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INTRODUCTION |
Sulfated glycosaminoglycans such as heparin and the related
heparan sulfate (HSGAGs)1 are
complex, linear carbohydrates possessing considerable chemical heterogeneity (2, 3). Their structural diversity is largely a
consequence of the variable number and position of sulfates present
within a single HSGAG chain. Because of their highly anionic character,
these polysaccharides historically have been relegated to an
exclusively structural role, namely as a sort of hydration gel and
scaffold comprising the extracellular matrix. Contrary to this limited
perception, however, HSGAGs actually play an important and dynamic
function in many critical biological processes ranging from development
(4) and tissue repair (5) to apoptosis (6, 7). These polysaccharides
are also central players in several pathological conditions such as
cancer (8, 9), angiogenesis (10, 11), certain neurodegenerative
diseases such as Alzheimer's (12), athleroscelerosis (13), and
microbial infectivity (14). HSGAGs do so as part of proteoglycans
found at the cell surface and within the extracellular matrix, where
they mediate signaling pathways and cell-cell communication by
modulating the bioavailability and temporal-spatial distribution of
growth factors, cytokines, and morphogens (15) in addition to various
receptors and extracellular adhesion molecules (16). We must now
therefore appreciate the fact that HSGAG structure and function are
inextricably related.
A study of the HSGAG structure-function paradigm (17) requires the
ability to determine both the overall composition of biologically
relevant HSGAGs as well as ultimately ascertaining their actual linear
sequence (fine structure). Invaluable toward this realization has been
the availability of several chemical and enzymatic reagents that are
able to cleave HSGAGs in a structure-specific fashion. One example
of an important class of GAG-degrading enzyme is the heparin lyases
(heparinases) originally isolated from the Gram-negative soil bacterium
Flavobacterium
heparinum.2 Each of the
three heparinases encoded by this microorganism cleaves both heparin
and heparan sulfate with a substrate specificity that is generally
based on the differential sulfation pattern that exists within each GAG
chain (18, 19). In fact, F. heparinum uses several
additional enzymes in an apparently sequential manner to first
depolymerize and then subsequently desulfate heparin/heparan sulfate.
The molecular cloning of each of these enzymes would be an important
first step toward their systematic use in vitro. In addition
to heparinase I (20), we have recently cloned one of these enzymes, the
4,5-unsaturated glycuronidase (1). This enzyme has also been
recombinantly expressed in Escherichia coli as a highly
active enzyme. Because of its rather unusual substrate specificity,
this enzyme has already proven to be a useful addition to our
property-encoded nomenclature-MALDI-based carbohydrate sequencing
methodology (21). The availability of specific HSGAG sulfatases would
make our enzyme repertoire complete.
In this paper, we likewise describe the molecular cloning of the
2-O-sulfatase from F. heparinum and its
recombinant expression in E. coli as a soluble, highly
active enzyme. This enzyme is a member of a highly conserved sulfatase
family as defined by a signature sulfatase domain located toward its
amino terminus. We also present a preliminary description of its
biochemical properties in addition to a kinetic description of the
enzyme's substrate specificity. Finally, we demonstrate the utility of
the sulfatase as a tool for probing HSGAG composition, especially when
the enzyme is used in tandem with the 4,5-glycuronidase. In the
accompanying paper (40), we describe the structure-function
relationship of this enzyme in greater detail, including the mapping of
a critically modified active cysteine to a formylglycine as well as the
construction of a structure-based homology model of the enzyme active site.
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EXPERIMENTAL PROCEDURES |
Reagents--
Heparin and chondroitin disaccaharides were
purchased from Calbiochem. Unfractionated heparin was obtained from
Celsus Laboratories (Cincinnati, OH). Materials for 
ZAP II genomic
library construction, screening, and phagemid excision, including
bacteriophage host strain XL1Blue MRF and the helper-resistant strain
SOLR, were obtained from Sratagene (La Jolla, CA) and used according to
the manufacturer's instructions. Restriction endonucleases and PCR enzymes were purchased from New England Biolabs (Beverly, MA). DNA
oligonucleotide primers were synthesized by an Invitrogen custom primer
service (Carlsbad, CA). TOP10 chemically competent cells for PCR
cloning and subcloning were also obtained from Invitrogen. [32P]dCTP radionuclides were purchased from PerkinElmer
Life Sciences. Additional molecular cloning reagents were
obtained from the manufacturers listed. All other reagents were from
Sigma unless otherwise noted.
Purification of the Flavobacterium Heparinum 2-O-Sulfatase and
Subsequent Proteolysis--
The 2-O-sulfatase was purified
from 20-liter fermentation cultures essentially as described (22).
Briefly, the large scale cultures were grown at 25 °C for 48 h.
Cell lysates were obtained by a repeated passage of a resuspended cell
pellet through an Aminco French pressure cell. The homogenate was
clarified by centrifugation (37,000 × g). The
2-O-sulfatase was purified from this cell-free supernatant
by employing five chromatographic steps carried out in the following
sequence: cation-exchange (CM-Sepaharose CL-6B) 
hydroxyapatite
(Bio-Gel HTP) gel filtration (Sephadex G-50) taurine-Sepharose
CL-4B blue Sepharose CL-6B. 2-O-Sulfatase activity was
measured at each chromatography step as described (22). Fractions from
initial CM-Sepharose chromatography were also assayed for heparinase,
chondroitinase (AC and B), and 4,5-glycuronidase activities as well
as any co-eluting 6-O- or N-sulfatase activities. The highly purified 2-O-sulfatase pool from the final blue
Sepharose chromatography step was free from any contaminating
glycosaminoglycan-degrading activity.
Generation of 2-O-Sulfatase Peptides and Protein
Sequencing--
In preparation for proteolysis, the purified
flavobacterial sulfatase was first desalted by reverse phase
chromatography (RP-HPLC) on a 150 × 4.6-mm C4 column (Phenomenex,
Torrance, CA). Protein was eluted by applying a linear gradient from 0 to 80% acetonitrile in 0.1% trifluoroacetic acid. During this
elution, both a major and minor protein peak were detected by UV
absorbance at 210 and 277 nm (Fig. 1A). The two separate
fractions were lyophilized to dryness and resuspended in 50 µl of
denaturation buffer (8 M urea, 0.4 M ammonium
bicarbonate, pH 7.5). Both protein fractions were digested with
modified trypsin for ~18 h at 37 °C. Trypsin was added at a 1:40
ratio (w/w) relative to each sulfatase fraction. Prior to proteolysis,
cysteines were first subjected to reductive carboxymethylation by the
addition of 5 mM dithiothreitol for 1 h at 50 °C,
followed by the addition of 20 mM iodoacetic acid for 30 min (room temperature). The alkylation reaction was quenched by the
addition of 50 µl of denaturation buffer. The resulting peptides were
resolved by RP-HPLC on a 250 × 2-mm C4 column using a linear
gradient of 2-80% acetonitrile in 0.1% trifluoroacetic acid carried
out over a 120-min time course. Select peptides corresponding to
chromatography peaks 2, 3, 4, 5, and 8 (Fig. 1B) were
sequenced using an on-line model 120 phenylthiohydantoin-derivative
analyzer (Biopolymers Laboratory, Massachusetts Institute of Technology).
Molecular Cloning of the Flavobacterial 2-O-Sulfatase--
The
2-O-sulfatase was cloned from a ZAP II flavobacterial
genomic library constructed and screened essentially as described for
the 4,5-glycuronidase (1). A 600-base pair DNA plaque hybridization
probe was generated by PCR using degenerate primers 5'-ATHGAYATHATHCCNACNATH-3' (forward) and 5'-DATNGTYTCATTNCCRTGYTG-3' (reverse). PCR was carried out for 35 cycles using a 52 °C annealing temperature and 2-min extensions at 72 °C. The specificity of this
probe was established by DNA sequence analysis, which indicated a
direct correspondence of its translated sequence to peak 1 tryptic peptides. Based on this information, the nondegenerate primers 5'-CATACACGTATGGGCGATTAT-3' (forward) and 5'-GATGTGGGGATGATGTCGAT-3' (reverse) were subsequently used in place of the original degenerate primers. PCR-amplified DNA probe was gel-purified and subsequently 32P-radiolabeled using the Prime-it II random priming kit
(Stratagene). Plaques were lifted on to nylon membranes (Nytran
Supercharge; Schleicher and Schuell), and DNA was cross-linked to each
filter by UV irradiation. Plaque hybridizations were completed
overnight at 42 °C according to standard methods and solutions (23).
Positive clones were visualized by phosphorimaging (Amersham
Biosciences) and/or 32P autoradiography. Clones were
further purified by secondary and tertiary screens, and the recombinant
phage was excised as a double-stranded phagemid (pBluescript) as
described by the manufacturer (Stratagene). Recombinants were confirmed
by DNA sequencing using both T7 and T3 primers. Insert size was
determined by restriction mapping of pBluescript inserts using
NotI, XbaI, and XhoI.
The full-length sulfatase gene (phagemid clone S4A) was subcloned into
the T7-based expression plasmid pET28a in three steps. In the first PCR
step, NdeI and XhoI restriction sites were
introduced at the 5' and 3' termini of the 2-O-sulfatase
coding sequence by using primers
5'-TGTTCTAGACATATGAAGATGTACAAATCGAAAGG-3' and 5'-GTCTCGAGGAT CCTTATTTTTTTAATGCATAAAACGAATCC-3',
respectively. At the same time, the NdeI restriction site
already present within the sulfatase gene starting at position 1050 (Fig. 2) was abolished by silent mutagenesis (CATATG CATCTG) using
the mutagenic primers 5'-GATATTATCCCCACCATCTGTGGCTTTGCCGGAA-3' and
5'-TTCCGGCAAAGCCACAGATGGTGGGGATAATATC-3', with the A
to C transversion underlined. In the second step, the final PCR product
was gel-purified and ligated into the TOPO/TA PCR cloning vector pCR
2.1 (Invitrogen) following the addition of 3'-dA overhangs with 0.5 units of Taq polymerase and 300 µM dATP (10 min, 72 °C). Ligated DNA was transformed into One-shot TOP10
chemically competent cells. Positive clones were identified by
blue/white colony selection and confirmed by PCR colony screening. In
the third step, the 1.5-kb sulfatase gene was excised from pCR 2.1 TOPO
and pasted into pET28a (Novagen) as an NdeI-XhoI cassette. Final expression clones were confirmed by plasmid DNA sequencing.
A 2-O-sulfatase amino-terminal truncation lacking the first
24 amino acids (2-O 1-24) was PCR-cloned as
above, except the forward primer
5'-TCTAGACATATGCAAACCTCAAAAGTAGCAGCT-3' was used in place
of the original outside 5' primer listed. In this DNA construct, the
2-O-sulfatase-specific sequence begins with
Gln25 (Fig. 2) and reads MQTSKVAASRPN.
Recombinant Expression and Protein Purification of a 6×
Histidine-tagged 2-O-Sulfatase (and 2-O
N1-24)--
Both the full-length enzyme and the
truncated enzyme (2-O N1-24) were
recombinantly expressed in the E. coli strain BL21 (DE3) (Novagen, Madison, WI) initially as NH2-terminal 6×
histidine fusion proteins to facilitate purification. The protocol for
their expression and subsequent one-step purification by nickel
chelation chromatography was as previously described for the
4,5-glycuronidase (1). Greater than 90% of the enzyme was eluted
from a 5-ml column in a single 12.5-ml fraction following the addition
of high imidazole elution buffer (50 mM Tris-HCl, pH 7.9, 0.5 M NaCl, and 250 mM imidazole). The enzyme
was immediately diluted with 2 volumes of cold enzyme dilution buffer
(50 mM Tris, pH 7.5, 100 mM NaCl). Cleavage of
the 6× histidine tag by thrombin was achieved by the stepwise addition
of 10 units of biotinylated thrombin (total of 50 units) to 30 ml of
diluted enzyme over the course of several hours while gently mixing by
inversion at 4 °C. Substantial precipitation of the sulfatase
routinely occurred during the cleavage reaction. Thrombin was recovered
by the addition of streptavidin-agarose using the thrombin cleavage
capture kit (Novagen). Capture was carried out at 4 °C for 2 h
with gentle mixing. Bound thrombin was collected by centrifugation for
5 min at 500 × g. Supernatant containing soluble
2-O-sulfatase was then dialyzed at 4 °C against 12 liters
of enzyme dilution buffer using 20.4-mm diameter Spectra/Por dialysis
tubing (Spectrum Laboratories, Rancho Dominguez, CA) with a 10,000 molecular weight cut-off. Following dialysis, the purified sulfatase
was concentrated using a Centriplus YM10 ultrafiltration device
(Millipore Corp.). Enzyme was stable for at least 2 weeks at 4 °C.
Long term storage was carried out at 
85 °C in the presence of 10%
glycerol without any substantial loss of activity due to freezing and thawing.
Protein concentrations were determined by the Bio-Rad protein assay and
confirmed by UV spectroscopy using a theoretical molar extinction
coefficient (
280) of 77,380 M
1
for 2-O N1-24 with the histidine tag
removed. Protein purity was assessed by silver staining of
SDS-polyacrylamide gels.
Molecular Mass Determinations by MALDI Mass
Spectrometry--
The molecular weight of the 2-O-sulfatase
NH2-truncated enzyme (2-O
N1-24) was determined by MALDI mass spectrometry
essentially as described (19). The NH2-terminal histidine
tag of the recombinant protein was cleaved by thrombin prior to mass
analysis. 1 µl of a 2-O-sulfatase solution (diluted in
water to 0.5 mg/ml) was added to 1 µl of a saturated sinapinic acid
matrix solution previously deposited on to the plate. The observed mass
of the recombinant enzyme was corrected according to an external
calibration using mass standards.
2-O-Sulfatase Assay and Determination of Biochemical Reaction
Conditions--
2-O-sulfatase activity was measured using
the unsaturated heparin trisulfated disaccharide
U2SHNS,6S3
or the disulfated disaccharide U2SHNS as
well as the disulfated disaccharide UHNS,6S lacking a
sulfate at the 2-OH-position. Standard reactions included 50 mM imidazole, pH 6.5, 50 mM NaCl, 500 µM disaccharide, and 25 nM enzyme (2-O
N1-24) in a 20-µl reaction volume. The reaction was
carried out for 30 s at 30 °C. Prior to its addition, the
enzyme was serially diluted to 250 nM in ice-cold 1×
imidazole buffer. The assay was initiated by the addition of 2 µl
of this 10× enzyme stock to 18 µl of reaction mixture. The enzyme
was inactivated by heating at 95 °C for 5 min in preheated 0.5-ml
Eppendorf tubes. Desulfation at the 2-OH-position of the disaccharide
was measured by capillary electrophoresis. Resolution of substrate and
product was achieved under standard conditions described for HSGAG
compositional analyses (19). Activity was generally measured as mol of
desulfated product formed and was calculated from the measured area of
the product peak based on molar conversion factors empirically
determined from standard curves. For the detection of mono- and
disulfated disaccharide products, total electrophoresis time was 20 min. Each unsaturated disaccharide peak was detected by UV absorption at 232 nm.
For pilot experiments measuring the relative effect of ionic strength
on 2-O-sulfatase activity, the NaCl concentration was varied
from 0.05 to 1 M in 50 mM MES buffer (pH 6.5)
that included 500 µM of the disulfated disaccharide
U2SHNS and 50 nM enzyme. The
effect of pH on sulfatase activity was assessed as a function of
catalytic efficiency by measuring kinetic parameters in the following
two overlapping pH buffer systems ranging from 5.0 to 8.0: 50 mM MES at pH 5.0, 5.5, 6.5, and 7.0; 50 mM MOPS
at pH 6.5, 7.0, 7.5, and 8.0. Assays included 25 nM enzyme,
50 mM NaCl, and varying concentrations of the disulfated
disaccharide substrate U2SHNS.
Km and kcat values were
extrapolated from Vo versus [S] curves
fit to the Michaelis-Menten equation by a nonlinear least squares
regression, and the relative
kcat/Km ratios were plotted
as a function of buffer pH. Based on this profile, relative enzyme
activity was also measured in four different buffers (MES, imidazole,
ADA, and sodium phosphate), each present as a 50 mM
concentration at pH 6.5. Relative activities were measured at a single
saturating substrate concentration (4 mM) using
U2SHNS.
Substrate Specificity and Kinetics Experiments Using Different
Disaccharide Substrates--
For substrate specificity experiments,
the following heparin disaccharide substrates were used:
U2SHNAc,
U2SHNAc,6S,
U2SHNS, and
U2SHNS,6S. In addition, the chondroitin
disaccharides U2SGalNAc,4S and
U2SGalNAc,6S were also studied. Disaccharide
concentrations for each respective substrate were varied from 0.1 to 4 mM. Initial rates (Vo) were extrapolated
from linear activities representing <20% substrate turnover and fit
to pseudo-first-order kinetics.
Tandem Use of 2-O-Sulfatase and 4,5-Glycuronidase in HSGAG
Compositional Analyses--
200 µg of heparin was first digested
with all three heparinases in an overnight digestion in glycuronidase
reaction buffer, which included 50 mM PIPES, pH 6.5, 50 mM NaCl, and a 100-µl reaction volume. The heparinase
digestion mix was split into four 20-µl reactions, which were
individually treated as follows: Tube 1, no addition (heparinase only
control); Tube 2, 5 µg of 4,5-glycuronidase, 30 °C for 1 h; Tube 3, 5 µg of 2-O-sulfatase (2-O
N1-24), 37 °C for 1 h; Tube 4, 2-O-sulfatase and 4,5-glycuronidase added simultaneously,
30 °C for 1 h. 4,5-Glycuronidase activity was ascertained by
a disappearance of unsaturated disaccharide peaks due to the loss of UV
absorption at 232 nm.
The substrate-product relationship between the two enzymes was examined
by directly measuring 4,5-glycuronidase activity either before or
following the addition of recombinant 2-O-sulfatase. Reactions were carried out at 30 °C and included 50 mM
MES, pH 6.5, 100 mM NaCl, and 2 mM
U2SHNS in a 100-µl reaction volume. In
these experiments, 250 nM 4,5-glycuronidase and 25 nM 2-O N1-24 were sequentially
added as follows: 4,5 alone, 4,5 followed by
2-O-sulfatase, or 2-O-sulfatase followed by
4,5. In each case, the first enzyme was added to the reaction in a
2-min preincubation step. 4,5-glycuronidase activity was measured
immediately following the addition of the second enzyme by determining
the rate of substrate disappearance as monitored by the loss of UV
absorption at 232 nm (1). 4,5 activity for the corresponding
2-O-desulfated disaccharide UHNS was also
measured under identical conditions.
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RESULTS |
Molecular Cloning and Recombinant Expression of the F. heparinum 2-O-Sulfatase--
As a first step toward the
cloning the 2-O-sulfatase gene, we purified the enzyme
directly from the native bacterium followed by a partial determination
of its amino acid sequence. Enzyme purification was essentially as
previously described (22). After a five-step chromatographic
fractionation of flavobacterial lysates, we achieved a greater than
3000-fold purification of sulfatase activity. Further fractionation of
this activity by reverse phase HPLC chromatography yielded two separate
polypeptides (Fig. 1A). Both
proteins were subjected to trypsin digestion, and the resultant peptides were likewise purified by reverse phase HPLC (Fig.
1B). From select peak 1 peptide sequences, degenerate
primers were synthesized. We initially screened primer pairs
corresponding exclusively to the peak 1 protein sequence (Table
I), given the fact that this sulfatase
fraction represented the major protein species present in the final
purification step. PCR amplification of genomic DNA using degenerate
primers corresponding to peptide peaks 3 and 5 yielded a discrete
600-bp DNA product. Sequence analysis of this amplified DNA indicated a
translated amino acid sequence to which three of the isolated peak 1 peptides mapped. We used this DNA, therefore, as a hybridization probe
to screen a ZAP flavobacterial genomic library and isolate a
full-length clone. Several positive clones were isolated; most of them
contained an average insert size between 4 and 5 kb. One genomic clone
of ~7 kb (clone S4A) was subjected to direct DNA sequencing. This clone contained at least one open reading frame in particular that
encodes a putative protein of 468 amino acids in length (464 amino
acids from the first methionine) and whose primary sequence includes
all of the sulfatase peptides for which we had obtained sequence
information (Fig. 2). Based on its amino
acid composition, the encoded protein is quite basic (theoretical pI of
8.75), with 67 basic side chains comprising ~14% on a molar basis.
The putative sulfatase also possesses 8 cysteines in addition to 46 aromatic amino acids.
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Fig. 1.
Flavobacterium 2-O-sulfatase
purification and proteolysis. A, final RP-HPLC
chromatography of blue Sepharose CL-6B-purified sulfatase.
B, C-4 RP-HPLC chromatographic resolution of sulfatase
peptides generated by a trypsin digestion of the major peak shown in
A.
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Table I
2-O-sulfatase peptides and corresponding degenerate primers
Select RP-HPLC-purified tryptic peptides (Fig. 1B) were
subjected to amino acid sequencing. Also shown are the corresponding
degenerate primer pairs for these peptides.
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Fig. 2.
F. heparinum
2-O-sulfatase coding sequence. Open reading
frame from genomic clone S4A. Translation initiation and termination
codons are shown in boldface type. Primers used
in the original PCR screen are noted by horizontal
arrows. An internal NdeI site is double
underscored. The amino acid sequence corresponding to select sulfatase
peptides are boxed. Sulfatase consensus sequence
CXPXRXXXX(S/T)G is boxed
and shaded with active site cysteine at position 82 noted by
an asterisk. Putative signal sequence is
overscored, with the predicted peptidase cleavage site
represented by a vertical arrow.
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Upon a closer examination of its primary sequence, we also identified a
conserved sulfatase domain as annotated in the Protein Family
Database.4 This signature
domain included the consensus sequence
(C/S)XPXRXXXX(S/T)G (25),
comprising, at least in part, the sulfatase active site and possessing
the cysteine (underlined) that is presumably modified as a
formylglycine in vivo (26). The putative
2-O-sulfatase that we cloned from F. heparinum
exhibits substantial homology to many members of a highly conserved
sulfatase family (data not shown) (27, 28). A structurally oriented
description of this homology and its correlation to enzyme function is
found in the accompanying paper (40).
From this sequence information, we were confident that we had indeed
cloned a sulfatase from the flavobacterial genome. To ultimately establish its functionality, we next set out to
recombinantly express this protein in E. coli. The
full-length gene (beginning at the first methionine noted in Fig. 2)
was subcloned into the T7-based expression vector, pET28a, for
expression as an NH2-terminal 6× histidine-tagged protein
to facilitate purification. Induction with
isopropyl-1-thio-
-D-galactopyranoside led to a limited
soluble expression of a polypeptide whose apparent molecular weight
roughly corresponded to the theoretical mass of the fusion protein
(~54 kDa). Using Ni2+ chelation chromatography, we were
able to partially purify this polypeptide from the bacterial lysate and
unequivocally measure 2-O-specific sulfatase activity using
the trisulfated, unsaturated heparin disaccharide
U2SHNS,6S as a substrate.
Both the total and soluble protein expression levels achieved were
unsatisfactory, however, especially given our previous successes with
recombinantly expressing other HSGAG-degrading enzymes cloned from
F. heparinum (1, 29, 30). As has been the case for most of
these enzymes, removal of their putative N-terminal signal sequences
greatly facilitated the recombinant expression of soluble protein
without compromising their respective specific activities. We
likewise identified a putative signal sequence for the flavobacterial
2-O-sulfatase comprised of the first 24 amino acids (see
Fig. 2). By engineering a 2-O-sulfatase N-terminal
truncation lacking this sequence (herein referred to as 2-O
1-24), we achieved high expression levels of soluble,
highly active enzyme. Protein yields exceeding 100 mg of relatively
pure sulfatase per liter of induced bacterial cultures were routinely
achieved using a single chromatographic step (Fig.
3). The specific activity of the
recombinant sulfatase was considerably enhanced following the removal
of the N-terminal 6× histidine tag by thrombin cleavage. Removal of
this purification tag resulted in a greater than 10-fold purification
of sulfatase activity relative to the crude bacterial lysate (Table
II). For this reason, we used the cleaved
protein in all subsequent experiments. The molecular mass of
this recombinantly expressed sulfatase as determined by MALDI mass
spectrometry is 50,120.8 daltons. This empirical value closely agrees
with its theoretical mass of 49,796 daltons, which is based entirely on its amino acid composition.
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Fig. 3.
Purification of recombinant
2-O-sulfatase from E. coli lysates by
Ni2+ chelation chromatography. Enzyme purity following
each fractionation step was assessed by silver staining of 12%
SDS-polyacrylamide gels. Approximately 200 ng of total protein was
loaded in each well. Lane 1, bacterial lysate from uninduced
(minus isopropyl-1-thio--D-galactopyranoside) control;
lane 2, whole cell lysate; lane
3, 20,000 × g supernatant (column preload);
lane 4, eluate from Ni2+ chelation
chromatography; lane 5, 2-O-sulfatase
following thrombin cleavage to remove the NH2 6× histidine
purification tag. Molecular weight markers (Mr)
and their corresponding masses are also shown.
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Table II
Purification of recombinant 2-O-sulfatase
200 ng of total protein from each purification step was assayed for
2-O-sulfatase activity as described under "Experimental
Procedures" using the unsaturated heparin disaccharide (DiS)
U2SHNS as a substrate. -Fold purification is
relative to crude bacterial lysate.
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To establish the recombinant enzyme's exclusivity for the uronic acid
2-O-sulfate, we initially compared two related unsaturated heparin disaccharides: U2SHNS,6S
versus UHNS,6S. The recombinant sulfatase
only hydrolyzed a single sulfate, namely the one found at the
2-OH-position (Fig. 5A). Other sulfated positions were not
hydrolyzed (data not shown).
Biochemical Conditions for Optimal in Vitro Activity--
Having
successfully achieved the recombinant expression and purification of
the flavobacterial sulfatase as a soluble enzyme as well as
demonstration of its unequivocal specificity for the uronic acid
2-O-sulfate, we next set out to define the reaction conditions required for optimal enzyme activity in vitro.
These parameters included pH, temperature, ionic strength, and possible divalent metal ion dependence. In brief, the enzyme exhibited a pH
activity range between 6.0 and 7.0, with optimum activity occurring at
pH 6.5 (Fig. 4A). The enzyme
was essentially inactive at the outlying pH values of 5.0 and 8.0. In
terms of different buffer systems (all at pH 6.5), an imidazole-based
buffer demonstrated the highest relative activity as compared with
buffers containing 50 mM MES, ADA, or phosphate. As
expected, phosphate buffer was clearly inhibitory (see
histogram, Fig. 4A, inset).
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Fig. 4.
In vitro biochemical reaction
conditions for the recombinant 2-O-sulfatase.
A, effect of pH. Sulfatase catalytic efficiency
(kcat/Km) was measured as a
function of varying pH from 5 to 8 using two overlapping buffers: 50 mM MES (solid circles) and 50 mM MOPS (open circles).
Inset, relative effect of three different assay buffers
(each at pH 6.5) on optimal enzyme activity. 1, 50 mM MES; 2, 50 mM imidazole;
3, 50 mM sodium phosphate. B, effect
of ionic strength. Shown here is percentage activity normalized
to 50 mM NaCl. C, effect of reaction
temperature. Data is normalized to 30 °C activity (100%). The
unsaturated disaccharide U2SHNS was used in
all three experiments.
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We also examined 2-O-sulfatase activity relative to ionic
composition. The recombinant enzyme was optimally active at ~50 mM NaCl. Activity was sharply inhibited by [NaCl]
exceeding 100 mM, with 50% inhibition occurring at less
than 250 mM NaCl (Fig. 4B). Maximal enzyme
activity was largely unaffected by the addition of EDTA up to a 1 mM concentration. The addition of exogenous CaCl2, MgCl2, or MnCl2 (up to 10 mM) also had no substantive effect, indicating that these
particular divalent metal ions are not required (data not shown). A
preincubation of the enzyme with 5 mM EDTA did result in an
~10% inhibition of activity using the trisulfated disaccharide as a substrate.
37 °C was the default temperature at which all of the preliminary
biochemical experiments were conducted. We measured both relative
enzyme activity and stability as a function of varying reaction
temperature (Fig. 4C). The 2-O-sulfatase was
active over a fairly broad temperature range (25-37 °C), with
optimal activity occurring at 30 °C. Enzyme activity was compromised
at 42 °C. Enzyme stability at this temperature was likewise affected
at this temperature as assessed in preincubation experiments conducted at varying temperatures (30-42 °C) prior to measuring
2-O-sulfatase activity at 30 °C (data not shown).
Determination of Disaccharide Substrate Kinetics and
Specificity--
We were interested in ascertaining any kinetic
discrimination the enzyme may possess for its disaccharide substrates
based on the following structural considerations: 1) the number and position of sulfates on the adjoining hexosamine; 2) the glycosidic linkage position (i.e. 1 4 versus
1 3); and 3) glucosamine versus galactosamine as
the adjoining hexosamine. We examined substrate saturation kinetics
measured under Michaelis-Menten conditions. For these experiments,
several heparin disaccharide substrates were used, each with a uronic
acid possessing a 2-O-sulfate and a 4,5-unsaturated bond
at the nonreducing end but differing in the degree of sulfation
within the glucosamine. In addition, the two unsaturated
chondroitin disaccharides U2SGalNAc,4S and U2SGalNAc,6S were also examined as possible
substrates. These latter two disaccharides differ from those derived
from heparin/heparan sulfate in possessing a 1 3 glycosidic
linkage and a galactosamine in place of a glucosamine. The results are
summarized in Fig. 5B and
Table III. All of the heparin
disaccharides examined were hydrolyzed at substantial rates that
included kcat values that varied from ~600 to
1700 s1. At the same time, the 2-O-sulfatase
did exhibit a kinetic preference directly related to the extent of
sulfation; the trisulfated disaccharide (U2SHNS6S) was clearly the preferred
substrate, whereas the monosulfated disaccharide
(U2SHNAc) was the least preferred substrate.
The disulfated disaccharides U2SHNS and
U2SHNAc,6S had
kcat/Km values intermediate
between these two extremes.
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Fig. 5.
2-O-sulfatase substrate
specificity. A, exclusive desulfation of the 2-OH-position
by the recombinant sulfatase. Enzyme desulfating activity was assayed
by capillary electrophoresis using the 2-O- containing
trisulfated heparin disaccharide U2SHNS,6S
or its disulfated counterpart UHNS,6S lacking a sulfate
at the 2-OH-position (data not shown). Only for the substrate
U2SHNS,6S is a loss of sulfate observed.
Minus enzyme control is shown as a dotted line.
B, steady-state kinetics for various unsaturated
disaccharide substrates. Initial rates were determined using 25 nM enzyme under standard conditions. Substrate saturation
data were fit to pseudo-first order Michaelis-Menten assumptions using
a nonlinear least squares analysis.  ,
U2SHNAc;  ,
U2SHNAc,6S;  ,
U2SHNS;  ,
U2SHNS,6S; +,
U2SGalNAc,6S.
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Table III
2-O-sulfatase disaccharide substrate specificity
Kinetic parameters were derived from a nonlinear regressional analyses
of substrate saturation data depicted in Fig. 5B.
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The 2-O-sulfated chondroitin disaccharide
U2SGalNAc6S was only negligibly hydrolyzed
under the same kinetic conditions. The enzyme did desulfate this
disaccharide to an appreciable extent, however, under reaction
conditions involving a 4 times higher enzyme concentration and a longer
incubation time (data not shown). Under these conditions, ~40% of
the substrate was desulfated over a 20-min period. In contrast, less
than 10% of chondroitin disaccharide U2SGalNAc4S was hydrolyzed during the same
time period. To determine whether either or both of these
2-O-sulfated chondroitin disaccharides could be
quantitatively desulfated under exhaustive conditions, we carried out
an 18-h incubation at 30 °C that included 5 mM substrate
and 5 µM enzyme. Under these conditions, both chondroitin disaccharides were greater than 95% desulfated at the 2-O-position (data not shown). This result indicates that whereas linkage position and/or hexosamine isomerization are discriminating kinetic factors, these physical parameters are not absolute determinants for
2-O-sulfatase substrate recognition.
The Substrate-Product Relationship between the 2-O-Sulfatase and
4,5-Glycuronidase--
As we have already noted, the flavobacterial
4,5-glycuronidase is unable to hydrolyze unsaturated saccharides
possessing a uronic acid 2-O-sulfate at the nonreducing end
(1). Considering this fact, an obligatory substrate-product
relationship between the 2-O-sulfatase and the
4,5-glycuronidase must apparently exist. We examined a possible
kinetic relationship between these two enzymes by looking at their
sequential action (Fig. 6). In this experiment, 4,5-glycuronidase activity was measured directly either
during or following the addition of the recombinant
2-O-sulfatase using the disaccharide substrate
U2SHNS. As expected, when this disaccharide
was incubated with the 4,5 enzyme alone, it was completely
refractory to glycuronidase-mediated hydrolysis as measured by a loss
of absorbance at 232 nm. A 2-min preincubation of the substrate with
the 2-O-sulfatase, however, resulted in robust linear
glycuronidase activity. This rate was comparable with the rate of
hydrolysis measured for the control substrate UHNS using
the 4,5 enzyme alone (data not shown). In the reciprocal experiment
(i.e. in which the 2-O-sulfatase was added
second), we observed an initial lag in 4,5 activity. This lag was
followed by a linear 4,5 activity, albeit at a slower rate than in
the case where the 2-O-sulfatase was added first. The
observed delay in activity was presumably due to the prerequisite
2-O-desulfation of the substrate, which must occur prior to
being acted on by the glycuronidase. This experiment clearly
demonstrates at least a functional linkage between these two
HSGAG-degrading enzymes. We were unable to show a kinetic coupling of
these two enzymes, however, in a related experiment where
2-O-sulfatase activity was measured directly using the same
disulfated disaccharide (U2SHNS) substrate
in the absence versus the presence of the
4,5-glycuronidase (data not shown).
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Fig. 6.
Substrate-product relationship between the
2-O-sulfatase and the
4,5-glycuronidase. A 2 mM
concentration of the unsaturated, 2-O-sulfated heparin
disaccharide U2SHNS was preincubated with
either 250 nM 4,5-glycuronidase or 25 nM
2-O-N1-24 for 2 min at 30 °C in a
100-µl reaction. Following this preincubation, the reciprocal enzyme
was added to the reaction for up to an extra 6 min.
4,5-glycuronidase activity was measured in real time as the rate of
substrate disappearance monitored by the loss of UV absorption at 232 nm. Zero time on the x axis represents the time following
the preincubation, during which the second enzyme was added.
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With the results just described, we considered the parallel use of
these two enzymes (along with the heparinases) as complementary tools
for HSGAG compositional analyses. The utility of this combinatorial approach is shown in Fig. 7. 200 µg of
heparin were first subjected to an exhaustive heparinase treatment.
Subsequent treatment of the cleavage products with the
4,5-glycuronidase resulted in the disappearance of select saccharide
peaks, namely those that did not possess a 2-O-sulfated
uronic acid at the nonreducing end (Fig. 7B). Conversely,
subsequent treatment of the heparinase-derived saccharides with the
2-O-sulfatase results in both the disappearance of
2-O-sulfated disaccharides and a concomitant appearance of their desulfated products (Fig. 7C). When both the
4,5-glycuronidase and the 2-O-sulfatase were added
simultaneously to the heparinase cleavage products, essentially all of
the saccharides were hydrolyzed by the 4,5-glycuronidase, as evident
by a lack of any UV-absorbable electrophoresis products (Fig.
7D).
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Fig. 7.
Tandem use of 2-O-sulfatase
and 4,5-glycuronidase in HSGAG compositional
analyses. A, 200 µg of heparin was first exhaustively
cleaved by heparinases I, II, and III. These heparinase-generated
saccharides were then subject to hydrolysis by the 4,5-glycuronidase
(B) or the 2-O-sulfatase (C) or the
2-O-sulfatase and 4,5-glycuronidase added simultaneously
(D). The seven disaccharide peaks (and one tetrasaccharide
peak) resolved by capillary electrophoresis are each numbered
separately. Their compositional assignments are as follows:
U2SHNS,6S (1);
UHNAc,6SGHNS,3S,6S tetrasaccharide
(2); U2SHNS (3);
UHNS,6S (4);
U2SHNAc,6S (5);
UHNS (6); U2SHNAc
(7); and UHNAc,6S (8).
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DISCUSSION |
Heparin and related heparan sulfate are structurally complex
glycosaminoglycans. Their extensive chemical heterogeneity plays out in
a functional context, namely in dictating how these linear oligosaccharides mediate a diverse number of biological signaling pathways. One of the more formidable challenges currently facing the
glycobiology field is the design of effective analytical methods to
study this structure-function relationship at the molecular level.
Structure-specific enzymatic tools are indispensable toward meeting
this challenge. The utility of HSGAG-degrading enzymes as such has been
considered for quite a long time. The heparin lyases (heparinases) from
F. heparinum are an excellent example of how such enzymes
have been put to such a use. However, whereas the heparinases may be an
important prototype, additional HSGAG-degrading enzymes with more
defined substrate specificities are now required to refine present
methods for sequencing these complex carbohydrates. Toward this end, we
have recently cloned from the flavobacterial genome an unsaturated
HSGAG glycuronidase (1) with a rather unusual substrate
specificity (31). The 4,5-glycuronidase can be recombinantly
expressed in E. coli as a soluble, highly active enzyme. We
have shown how this readily purified enzyme can be used as an enzymatic
tool to refine our property-encoded nomenclature-MALDI-based method for
sequencing heparinase-derived HSGAG oligosaccharides (21). In this
paper, we describe the molecular cloning, recombinant expression, and
biochemical characterization of another highly useful flavobacterial
HSGAG-degrading enzyme, the 2-O-sulfatase.
The cloned, full-length gene encodes an open reading frame of 468 amino
acids (Fig. 2), with a predicted molecular mass of 51.9 kDa. This
theoretical molecular weight is ~10,000 less than the value
reported in the literature (22). Based on its amino acid composition,
the encoded protein is quite basic (theoretical pI of 8.75). A further
analysis of its primary amino acid sequence unequivocally places this
open reading frame as a member of a larger sulfatase family. Members of
this superfamily all possess a defining sulfatase domain that resides
within the amino terminus and includes the signature motif
(C/S)XPXRXXXX(S/T)G (25). Demarcating this consensus sequence is the invariant active site cysteine or serine
that is covalently modified as a formylglycine. From our sequence
alignment, we predict cysteine at position 82 to be, in fact, the
covalently modified residue. An empirical demonstration of this
active-site aldehyde at this position is made in the following paper
(40).
Whereas the cloned flavobacterial sulfatase exhibits the highest
sequence similarity to the bacterial arylsulfatases (especially the
arylsulfatase from Pseudomonas aeruginosa), we point out
that a limited homology of the 2-O-sulfatase does extend to
the mammalian glycosaminoglycan sulfatases functioning in the lysosomal
degradation pathway (see Fig. 1 in the accompanying paper (40)). As is
the case for the bacterial enzymes, this sequence homology is strongest in the NH2 terminus, where the putative sulfatase domain
resides. Among the human lysosomal enzymes, it is the galactosamine
(N-acetyl)-6-sulfate sulfatase (chondroitin
6-O-sulfatase) that exhibits the closest similarity with the
flavobacterial 2-O-sulfatase; the two enzymes possess
~26% identity when comparing their entire protein sequences. There
are also two functionally related lysosomal sulfatases, which
specifically hydrolyze the 2-OH-position of uronic acid. These enzymes
are the iduronate-2-sulfate sulfatase (32) and the glucuronic 2-sulfate
sulfatase (33). The iduronate-2-sulfate sulfatase and flavobacterial
2-O-sulfatase exhibit only a limited sequence homology (less
than 22% identity), however.
Like the 4,5-glycuronidase, we successfully expressed the
2-O-sulfatase in E. coli, from which
several milligrams of highly active, soluble enzyme were readily
purified. As was also the case for the glycuronidase, we found that the
yield of soluble recombinant enzyme was greatly improved by the
engineered removal of the hydrophobic N-terminal signal sequence
composed of the first 24 amino acids. This signal sequence was
predicted by the Von Heinje method, which also identified the likely
signal peptidase cleavage recognition sequence
AXAXA. With only one exception, we have
identified a putative signal sequence in all of the flavobacterial HSGAG-degrading enzymes examined to date. The general presence of this
tag, therefore, would suggest a common periplasmic locale within the
flavobacterium for glycosaminoglycan degradation.
Our initial assessment of 2-O-sulfatase activity was based
upon the use of a few select unsaturated heparin disaccharide
substrates. Desulfation was unequivocally specific for the 2-O-position
(Fig. 5A). Our examination of the biochemical conditions for
optimal enzymatic activity yielded several observations. First,
2-O-sulfatase activity exhibited a pH profile generally in
agreement with previously published results (22), albeit with a
narrower pH range (6.0-7.0) in which the enzyme was most active. The
enzyme exhibited maximal catalytic efficiency at pH 6.5 with
essentially no activity observed at the outlying pH values of 5 and 8 (Fig. 4A). A sharply defined pH optima of 6.5 clearly
implicates a catalytic role of one or more histidines. Of the 8 histidines present in the flavobacterial 2-O-sulfatase,
His-136 and His-191 are highly conserved among the sulfatases examined.
Catalytically important histidines have been observed within the active
site of several sulfatase crystal structures including human lysosomal
N-acetylgalactosamine-4-sulfatase (arylsulfatase B) (27) and
arylsulfatase A (25) as well as the arysulfatase from P. aeriginosa (34) to which the flavobacterial 2-O-sulfatase appears to be most closely related. We have in
fact used the structures of these related enzymes to construct a highly informative homology model for the flavobacterial
2-O-sulfatase. This model and its structure-function
implications are presented in the accompanying paper (40).
Second, the observed NaCl titration profile (Fig. 4B)
demonstrates a clearly inhibitory effect of ionic strength on sulfatase activity, even at relatively low NaCl concentrations. In other words,
whereas 50% inhibition occurred in the presence of ~200 mM NaCl, even 100 mM NaCl was slightly
inhibitory to 2-O-sulfatase activity. We have likewise noted
this rather sharp activity transition for both the 4,5-glycuronidase
and other recombinantly expressed F. heparinum GAG-degrading
enzymes. This correlation between activity and the ionic buffer
composition is not surprising, given the anionic character of the
saccharide substrates conferred by both the presence of sulfates and
the uronic acid carboxylates within each disaccharide unit. For the
2-O-sulfatase in particular, charge interactions between
basic side chains and the sulfate oxygen anion would be clearly
important for substrate orientation. Additional interactions with the
uronic acid C-5 carboxylate are also likely to occur. A masking of
these important charges by exogenous ions would logically interfere
with their catalytic function.
The flavobacterial 2-O-sulfatase also possesses 52 acidic
amino acids, at least three of which are highly conserved
(e.g. Asp-42, Asp-63, and Asp-295). Four acidic side chains
are also found in a consensus active site derived from known crystal
structures. In this snapshot, these four carboxylates appear to
coordinate a divalent metal ion (typically calcium). We could find no
empirical evidence, however, for the requirement of divalent metal ions for maximal sulfatase activity. The enzyme was largely unaffected by
either a preincubation with EDTA (up to 5 mM) or the
addition of Ca2+, Mg2+, or Mn2+. We
were somewhat surprised by this in vitro observation given the structural inference of a common metal binding site and a report in
the literature describing inactivation of the glucuronate-2-sulfatase with EDTA (35). Whereas this may be so, not all sulfatases described to
date have been shown in fact to be inactivated by EDTA. The issue of
metal ion chelation is discussed in greater detail in the paper that
follows (40).
As stated earlier, the recombinant flavobacterial
2-O-sulfatase acted on every unsaturated heparin
disaccharide examined, provided each saccharide possessed a
2-O-sulfated uronic acid at its nonreducing end. We were
particularly interested, however, in ascertaining any kinetic
discrimination the enzyme may possess for its substrates based on
additional structural considerations, namely the number and position of
sulfates on the adjoining hexosamine, glycosidic linkage position
(i.e. 14 versus 13) and the
2-OH-position of the anomeric carbon within this hexosamine
(i.e. glucosamine versus galactosamine). For the
2-O-sulfated, unsaturated heparin disaccharides examined,
the enzyme exhibited kcat and
Km values roughly in the range of 600-1700
s1 and 0.1-1 mM, respectively. Both of these
parameters generally agree with values described in earlier studies
using the sulfatase directly purified from flavobacterial lysates. In
directly comparing the four heparin disaccharide substrates tested,
however, we noted a clear kinetic discrimination exhibited by the
recombinant enzyme; one based on the extent of sulfation and largely
manifested as a Km effect. In particular, it appears
that substrate binding is most favorably conferred by the presence of a
sulfate at the 6-OH-position of the adjoining glucosamine. Sulfation of the amine likewise confers a positive effect, albeit to a somewhat lesser degree relative to the 6-OH-position.
Interestingly, the 2-O-sulfatase was also able to desulfate
unsaturated chondroitin disaccharides at the 2-OH-position. The recombinant enzyme is, therefore, both a heparin/heparan and
chondroitin 2-O-sulfatase. Compared with the corresponding
heparin substrate, the rate of hydrolysis was markedly slower, however
(Fig. 5B). This result indicates that whereas linkage
position and/or hexosamine isomerization are discriminating kinetic
factors, these physical parameters are not absolute determinants for
2-O-sulfatase substrate recognition. It is interesting to
consider this latter observation in the context of the lysosomal
pathway for glycosaminoglycan degradation in mammals where one enzyme
desulfates both chondroitin and HS oligosaccharides at this position.
In our studies, we exclusively used unsaturated saccharide substrates
on the presumption that they are the naturally occurring substrate
in vivo. If the 2-O-sulfatase ordinarily acts on
heparinase-derived saccharides in a sequential HSGAG degradation
pathway, we can also place the activity of the 2-O-sulfatase
"upstream" from the hydrolysis of the unsaturated uronic acid by
the 4,5-glycuronidase. The obligatory substrate-product relationship
between the 2-O-sulfatase and the 4,5-glycuronidase has
been described by our laboratory (1) and others (36). In this paper, we
further demonstrate this correlation in two experiments summarized in
Figs. 6 and 7. Fig. 7 also illustrates how these two enzymes (along
with the heparinases) can be used in tandem as analytical tools for
HSGAG compositional analyses.
As we pointed out earlier, there are 2-O-sulfatases present
in the mammalian glycosaminoglycan degradation pathway. Whereas both of
these enzymes desulfate heparan sulfate, the iduronate-2-sulfate sulfatase also acts on dermatan sulfate. Both enzymes possess an acidic
pH optimum for activity, a fact consistent with their location within
the lysosome. The two sulfatases initially exist as precursors that
must be proteolytically processed for activity. The native molecular
mass of the human iduronate-2-sulfate sulfatase precursor has
been reported in the range of 42-65 kDa (32), whereas its theoretical
mass based entirely on its amino acid composition is ~62 kDa. As
such, the mammalian lysosomal iduronate-2-sulfate sulfatase is somewhat
larger than its flavobacterial counterpart, while also requiring
substantial posttranslational modification for maximal enzyme activity.
The acidic pH optima for the lysosomal enzymes would also appear to
limit their in vitro use for the determination of HSGAG
composition, at least when used in tandem with other flavobacterial
HSGAG-degrading enzymes such as the heparinases or the
4,5-glycuronidase; these latter enzymes all possess a pH optimum
much closer to neutrality.
The obvious fact that we are able to measure robust,
2-O-specific sulfatase activity for the recombinantly
expressed enzyme argues, at least from a functional perspective, for
the presence of the critical catalytic formylglycine within the
sulfatase active site. This functionality clearly validates our use of
E. coli as a recombinant expression system for the large
scale production of active enzyme. Our results also logically point to
the existence of the necessary sulfatase-modifying enzyme(s) encoded by
the E. coli genome. The heterologous expression of other
catalytically active sulfatases in E. coli has been reported
(34, 37). One notable exception of which we are aware is the
mucin-desulfating enzyme from Prevotella sp. (38).
Interestingly, this particular desulfating enzyme possesses a modified
serine rather than a cysteine within its active site, a fact supported
by our own multiple sequence alignment (see Fig. 1 in the accompanying
paper (40)). To the best of our knowledge, nearly every active,
recombinantly expressed sulfatase reported in the literature possesses
a cysteine (and not a serine) within the active site sequence
(C/S)XPXRXXXX(S/T)G. It seems likely,
therefore, that a cysteine-specific modifying machinery functionally
exists in E. coli. Our initial attempts to produce a
recombinant cysteine serine 2-O-sulfatase variant led to
the production of insoluble protein when expressed in E. coli. Consequently, we are currently unable to empirically address this hypothesis using the 2-O-sulfatase as a prototype. We
point out the curious fact that the E. coli genome encodes
for at least three different putative sulfatase genes, in addition to
the atsB gene, which, by homology, has been proposed to
encode for this cysteine-specific modifying activity. All of these
genes are located as a cluster within the bacterial chromosome (39). It
would appear, however, that the E. coli sulfatase genes are
normally cryptic. At the very least, E. coli lacks the
specific enzymes for desulfating heparin/heparan sulfate
glycosaminoglycans. In any case, the bacterium fortuitously provides
the necessary enzymology to effectively modify select heterologous
sulfatases such as the 2-O-sulfatase. We look forward to the
use of this facile expression system to recombinantly express other
flavobacterial glycosaminoglycan sulfatases as their respective genes
are identified.
§,
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TOP
ABSTRACT
INTRODUCTION
