Tài liệu Báo cáo khoa học: Inorganic pyrophosphatase in the roundworm Ascaris and its role in the development and molting process of the larval stage parasites doc

Inorganic pyrophosphatase in the roundworm
Ascaris
and its role in the development and molting process
of the larval stage parasites
M. Khyrul Islam
1
, Takeharu Miyoshi
1
, Harue Kasuga-Aoki
1
, Takashi Isobe
1
, Takeshi Arakawa

2
,
Yasunobu Matsumoto
3
and Naotoshi Tsuji
1
1
Laboratory of Parasitic Diseases, National Institute of Animal Health, National Agricultural Research Organization, 3-1-5,
Kannondai, Tsukuba, Ibaraki, Japan;
2
Division of Molecular Microbiology, Center of Molecular Biosciences, University of the
Ryukyus, Senbaru, Okinawa, Japan;

3
Laboratory of Global Animal Resource Science, Graduate School of Agricultural and
Life Sciences, University of Tokyo, Yayoi, Bunkyo, Tokyo, Japan
Inorganic pyrophosphatase (PPase) is an important
enzyme that catalyzes the hydrolysis of inorganic pyro-
phosphate (PP
i
)intoortho-phosphate (P
i
). We report
here the molecular cloning and characterization of a gene
encoding the soluble PPase of the roundworm Ascaris

suum. The predicted A. suum PPase consists of 360 amino
acids with a molecular mass of 40.6 kDa and a pI of 7.1.
Amino acid sequence alignment and phylogenetic analysis
indicates that the gene encodes a functional Family I
soluble PPase containing features identical to those of
prokaryotic, plant and animal/fungal soluble PPases. The
Escherichia coli-expressed recombinant enzyme has a spe-
cific activity of 937 lmol P
i
Æmin
)1
Æmg

)1
protein corres-
ponding to a k
cat
value of 638 s
)1
at 55 °C. Its activity was
strongly dependent on Mg
2+
and was inhibited by Ca
2+
.

Native PPases were expressed in all developmental stages of
A. suum. A homolog was also detected in the most closely
related human and dog roundworms A. lumbricoides and
Toxocara canis, respectively. The enzyme was intensely
localized in the body wall, gut epithelium, ovary and uterus
of adult female worms. We observed that native PPase
activity together with development and molting in vitro of
A. suum L3 to L4 were efficiently inhibited in a dose-
dependent manner by imidodiphosphate and sodium
fluoride, which are potent inhibitor of both soluble- and
membrane-bound H
+

-PPases. The studies provide evi-
dence that the PPases are novel enzymes in the roundworm
Ascaris, and may have crucial role in the development and
molting process.
Keywords: roundworm; inorganic pyrophosphatase;
sodium fluoride; imidodiphosphate; molting.
Geohelminth parasites are among the commonest and
widespread of human infections, particularly in the regions
where public health hygiene and nutritional status are
poorly maintained. The most prevalent geohelminth is
Ascaris lumbricoides (originally described by Linnaeus in
1758), which colonizes the small intestine of children, and is

estimated to infect a quarter of the world’s population [1].
Ascaris suum (originally described by Goeze in 1782) of pigs
is a very closely related species to A. lumbricoides,whichcan
develop in human hosts, indicating its zoonotic significance
[2,3]. Childhood infections with Ascaris worms are reported
to be associated with stunting growth, malabsorption,
deficiencies of macro- and micro-nutrients and damage of
the small intestinal mucosa [4,5]. In addition, concurrent
Ascaris infection may have potential immunomodulatory
effects on the immune response to other infections [6,7]. It
is therefore of considerable interest to investigate the
biochemical aspects of Ascaris worms to identify potential

drug targets and vaccine candidates.
Inorganic pyrophosphatases (PPases), which catalyze the
hydrolysis of inorganic pyrophosphate (PP
i
) into inorganic
ortho-phosphate (P
i
), are widely distributed among living
cells. The enzymes play an important role in energy
metabolism, providing a thermodynamic pull for many
biosynthetic reactions [8], and have been shown to be
essential to life [9–11]. There are two major categories of

PPases, the soluble PPases and the membrane-bound H
+
-
translocating PPases (H
+
-PPase). Two families of soluble
PPases have been recognized to date, Family I includes most
of the currently known soluble PPases [12], and Family II
comprises recently discovered Bacillus subtilis PPase as well
as PPases of four other putative members, two streptococcal
and two archeal [13,14]. These two families do not show
any sequence similarity to each other. Family I soluble

PPases have been further divided into three subfamilies,
Correspondence to N. Tsuji, Laboratory of Parasitic Diseases,
National Institute of Animal Health, National Agricultural Research
Organization, 3-1-5 Kannondai, Tsukuba, Ibaraki 305-0856, Japan.
Fax: + 81 29 8387749, Tel.: + 81 29 8387749,
E-mail: tsujin@affrc.go.jp
Abbreviations: PPase, inorganic pyrophosphatase; H
+
-PPase, proton-
translocating pyrophosphatase; AsPPase, Ascaris suum inorganic
pyrophosphatase; rAsPPase, recombinant A. suum inorganic pyro-
phosphatase; L3, third-stage infective larvae; L4, fourth-stage larvae;

ES, excretory and secretory; IDP, imidodiphosphate.
Enzyme: Soluble inorganic pyrophosphatase (EC 3.6.1.1).
Note: The nucleotide sequences reported in this paper has been
submitted to the DDBJ/EMBL/GenBank with accession number
AB091401.
(Received 4 March 2003, revised 7 May 2003, accepted 9 May 2003)
Eur. J. Biochem. 270, 2814–2826 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03658.x
prokaryotic, plant and animal/fungal PPases. Among the
subfamilies, plant PPases bear a closer similarity to prok-
aryotic than to animal/fungal PPases [12]. The H
+
-PPases,

which appear to work as a reversible H
+
-pump, are much
larger and do not have any sequence similarity to either of
the two families of soluble PPases [15–17]. All known soluble
PPases are homologous proteins, whose active site residues
are highly conserved evolutionarily [18]. Site-directed mut-
agenesis and high-resolution X-ray crystallography studies
on PPases from E. coli and Saccharomyces cerevisiae have
implicated 17 amino acid residues as being important for
enzyme activity [19,20]. However, depending on the choice
of alignment parameters, 14–17 of the putative active site

residues described by Terzyan et al. [21] are conserved in
sequence alignments of E. coli and yeast PPases [18]. Recent
studies have demonstrated that only 17 residues are
conserved in all currently known Family I soluble PPases,
of which 13 are functionally important active site residues
[12,13]. PPases are strongly divalent metal ion-dependent,
with Mg
2+
conferring the highest PP
i
hydrolysis activity
[22]. Mg

2+
has several roles: it activates the enzyme and, as a
Mg
2+
-PP
i
complex, forms a true substrate for soluble
PPases; Mg
2+
also stabilizes the enzyme. Ca
2+
is reported

to be a potent inhibitor of soluble PPases [23].
While PPases from diversified sources have been des-
cribed in some detail, no PPase has ever been studied in any
metazoan helminth parasite including the roundworm
Ascaris. To address this, we describe here the cloning,
sequencing and heterologous expression in E. coli of a gene
encoding PPase of A. suum. The amino acid sequence of
A. suum PPase (AsPPase) indicates that it is an authentic
member of the Family I soluble PPases. We also provide
information concerning the kinetics and properties of the
enzyme. More strikingly, we show a novel role of the PPase
enzyme in the development and molting process of A. suum

larvae in vitro.
Materials and methods
Parasites
Adult A. suum were obtained from infected pigs at a
slaughterhouse in Shimotsuma, Japan. Adult A. lumbric-
oides and T. canis were obtained from patients after
treatment with piperazine in Bac Gian, Vietnam and, from
an infected dog in Miyazaki, Japan, respectively. Unem-
bryonated and embryonated eggs were obtained essentially
as described elsewhere [24]. Third-stage infective larvae (L3)
from embryonated eggs and lung-stage L3 were obtained as
described previously [25,26]. Excretory and secretory (ES)

products from L3, lung-stage L3 and adult worms were
collected as described previously [27]. Animal studies were
performed in accordance with the approval of the National
Institute of Animal Health Animal Care and Use Commit-
tee (Approval no. 23).
RNA was isolated from embryonated eggs using an
RNA isolation kit (Clontech). Poly(A)
+
mRNA was
prepared from total RNA using a polytract mRNA
isolation kit (Clontech) and first-strand cDNA synthesis
was performed using a cDNA synthesis kit and an oligo

(dT)
15
primer from Amersham Pharmacia Biotech. An
A. suum adult female worm cDNA expression library was
constructed in UniZap XR vector (Stratagene) according to
the manufacturer’s instructions as previously described [28].
Protein concentrations of NaCl/P
i
-soluble parasite antigens
andESproductsweremeasuredusingMicroBCAprotein
assay reagent (Pierce).
Immunoscreening of a cDNA expression library

An adult female worm cDNA expression library was
immunoscreened with rabbit antibodies raised against
A. suum embryonated egg trickle inoculations. Phages were
plated onto a lawn of E. coli XL-1 Blue at a density of
50 000 phage per dish and grown at 37 °C for 4 h. When
plaques were visible, isopropyl thio-b-
D
-galactoside-impreg-
nated filters were placed on the plates for 3 h to obtain a
plaque lift. After blocking in Tris/HCl, pH 8.0, with 0.05%
Tween 20, the filters were incubated in rabbit immune sera
overnight at 4 °C. Antibody reactivity with recombinant

proteins was revealed by incubation of the filters with
alkaline phosphate-conjugated goat anti-rabbit IgG (ICN)
for 1 h and developed with 5-bromo-4-chloro-3-indolyl-
phosphate/nitroblue tetrazolium (Gibco/BRL). Clones that
were reactive with the antibody were plaque-purified by
repeated cycles of immune selection. Plaque-purified clones
were converted using ExAssist
TM
helper phages and SOLR
E. coli (Stratagene) according to the in vivo excision
protocol described in the Stratagene ZAP-cDNA Synthesis
Kit (Stratagene). The nucleotide sequences of the cDNAs

were determined by the Sanger dideoxy chain termination
method using a PRISM
TM
Ready Dye Terminator Cycle
Sequencing Kit (PerkinElmer). DNA samples were ana-
lyzed using an automated sequencer (373A DNA sequencer,
Applied Biosystems).
BLASTX
(NCBI, National Institute of
Health) searches were performed to obtain cDNA clones
coding low similarity against mammalian proteins stored at
the current database. The

GENETYX
-
WIN
TM
DNA Sequence
Analysis Software System (Software Inc) and the
BLAST
network server of the National Center for Biotechnology
Information (NCBI) were used to analyze the nucleotides
and deduce the amino acid sequences in determining
similarities with previously reported sequences in GenBank.
A primary sequence motif was identified using the

PROSITE
network server at EMBL. Analysis of the signal sequence
was performed using
SIGNALP
v1.1 at the Center for
Biological Sequence Analysis ( />services/SignalP/index.html). Sequences were aligned by the
program
CLUSTALW
1.8 ( />with the
BLOSUM
amino acid substitution matrix using gap
penalties of 10.0 and 0.05 for gap opening and extension,

respectively. Phylogenetic trees were generated from
homologies of the PPase amino acid sequences by the
neighbor-joining method and the confidence of the branch-
ing order was verified by making 1000 bootstrap replicates
using the program
CLUSTALW
1.8. The tree was viewed
and converted to graphic format with
TREEVIEW
(http://
taxonomy.zoology.gla.ac.uk/rod/treeview.html).
Expression and purification of recombinant AsPPase

proteins
A full length cDNA (lacking signal peptides) was amplified
by PCR as previously described [29]. A sense primer
Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2815
(5¢-CCGAGCTCGAGACGTGAAGCGACAATCTCGC
AATCT-3¢) containing an XhoI (Promega) site upstream of
the start codon and an antisense primer (5¢-CAGCCAA
GCTTCTCACTCTTTGATGAAATGCATCT-3¢) con-
taining a HindIII (Promega) site just downstream of amino
acid residue were used. The PCR fragments digested with
XhoIandHindIII were ligated into plasmid expression
vector pTrcHisB

TM
(Invitrogen), which had also been
digested with the same enzymes according to the manufac-
turer’s instructions. The resultant plasmid was transformed
into E. coli strain TOP10F¢ (Invitrogen). The transformed
cellsweregrowntoaD
600
at 37 °CinSOBmedium
supplemented with 50 lgÆmL
)1
ampicillin. To induce pro-
tein expression, isopropyl thio-b-

D
-galactoside was then
added to a final concentration of 1 m
M
and the culture was
grown for an additional 4 h at 37 °C. The E. coli cells were
pelleted and resuspended in lysis buffer [50 m
M
NaH
2
PO
4

(pH 8.0), 10 m
M
Tris/HCl (pH 8.0), 100 m
M
NaCl]. Lyso-
zyme was added at 100 lgÆmL
)1
, and the cell suspension
was incubated on ice for 15 min. The cell suspension was
disrupted using an ultrasonic processor (VP-5, TAITEC) on
ice. The E. coli lysate was centrifuged at 26 000 g for 30 min
at 4 °C. The supernatants containing recombinant proteins

of AsPPase were purified using ProBond
TM
resin (Invitro-
gen) under nondenaturing conditions and subsequently
eluted with a stepwise gradient of imidazole (50–500 m
M
).
The eluted fractions were concentrated by Centrisart I (cut
off MW 10 000; Sartorius) and then dialyzed extensively at
4 °C with several successive changes of 20 m
M
Tris/HCl,

pH 7.5 and a decreasing concentration of NaCl in a Slide-
A-Lyzer Dialysis Cassette (Pierce). Fractions were collected
and the presence and purity of recombinant protein was
detected by 10% SDS/PAGE [30] and immunoblot [31]
using anti-T7 tag Ig (Invitrogen). Protein concentrations
were measured with the Micro BCA protein assay reagent
(Pierce).
Production of mouse polyclonal antibodies
BALB/c mice were immunized first with a subcutaneous
injection of 50 lg of recombinant AsPPase (rAsPPase)
emulsified with TiterMax Gold
TM

(CytRx), followed by
another injection 2 weeks later in the same adjuvant. The
mice were bled 2 weeks after the second injection. The
antiserafromthemiceweremixedandstoredat)20 °C
until used. Anti-(mouse rAsPPase) IgG from immune sera
and mouse preimmune IgG were affinity purified using
UltraLink
TM
immobilized protein G according to manu-
facturer’s instructions (Pierce) and used for evaluating the
native AsPPase-neutralizing activity.
Two-dimensional electrophoresis

Parasite extracts were treated with an equal volume of urea
mixture consisting of 9
M
urea, 4% Nonidet P-40, 0.8%
ampholine (pH 3.5–10; Pharmacia) and 2% 2-mercapto-
ethanol, and then subjected to 2D PAGE. Nonequilibrium
pH gradient electrophoresis was performed [32] in the first
dimension using a rectangular gel electrophoresis apparatus
(AE-6050 A; ATTO). After electrophoresis at 400 V for
2 h, the gels were incubated in the equilibration buffer
for 10 min on a shaker. Electrophoresis in the second
dimension was performed on 8% SDS/PAGE gels under

reducing conditions. The proteins were either stained using
a silver staining kit (Dai-ichi Pure Chemicals) or transferred
to nitrocellulose membranes.
Immunoblot analysis
Immunoblot analysis was carried out as previously des-
cribed [29]. Anti-(mouse rAsPPase) serum was used at a
dilution of 1: 500. The proteins bound to the secondary
antibody were visualized with 5-bromo-4-chloro-3-indolyl-
phosphate/nitroblue tetrazolium.
Immunohistochemistry
Adult females of A. suum and A. lumbricoides were fixed in
4% paraformaldehyde in 0.1

M
phosphate buffer, pH 7.2,
overnight and embedded in paraffin. Thin transverse
sections were made from paraffin-embedded fixed worms.
The sections on glass slides were deparaffinized and
dehydrated using a graded series of alcohol and then
rehydrated in NaCl/P
i
. The slides were blocked for 30 min
in 1% H
2
O

2
in NaCl/P
i
containing 10% ethanol to inacti-
vate endogenous peroxidases. For immunolocalization, the
slides were blocked in NaCl/P
i
containing 10% (v/v) goat
serum (Wako) for 30 min at room temperature. They were
then flooded with anti-(mouse rAsPPase) Ig diluted 1 : 100
in NaCl/P
i

/E. coli lysate, overnight at 4 °C. Afterwards, the
slides were rinsed thoroughly with NaCl/P
i
,andthe
antibody binding was resolved with a peroxidase-labeled
anti-mouse IgG and the substrate 3¢,3¢-diaminobenzidine
tetrahydrochloride (Sigma Fast
TM
tablets, Sigma). After
color development, the slides were dehydrated in a graded
series of alcohol and cleared in xylene. The slides were then
covered with cover slips and observed with a microscope

(Axiophot; Carl Zeiss).
Enzyme assay
The rAsPPase activity was determined spectrophotometri-
cally by measuring the rate of liberation of P
i
from PP
i
using a molybdate-blue based colorimetric assay [33]. The
recombinant protein was assayed in the standard reaction
mixture containing 5 m
M
Mg

2+
, 100 m
M
Tris/HCl
(pH 7.5) and 1 m
M
PP
i
(Na
4
P
2

O
7
), in a total volume of
200 lL together with the protein solution, at 55 °C. The
assay was started by adding 10 lL of diluted rAsPPase
solution into the standard reaction mixture. The reaction
was stopped by adding 1 mL of 200 m
M
glycine/HCl,
pH 3.0. Then, 125 lL of 1% ammonium molybdate (in
25 m
M

H
2
SO
4
)and125lL of 1% ascorbic acid (in 0.05%
KHSO
4
) were added to the mixture, and incubated for
30 min at 37 °C. Protein concentrations and reaction times
were chosen in order to obtain the linearity of the reactions.
As positive and negative controls, pure yeast-soluble PPase
from Sigma (1-1643) and an unrelated A. suum 14-kDa

recombinant protein (As14; [28]) were used, respectively.
The concentrations of individual components were varied as
indicated for the determination of Mg
2+
and pH dependent
rAsPPase activity. The amount of P
i
liberated from the
hydrolysis of PP
i
during the course of the reaction was
measured in comparison to a standard P

i
sample using a
2816 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003
spectrophotometer (Model 600, Shimadzu) at an optical
density of 700 nm. The specific activity of rAsPPase was
defined as lmol P
i
released min
)1
Æmg
)1
of protein.

Enzyme kinetic study
The K
m
(Michaelis constant) and V
max
(maximum velocity)
values were determined by incubating the diluted recombin-
ant proteins in the standard reaction mixture in the presence
of increasing concentrations of PP
i
(0.05–0.5 m
M

)at55°C.
Data were fit to the appropriate equation using
GRAFIT
version 3.09b (Erithacus Software). K
m
and V
max
values were
reported with their standard errors derived from the fit.
Native AsPPase activity and NaF sensitivity
To investigate the native AsPPase activity during A. suum
larval development and molting, the L3 soluble extracts (in

20 m
M
Tris/HCl, pH 7.5) and the L3 ES products in the
culture fluids (dialyzed against 20 m
M
Tris/HCl, pH 7.5),
were assayed in the standard reaction mixture as described
above. Anti-(mouse rAsPPase) IgG were evaluated for
AsPPase-neutralizing activity. Recombinant AsPPase pro-
teins or A. suum L3 extracts were preincubated in the
standard reaction mixture containing 5 lgÆmL
)1

pre-
immune or anti-(mouse rAsPPase) IgG (15 min, 37 °C)
before adding PP
i
. The sensitivity of the native AsPPase to
inhibition by sodium fluoride (NaF, S-7920; Sigma) was
tested in the present study. The L3 extracts were assayed in
the standard reaction mixture for PP
i
hydrolysis, in the
presence of increasing concentrations of NaF. The PPase
activated PP

i
hydrolysis rate was then calculated.
Larval molting inhibition assay
To confirm whether the native PPase enzyme is involved in
the molting process, we examined the effects of two PPase
specific inhibitors, imidodiphosphate (IDP, 1-0631; Sigma)
and NaF on development and molting of A. suum lung-
stage L3 to fourth-stage larvae (L4) in vitro. The lung-stage
L3 were obtained from the lungs of New Zealand white
rabbits 7 days after inoculation with 2.5 · 10
5
embryonated

infective eggs of A. suum [26]. Briefly, the rabbits were killed
by an overdose of ketamine hydrochloride (50 mgÆkg
)1
body weight, i.v.) followed immediately by decapitation and
the lungs were removed and minced with a surgical knife.
The minced tissue was wrapped in cotton gauze and
suspended in NaCl/P
i
containing 100 lgÆmL
)1
penicillin/
streptomycin at 37 °C for 3 h. After incubation the tissues

were removed, and the larvae were collected from the
bottom of the tube. The recovered L3 were washed several
times with warm NaCl/P
i
containing 50 lgÆmL
)1
penicillin/
streptomycin, and subjected to molting inhibition assay.
Briefly, 50–100 L3 in 1 mL of RPMI 1640 medium (Gibco/
BRL), pH 6.8 supplemented with 10% (v/v) fetal bovine
serum (Sigma), 50 lgÆmL
)1

penicillin/streptomycin were
cultured in 24-well flat-bottomed tissue culture plates
(CostarÒ). The cultures were incubated at 37 °Cina
humidified 5% CO
2
incubator in the absence (control) and
presence of increasing concentrations of inhibitors for
10 days, and the number of molting larvae was determined.
Molting was manifested by shedding of the L3 cuticle.
Numbers of molted larvae in a culture well were therefore
determined by counting the L3 cuticles shed from the larvae.
Furthermore, molted larvae (that had already shed their

cuticles) exhibited an intense motility compared with
unmolted larvae (that had not shed their cuticle). Aliquots
of larvae were removed at different days of postcultures and
photographs were taken.
Results
Identification of cDNA encoding
A. suum
inorganic
pyrophosphatases
A clone designated AdR44 was isolated initially by immu-
noscreening an A. suum female worm cDNA library with
serum obtained from a rabbit immunized with A. suum

infective eggs. AdR44 was selected for further characteriza-
tion because of its sequence homology to the inorganic
pyrophosphatase family of proteins. Sequence analysis
showed that AdR44 was 1,375-bp long with an open
reading frame (ORF) coding for 360 amino acids. The ATG
initiation codon is predicted to be at nucleotides 79–81 and
is followed by a region encoding a hydrophobic sequence of
17 amino acids, which may function as a signal peptide. The
3¢ untranslated region contained 224 bp and ended with
17-bp poly(A)
+
tail that began 14-bp down-stream from the

sequence AATAAA, which is the eukaryotic consensus
polyadenylation signal. An entire ORF of the AdR44
cDNA encodes a sequence of 360 amino acids, predicting a
40 600-Da polypeptide with an isoelectric point of 7.1.
Removal of the signal peptide resulted in a putative mature
protein with molecular weight 38 771 Da. Two potential
sites for N-glycosylation (residues 50–53, 246–249) were
predicted in AsPPase. The three conserved aspartates that
are involved in the binding of cations in PPases (D-[SGDN]-
D-[PE]-[LIVMF]-D-[LIVMGAC]) were found at position
192–197. A search of the protein database conducted using
the information obtained from the NCBI revealed that

AsPPase shared a high degree of sequence similarity to those
of animal/fungal PPases in Family I soluble PPases.
Figure 1 shows a comparison of the AsPPase sequence to
five other sequences of animal/fungal soluble PPases. The
deduced amino acid sequence of AsPPase shows 74%
similarity (56% identical) to the Drosophila melanogaster
PPase sequence, 69% similarity (55% identical) to the
sequence of Caenorhabditis elegans, 72% similarity (55%
identical) to the sequence of Schizosaccharomyces pombe,
70% similarity (51% identical) to the sequence of Bos taurus
and 67% similarity (51% identical) to the sequence of
S. cerevisiae. Sequence similarities occur throughout the

protein but few are at both ends. Sequence analysis revealed
that all 13 functionally important active site residues
(AsPPase numbering: E-125, K-133, E-135, R-155, Y-170,
D-192, D-194, D-197, D-224, D-229, K-231, Y-269 and
K-270) (Fig. 1), which have been reported previously to be
evolutionarily well conserved in Family I soluble PPases
[12,13,18,34,35], are identical in AsPPase.
Phylogenetic analysis of available PPases
We have constructed a phylogenetic tree using Family I
soluble PPase sequences by the neighbor-joining method
Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2817
and the confidence of the branching order was verified by

making 1000 bootstrap replicates with the
CLUSTALW
program (Fig. 2). The neighbor-joined trees reveal that
animal and fungal PPases including AsPPase represent a
separate group from plant and prokaryotic PPases. Fur-
thermore, within the animal/fungal subgroup, AsPPase is
more closely clustered with PPases from the free-living
model nematode C. elegans and the insect D. melanogaster.
Characterization of recombinant AsPPase
The gene encoding the soluble PPase of A. suum was
amplified by PCR with A. suum female worm cDNA.
AsPPase was then overexpressed in E. coli strain TOP10F¢

using pTrcHisB
TM
vector, to test whether the clone indeed
has an inorganic pyrophosphatase activity. Recombinant
AsPPase was expressed in E. coli with a yield of 1 mgÆL
)1
of
bacterial culture. The rAsPPase was 99% pure as determined
by SDS/PAGE analysis. The observed molecular mass of
rAsPPase corresponded well to the calculated mass of the
AdR44 cDNA (data not shown). The functional activity of
the purified rAsPPase was determined using a PP

i
hydrolysis
assay in a standard reaction mixture containing 5 m
M
Mg
2+
,
100 m
M
Tris/HCl, pH 7.5 and 1 m
M
PP

i
. The recombinant
protein showed a specific activity of 937 lmol P
i
Æmin
)1
Æmg
)1
protein corresponding to a k
cat
value of 638 s
)1

that could be
abolished by Ca
2+
or removal of Mg
2+
(Table 1). This
activity could not be due to copurification of endogenous
E. coli PPase as, the rAsPPase contained a His-tag that was
used for purification and the recombinant protein was
determined to be pure by SDS/PAGE analysis.
Expression and immunohistochemical detection
of native AsPPase

We performed 2D immunoblot analysis to identify native
AsPPase in adult female A. suum. Anti-(mouse rAsPPase)
serum reacted strongly with a protein having a molecular
mass of 39 kDa with a pI of 7.1 (Fig. 3A) confirming that it
corresponded to the predicted size of the putative mature
protein (38.771 kDa) calculated from the AsPPase amino
acid sequence except for a signal peptide. In addition, a native
AsPPase was identified on silver-stained 2D gels on which
more than 200 visible protein spots appeared (data not
shown). To determine the N-terminal residues, we excised the
native AsPPase spots from 2D immunoblotted polyvinylid-
ene difluoride membranes and subjected them to analysis by

the automatic Edman degradation method. The sequence 1-
MALAASATIS-10 of native AsPPase was identical to that
of the putative mature protein. This confirmed that our clone
encoded a soluble PPase of A. suum. A spot reacting with the
anti-mouse rAsPPase was also seen in parasite extracts and
ES products from various developmental stages, including
embryonated eggs, L3, lung-stage L3 and adult male and
female worms, indicating that native AsPPases were
expressed in all lifecycle stages of A. suum (data not shown).
Interestingly, enzyme homologs were also expressed in the
human roundworm A. lumbricoides and the dog roundworm
T. canis (Fig. 3B). Native AsPPases that reacted with mouse

polyclonal antibodies against rAsPPase were localized in
various structures such as the hypodermis, dorsal and lateral
hypodermalchord, in muscle cells, gut epithelium and, in the
uterus and ovary of adult female A. suum (Fig. 4B–D). No
labeling was, however, seen in sections probed with mouse
preimmune sera (Fig. 4A). This study also detected the
ubiquitous presence of AsPPase homologs in various organs
of A. lumbricoides (data not shown).
Enzymatic properties of recombinant proteins
The PP
i
dependence of the maximum hydrolytic velocity

(V
max
) of the recombinant AsPPase protein was shown to
be 849.005 ± 14.635 lmol P
i
Æmin
)1
Æmg
)1
protein with a
K
m

(Michaelis Constant) value of 0.117 ± 0.006 m
M
for PP
i
from three independent experiments (Fig. 5A).
The K
m
value is significantly higher than the values of
Fig. 1. Sequence alignment of representative members of Family I sol-
uble PPases.
CLUSTALW
alignment of soluble PPases (GenBank

accession numbers are indicated in parentheses): A. suum (AB091401),
C. elegans (CAA93107), D. melanogaster (O77460), Bos taurus
(P37980), S. cerevisiae (2781300) and Schizosaccharomyces pombe
(P19117). Identical residues among PPases are marked with asterisks.
The 13 essential, active site residues that are conserved in all Family I
soluble PPase sequences currently available in the GenBank are further
emphasized by bold typeface. The signal peptides are underlined.
Dashes indicate gaps inserted to optimize the alignment. The num-
bering is for the sequence of A. suum (As) PPase. As, A. suum,Ce,
C. elegans,Dm,D. melanogaster,Bt,B. taurus,Sc,S. cerevisiae,Sp,
S. pombe.
2818 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003

0.0009–0.00147 m
M
from bovine retinal PPase [23],
0.005 m
M
from rat liver PPase [36] and 0.026 m
M
from
bovine rod outer segment PPase [37]. These discrepancies in
K
m
values for PP

i
are not entirely surprising in view of the
many differences in enzyme purity and assay methodology.
The rAsPPase was shown to absolutely require Mg
2+
for
PP
i
hydrolysis. The maximum enzyme activity was found
with 5 m
M
Mg

2+
using 1 m
M
PP
i
, which then gradually
declined with increased concentrations of Mg
2+
(Fig. 5B).
A drop in enzyme activity due to increased concentrations
of Mg
2+

(> 5 m
M
Mg
2+
) has not been investigated in the
present study. Although excess of Mg
2+
is known to inhibit
the PPases, however, the mechanism is yet unclear. Both
E. coli and yeast PPases have four (M1–M4) subsites for
binding metal ions for catalysis [38,39]. It has been urgued
that binding of Mg

2+
at three subsites is required for
Fig. 2. Phylogenetic tree based on alignment of available Family I soluble PPase sequences. The sequences shown are those from (GenBank accession
numbers are indicated in parentheses): A. suum (AB091401), S. cerevisiae (2781300), Kluyveromyces lactis (P13998), Pichia pastoris (O13505),
S. pombe (P19117), D. melanogaster (O77460), C. elegans (CAA93107), B. taurus (P37980), Homo sapiens,from([12]),S. cerevisiae mitochondria
(P28239), Hordeum vulgare (O23979), Zea mays (O48556), Solanum tuberosum (O43187), Arabidopsis thaliana (AAC33503), Oryza sativa
(AAC78101), Chlamydia pneumoniae (AAD19056), Chlamydia trachomatis (O84777), Mycoplasma pneumoniae (P75250), Mycoplasma genitalium
(P47593), Bacillus stearothermophilus (BAA19837), Synechocystis (PCC6803, P80507), Thermoplasma acidophilum (P37981), Methanobacterium
thermoautotrophicum (O26363), Thermococcus litoralis (P77992), Pyrococcus horikoshii (O59570), T. thermophilus (P38576), Mycobacterium leprae
(O69540), Mycobacterium tuberculosis (CAB08851), Haemophilus influenzae (1170585), Sulfolobus acidocaldarius (P50308), Aquifex aeolicus
(O67501), Helicobacter pylori (P56153), Gluconobacter suboxydans (O05545), Bartonella bacilliformis (P51064), Rickettsia prowazekii (CAA15034),
Legionella pneumophila (O34955) and E. coli (P17288). The bar indicates the numbers of substitutions per site. Unrooted neighbor-joining trees

were generated from homologies of soluble PPase sequences and the confidence of the branching order was verified by making 1000 bootstrap
replicates using the
CLUSTALW
program. The tree was viewed and converted to graphic format with
TREEVIEW
.
Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2819
catalysis to proceed, whereas binding at M4 causes inhibi-
tion [38,40]). Omission of Mg
2+
from the reaction medium
abolished PPase-mediated PP

i
hydrolysis. The enzyme
activity was found to be optimum in the pH range 7.0–8.0
(Fig. 5C). The pH profile showed a dramatic drop in
activity at high pH. These results are consistent with other
soluble PPases from various sources [35,36,41].
Detection of native AsPPase activity and inhibition
by IDP and NaF of larval development and molting
Native enzyme activity in L3 soluble extracts and in
ES products was detected by PPase-activated PP
i
hydroly-

sis. The L3 extracts showed an activity of 1.58 ± 0.02 lmol
P
i
Æmin
)1
Æmg
)1
, whereas L3 ES exhibited that of
0.51 ± 0.03 lmol P
i
Æmin
)1

Æmg
)1
protein for PP
i
hydrolysis.
Anti-(mouse rAsPPase) IgG partially inhibited recombinant
AsPPase (6 ng) activity up to 22% in the presence of
5 lgÆmL
)1
anti-(mouse rAsPPase) IgG relative to AsPPase
activity determined in the presence of 5 lgÆmL
)1

mouse
preimmune IgG (data not shown). Also, native AsPPase
activity in L3 extracts was shown to be partially inhibited
(25%) by anti-(mouse rAsPPase) IgG (data not shown)
indicating that AsPPase is responsible for the hydrolyzing
activity of PP
i
in A. suum L3 extracts. NaF, an anion, is a
potent inhibitor of PPases and was able to inhibit native
AsPPase activity at micromolar concentrations in a dose-
dependent manner (Fig. 6A). This agent is also known to
inhibit the H

+
-PPases from plants [42], trypanosomatids
[43,44] and apicomplexan protozoa [45].
The L3 of A. suum develop and molt to L4 in the lungs of
their vertebrate hosts that can also occur during in vitro
cultivation. To determine whether this complex process is
regulated by PPase enzyme, we examined IDP, a non-
hydrolyzable PP
i
analogue, and NaF for their possible
in vivo ability to inhibit/arrest development and the molting
process by blocking the PPase activated PP

i
hydrolysis, as
the native PPases in L3 extracts were found to be very
sensitive to inhibition by NaF (Fig. 6A). As IDP interferes
with the colorimetric assay, it was, however, not possible
to examine enzyme sensitivity with this compound in the
present study. In vitro molting inhibition experiments
demonstrated that IDP and NaF inhibited molting of
A. suum L3 to L4 with varying success, in a dose-dependent
manner (Fig. 6B,C). Up to 55% molting was inhibited at a
maximum concentration of 10 m
M

IDP without affecting
the growth and viability of the L3. In contrast, NaF
inhibited 65% molting at 1 m
M
concentration. However, at
higher concentrations (>1 m
M
NaF) molting inhibition was
increased drastically up to 100% with an apparent growth
inhibition of the L3 observed on day 5 postculture and
onwards. A mild larvicidal effect of 10 m
M

NaF with
progressive damage of the body wall and intestine was seen
on day 5 postculture and onwards (data not shown). The
molted L3 developed well to L4 in control culture with
increased body length and width (data not shown), and
changes in the structure of the head and tail (Fig. 7A–C)
compared with unmolted L3 which achieved little or no
development, being inhibited by IDP/NaF (Fig. 7D,E).
Under light microscopy, it was however, not possible to
detect the formation and/or separation of new cuticles of
unmolted L3 exposed to inhibitors that might be carried out
by electron microscopy. The mean molting percentage in

control culture was recorded as, 52.59 ± 4.12.
Discussion
Although PPases are distributed widely among living cells,
most of the previous studies have focused on microbial and
plant enzymes, and very little is known about the enzyme
Fig. 3. Identification of A. suum native PPase in adult female worm.
(A) Fifty micrograms of female worm extract was separated by 2D
nonequilibrium pH-gradient gel electrophoresis, and the proteins were
then transferred to a nitrocellulose membrane. The native AsPPase
bound to the anti-(mouse rAsPPase) serum was found by alignment of
the stained gel and immunoblot membrane. (B) Expression of AsPPase
homologs in ascarid roundworms. Sixty or 80 mg of protein equiva-

lents of each parasite extract were electrophoresed on a 10% SDS/
PAGE and blotted onto a nitrocellulose membrane. The AsPPase
homologs bound to the anti-(mouse rAsPPase) serum were detected by
5-bromo-4-chloro-3-indolylphosphate/nitroblue tetrazolium. Lane 1,
A. lumbricoides;lane2,T. canis;lane3,A. suum.
Table 1. Recombinant A. suum PPase activity. Pyrophosphatase
activity was assayed as described in the Materials and methods section;
–, no detectable activity. Data represent the mean ± SE from three
independent experiments. An unrelated A. suum 14-kDa recombinant
protein was used as the negative control (As 14; [28]).
Assay conditions
Activity

(lmol P
i
Æmin
)1
Æmg protein
)1
)
A. suum PPase +5.0 m
M
Mg
2+
937.76 ± 39.76

A. suum PPase +0.0 m
M
Mg
2+
–
A. suum PPase +5.0 m
M
Ca
2+
–
Yeast-PPase +5.0 m
M

Mg
2+
(positive control)
13232.36 ± 183.42
As14 + 5.0 m
M
Mg
2+
(negative control)
–
2820 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003
from mammalian tissues. In contrast, we do not have any

evidence of PPases from parasitic helminths. We report here
the cloning, sequencing and biochemical and functional
characterization of a novel PPase from the important
pathogenic roundworm A. suum. The deduced amino acid
sequence of AsPPase shows significant similarity with
animal/fungal PPase sequences in Family I soluble PPases
(Fig. 1). All members of Family I soluble PPases currently
available in the database have been shown to contain 13
functionally important active site residues that are evolu-
tionarily well conserved, and were found to be identical in
AsPPase. Several highly conserved regions, the most
prominent of which is an eight residue long sequence

(224-DEGETDWK-231), are also seen in the AsPPase
sequence. It will be interesting to see the significance of this
highly conserved region in AsPPase structure and function-
ing. Over 37 Family I soluble PPases have been identified.
The prokaryotic PPases are hexamers of  20 kDa and
reported to contain 162–220 amino acid residues per
subunit, while eukaryotic PPases are dimers of 30–36 kDa
with 211–310 residues per subunit [12,13,18]. AsPPase, with
360 amino acid residues having a calculated molecular mass
of 40.6 kDa, resembles eukaryotic PPases and thus is the
largest among the Family I soluble PPases stored in the
current protein database. This is largely due to a longer

N-terminal region compared with other PPases (Fig. 1).
The membrane-bound H
+
-PPases that are found in plants
[17], certain bacteria [46], and more recently identified from
trypanosomatids [47] and apicomplexan protozoa [48] differ
greatly in structure and function from soluble PPases. The
H
+
-PPases are much larger (660–770 amino acid residues
per monomer) and do not have any sequence similarity to
soluble forms [15,16,49]. The AsPPase described here is

clearly a soluble PPase and it does not have any sequence
similarity to plant/protist H
+
-PPases. Together, these
findings suggest that AsPPase is a distinct Family I soluble
PPase. Phylogenetically, AsPPase is, within the subfamily
of animal/fungal soluble PPases, closer to C. elegans and
D. melanogaster PPases than to fungal and mamma-
lian PPases (Fig. 2). Moreover, E. coli-expressed purified
rAsPPase protein has shown enzymatic activity
(937 lmol P
i

Æmin
)1
Æmg protein
)1
)byPP
i
hydrolysis assay
that was found to be closer to those of the highly purified
and crystallized E. coli (2000 lmol P
i
Æmin
)1

Æmg
)1
[50]),
yeast (655 lmol P
i
Æmin
)1
Æmg
)1
[51]), rat liver 600–
700 lmol P
i

Æmin
)1
Æmg
)1
[36]) and bovine retinal PPases
(> 8 8 5 lmol P
i
Æmin
)1
Æmg
)1
[23]). Thus, AsPPase represents

the first member of Family I soluble PPase enzymes to be
identified from the parasitic helminths.
The rAsPPase activity was shown to be strictly Mg
2+
-
dependent. On the contrary, Ca
2+
inhibited the activity to
some degree in the presence of Mg
2+
(data not shown).
These distinctive features have been well demonstrated for

Family I soluble PPases [23,52,53]. The rAsPPase enzyme,
however, requires a higher concentration of Ca
2+
for
inhibition (data not shown), and this finding is fairly
Fig. 4. Immunohistochemical localization of A. suum native PPase in adult female worm. A. suum female worms were fixed in paraformaldehyde,
embedded in paraffin, sectioned (7-lm thickness) and exposed to either mouse preimmune serum as a control (A) or mouse anti-rAsPPase serum
diluted 1: 100 (B). (C) and (D) (both 25·) are magnified areas of (B). Arrows indicate antibody-labeled regions; cu, cuticle; hd, hypodermis;
hc, hypodermal chord; mu, muscle; gu, gut; ov, ovary; ut, uterus.
Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2821
consistent with other animal PPases [36,54] but contrasts
with the reports on yeast PPase and on porcine brain and

bovine retinal PPases [23,41] that demonstrated much lower
concentrations of Ca
2+
were needed for enzyme inhibition.
Prior studies have shown that free PP
i
is a potent inhibitor,
and free Mg
2+
activates the enzyme and binds with PP
i
to

form a true substrate, Mg
2+
PP
i
for soluble PPases [38,55].
Family II PPases are easily distinguishable from Family I
PPases in having a preference for Mn
2+
over Mg
2+
as the
activator, and are not inhibited by Ca

2+
,ratherCa
2+
Fig. 5. PP
i
(A), Mg
2+
(B) and pH (C) dependence of A. suum recom-
binant PPase activity. (A) Diluted recombinant proteins were run in the
standard reaction mixture (as described in Materials and methods) for
PPase assays at 55 °C, in the presence of increasing concentrations of
PP

i
(0.05–0.5 m
M
). Data were analyzed using a computer assisted
program (
GRAFIT
version 3.09b). The theoretical curve drawn is for
thebestfitvaluesofK
m
¼ 0.117 ± 0.006 m
M
,andV

max
¼
849.005 ± 14.635 lmolÆmin
)1
Æmg protein
)1
. The inset in (A) repre-
sents the linear transformation of the curve. (B) Mg
2+
dependency was
determined as described in (A) in the presence of increasing concen-
trations of Mg

2+
. (C) pH dependency of the enzyme was examined as
described above using several buffers with increasing pH values. The
buffers used were (100 m
M
), sodium acetate (pH 5.0–5.5), Mops
(pH 6.0–6.5), Tris/HCl (pH 7.0–8.5) and glycine/NaOH (9.0–10.5).
Data represent mean ± SEM from three independent experiments.
Fig. 6. Inhibition of A. suum native PPase activity and A. suum L3
molting by IDP and NaF. (A) Aliquots of A. suum L3 soluble extracts,
17 mg proteinÆmL
)1

was run in the standard reaction mixture
for PPase assays at 55 °C, in the presence of increasing concentrations
of NaF. Percentage activity compared to the control in the absence
of NaF (100%). Control activities were 1.58 ± 0.01 lmol
P
i
Æmin
)1
Æmg protein
)1
for PP
i

hydrolysis. Data represent mean ±
SEM from three independent experiments. (B) Lung-stage A. suum L3
were cultured for molting inhibition assays, in the presence of
increasing concentrations of IDP and (C) NaF. Molting percentage
was determined on day 10. Percentage activities are relative to the
control in the absence of inhibitor (100%). Molting percentage of
control was 52.59 ± 4.12. Data represent mean ± SEM of triplicates.
2822 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003
activates the enzymes preincubated with Mn
2+
[13,53]
further indicating that AsPPase reported here is an authen-

tic member of Family I soluble PPases.
An intense expression of native AsPPase in metabolically
active tissues, such as the body wall, gut epithelium and
reproductive organs, of adult female worms suggests a
critical role of the enzyme in these organs. The presence of
AsPPase in embryonated eggs, L3, lung-stage L3, adult
worms and their ES products together with its direct
detection in L3 soluble extracts and in ES products
(1.58 ± 0.02 lmol P
i
Æmin
)1

Æmg
)1
and 0.51 ± 0.03 lmol
P
i
Æmin
)1
Æmg
)1
protein for PP
i
hydrolysis, for L3 extracts and

ES products, respectively) strongly suggested the important
roles of the PPase enzyme throughout the developmental
cycle of Ascaris parasites. The results of neutralization
studies indicate that AsPPase-specific IgG may interfere
with the development and molting process of Ascaris larvae.
We showed that the native AsPPases are very sensitive to
inhibition by NaF in the micromolar range (Fig. 6A). This
value is much lower than previously reported data for NaF
against H
+
-PPases from parasitic protozoa [44]. These
results prompted us to investigate the role of PPase enzymes

in the development and molting process of A. suum larvae
and to test whether this could be targeted by inhibitors. We
used IDP, a nonhydrolyzable PP
i
analogue, and NaF, a well
known inhibitor of Family I and Family II soluble PPases
(NaF competes with the hydroxide ion for binding to Mg
2+
in the active site of the enzyme [35,53]), to block enzyme
activity. We demonstrated for the first time that NaF is
highly effective in inhibiting the development and molting of
A. suum L3 to L4, in a concentration-dependent manner,

whereas, IDP has shown only partial inhibitory effect
(Fig. 6B,C). However, a much higher concentration of NaF
(> 1 m
M
) is required to completely block development and
molting of L3 as against micromolar concentration is needed
for the inhibition of native enzyme in vitro. This difference
may in part be attributed to the difference between live
parasites and their soluble extracts used in the assay system.
We observed that during in vitro cultivation, the L3 could
not develop and molt to L4 in the presence of inhibitor, even
at the end when the culture had terminated (Fig. 7D,E).

These observations indicate that PPase enzymes are prob-
ably involved in the development and molting process of
A. suum L3 to L4. However, the mechanisms of inhibition of
this complex process by PPase inhibitors are yet to be
elucidated. Although aminopeptidase, cysteine protease and
hyaluronidase enzymes so far have been reported to be
involved in the development and molting process of A. suum
L3 to L4 in vitro, virtually none had been characterized in
relation to the actual mechanism of the molting process in
this roundworm [56]. The basic structure of the body wall of
parasitic roundworms consists of the cuticle, an underlying
syncytial or cellular layer called the hypodermis, and the

longitudinally oriented somatic musculature. The ecdysis of
an old cuticle and deposition of the components of a new
cuticle that are synthesized in the hypodermis and are
secreted across the hypodermal membrane into the space
between it and the old cuticle, occur at each of four molts
during the lifecycle of all roundworms [57]. The molecular
mechanisms regulating this complex process are, however,
Fig. 7. In vitro development and molting of A. suum lung-stage L3 to L4 in the absence of inhibitor (control; A–C) and its presence (D,E). (A) L3 on
day 0 culture from control. (B) L3 had initiated molting on day 5 postculture from control. Arrow indicates an entirely distended L3 cuticle. (C) L4
(molted L3) on day 10 postculture from control. A cuticle which had shedded from L3 is indicated by an arrow. (D) L3 had not initiated molting on
day 5 postculture with inhibitors, IDP/NaF. (E) L3 had not molted on day 10 postculture with IDP/NaF. Photographs were taken using differential
interference contrast microscopy.

Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2823
still poorly understood. Based on our results presented
above, it is assumed that the target molecules of IDP and
NaF might be PPase enzymes in the hypodermis of L3
(Fig. 4 indicates the abundance of native PPases in the
hypodermal cells of sectioned adult worms in immuno-
histochemical staining), and that the inhibition of PPase-
catalyzed PP
i
hydrolysis by these inhibitors, is likely to
prevent the synthesis of the new cuticle from the hypodermis
and/or ecdysis of the old cuticle. This assumption is

supported by the fact that PPase-activated PP
i
hydrolysis
is essential to maintain the forward direction of many
biosynthetic reactions like synthesis of DNA, RNA, proteins
and polypeptides [8]. In addition, several investigators have
demonstrated that both IDP and NaF selectively inhibited
the plant and protist H
+
-PPases [42–45] and other potential
PP
i

analogues (bisphosphonates) selectively inhibited the
proliferation of acidocalcisome-containing parasites [58].
Recent studies have shown that fluoride can inhibit several
metabolic and defending enzymes of microbial origin and
can alter expression of protein essential for survival and
virulence in unicellular bacteria [59–61]. It is thought
important that the hypodermis of the body wall, which
synthesizes components of the cuticle, may offer a useful
target for studies into the mechanism of the development
and molting process in the roundworms. Investigation into
the effects of fluoride exposure to A. suum L3 on protein
expression to explore the PPase-regulated development and

molting process of the Ascaris parasites is currently being
undertaken in our laboratory.
We conclude that PPase is a novel enzyme in A. suum
that may play an important role in the development and
molting of the larval stage parasites. Furthermore, the
biochemical as well as the in vivo/in vitro functional
characterization of this enzyme (this study) using some well
known PPase-specific inhibitors (e.g. NaF, IDP), and
rAsPPase-specific mouse IgG may give new insight into
the PPase-regulated worm metabolism/development and
may provide prospects for the design of novel chemothera-
peutic agents to control Ascaris and other geohelminth

parasites.
Acknowledgements
This work was supported by a grant from the Program for Promotion
of Basic Research Activities for Innovative Biosciences (PROBRAIN).
M. K. I. was supported by the Japan Society for the Promotion of
Science postdoctoral fellowship. We would like to thank Dr M. S. H.
Bhuiyan, Enzymology Laboratory of National Food Research Institute
for the enzyme assay protocol, Mr M. Kobayashi of the Histopatho-
logy Core Group for preparing paraffin-sectioned glass slides, and
Dr Y. Ando, Mr T. Fujisawa and Ms. Y. Kinoshita for their excellent
technical assistance.
References

1. Chan, M.S. (1997) The global burden of intestinal nematode
infections-fifty years on. Parasitol. Today 13, 438–443.
2. Anderson, T.J., Romero-Abal, M.E. & Jaenike, J. (1993) Genetic
structure and epidemiology of Ascaris populations: patterns of
host affiliation in Guatemala. Parasitology 107, 319–334.
3. Peng, W., Anderson, T.J., Zhou, X. & Kennedy, M.W. (1998)
Genetic variation in sympatric Ascaris populations from humans
andpigsinChina.Parasitology 117, 355–361.
4. Tripathy, K., Duque, E., Bolanos, O., Lotero, H. & Mayoral,
L.G. (1972) Malabsorption syndrome in ascariasis. Am.J.Clin.
Nutr. 25, 1276–1281.
5. Hlaing, T. (1993) Ascariasis and childhood malnutrition. Para-

sitology 107, S125–S136.
6. Cooper,P.J.,Chico,M.,Sandoval,C.,Espinel,I.,Guevara,A.,
Levine, M.M., Griffin, G.E. & Nutman, T.B. (2001) Human
infection with Ascaris lumbricoides is associated with suppression
of the interleukin-2 response to recombinant cholera toxin B
subunit following vaccination with the live oral cholera vaccine
CVD 103-HgR. Infect. Immun. 69, 1574–1580.
7. Paterson, J.C.M., Garside, P., Kennedy, M.W. & Lawrence, C.E.
(2002) Modulation of a heterologous immune response by the
products of Ascaris suum. Infect. Immun. 70, 6058–6067.
8. Kornberg, A. (1962) On the metabolic significance of phos-
phorolytic and pyrophosphorolytic reactions. In Horizons in Bio-

chemistry (Kasha, M. & Pullman, B., eds), pp. 251–264. Academic
Press, Inc., New York, USA.
9. Chen,J.,Brevet,A.,Fromant,M.,Le
´
veˆ que, F., Schmitter, J.M.,
Blanquet, S. & Plateau, P. (1990) Pyrophosphatase is essential for
growth of Escherichia coli. J. Bacteriol. 172, 5686–5689.
10. Lundin, M., Baltscheffsky, H. & Ronne, H. (1991) Yeast PPA2
gene encodes a mitochondrial inorganic pyrophosphatase that is
essential for mitochondrial function. J. Biol. Chem. 266,
12168–12172.
11. Sonnewald, U. (1992) Expression of Escherichia coli inorganic

pyrophosphatase in transgenic plants alters photoassimilate par-
titioning. Plant J. 2, 571–581.
12. Sivula, T., Salminen, A., Parfenyev, A.N., Pohjanjoki, P., Gold-
man, A., Cooperman, B.S., Baykov, A.A. & Lahti, R. (1999)
Evolutionary aspects of inorganic pyrophosphatase. FEBS Lett.
454, 75–80.
13. Shintani, T., Uchiumi, T., Yonezawa, T., Salminen, A., Baykov,
A.A.,Lahti,R.&Hachimori,A.(1998)Cloningandexpression
of a unique inorganic pyrophosphatase from Bacillus subtilis:
evidence for a new family of enzymes. FEBS Lett. 439, 263–266.
14. Young, T.W., Kuhn, N.J., Wadeson, A., Ward, S., Burges, D. &
Cooke, G.D. (1998) Bacillus subtilis ORF YybQ encodes a man-

ganese-dependent inorganic pyrophosphatase with distinctive
properties: the first of a new class of soluble pyrophosphatases?
Microbiology 144, 2563–2571.
15. Rea, P.A., Kim, Y., Sarafian, V., Poole, R.J., Davies, J.M. &
Sanders, D. (1992) Vacuolar H
+
-translocating pyrophosphatase:
a new category of ion translocase. Trends Biochem. Sci. 17, 348–
353.
16. Zhen, R.G., Kim, E.J. & Rea, P.A. (1997) The molecular and
biochemical basis of pyrophosphate-energized proton transloca-
tion at the vacuolar membrane. Adv. Bot. Res. 25, 297–337.

17. Sato, M.H., Kasahara, M., Ishii, N., Homareda, H., Matsui, H. &
Yoshida, M. (1994) Purified vacuolar inorganic pyrophosphatase
consisting of a 75-kDa polypeptide can pump H
+
into reconsti-
tuted proteoliposomes. J. Biol. Chem. 269, 6725–6728.
18. Cooperman, B.S., Baykov, A.A. & Lahti, R. (1992) Evolutionary
conservation of the active site residue of soluble inorganic pyro-
phosphatase. Trends Biochem. Sci. 17, 262–266.
19. Kankare, J., Salminen, T., Lahti, R., Cooperman, B.S., Baykov,
A.A. & Goldman, A. (1996) Crystallographic identification of
metal-binding sites in Escherichia coli inorganic pyrophosphatase.

Biochemistry 35, 4670–4677.
20. Heikinheimo, P., Lehtonen, J., Baykov, A.A., Lahti, R., Coo-
perman, B.S. & Goldman, A. (1996) The structural basis for
pyrophosphatase catalysis. Structure 4, 1491–1508.
21. Terzyan, S.S., Voronova, A.A., Smirnova, E.A., Kuranova, I.P.,
Nekrasov, Y.V., Arutyunyan, E.G., Vainstein, B.K., Hohne, W. &
Hansen, G. (1984) Spatial structure of yeast inorganic pyrophos-
phatase at a resolution of 3 A
˚
. Bioorg. Khim. 10, 1469–1482.
2824 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003
22. Cooperman, B.S. (1982) The mechanism of action of yeast

inorganic pyrophosphatase. Methods Enzymol. 87, 526–548.
23. Yang, Z. & Wensel, T.G. (1992) Inorganic pyrophosphatase from
bovine retinal rod outer segments. J. Biol. Chem. 267, 24634–
24640.
24. Crompton, D.W. (2001) Ascaris and ascariasis. Adv. Parasitol. 48,
285–375.
25. Geenen, P.L., Bresciani, J., Boes, J., Pedersen, A., Eriksen, L.,
Fagerholm, H.P. & Nansen, P. (1999) The morphogenesis of
Ascaris suum to the infective third-stage larvae within the egg.
J. Parasitol. 85, 616–622.
26. Tsuji, N., Kasuga-Aoki, H., Isobe, T. & Yoshihara, S. (2000)
Cloning and characterization of a peroxiredoxin from the swine

roundworm Ascaris suum. Int. J. Parasitol. 30, 125–128.
27. Douvres, F.W. & Urban, J.F. Jr (1987) Nematode except parasites
of insects. In In vitro Methods for Parasite Cultivation (Taylor,
A.E.R. & Baker, J.R., eds), pp. 318–378. Academic Press, Lon-
don, UK.
28. Tsuji, N., Suzuki, K., Kasuga-Aoki, H., Matsumoto, Y., Ara-
kawa, T., Ishiwata, K. & Isobe, T. (2001) Intranasal immunization
with recombinant Ascaris suum 14-kilodalton antigen coupled
with cholera toxin B subunit induces protective immunity to A.
suum infection in mice. Infect. Immun. 69, 7285–7292.
29. Tsuji, N., Morales, T.H., Ozols, V.V., Carmody, A.B. & Chan-
drashekar, R. (1998) Molecular characterization of a calcium-

binding protein from the filarial parasite Dirofilaria immitis. Mol.
Biochem. Parasitol. 97, 69–79.
30. Laemmli, U.K. (1970) Cleavage structural proteins during the
assembly of the head of bacteriophage T4. Nature 227, 680–685.
31. Towbin, H., Staehelin, T. & Gordon, J. (1979) Electrophoretic
transfer of proteins from polyacrylamide gels to nitrocellulose
sheets: procedures and some applications. Proc. Natl Acad. Sci.
USA 76, 4350–4354.
32. O’Farrell, P.Z., Goodman, H.M. & O’Farrell, P.H. (1977) High
resolution two-dimensional electrophoresis of basic as well as
acidic proteins. Cell 12, 1133–1141.
33. Saheki, S., Takeda, A. & Shimazu, T. (1985) Assay of inorganic

phosphate in the mild pH range, suitable for measurement of
glycogen phosphorylase activity. Anal. Biochem. 148, 277–281.
34. Pohjanjoki, P., Lahti, R., Goldman, A. & Cooperman, B.S. (1998)
Evolutionary conservation of enzymatic catalysis: quantitative
comparison of the effects of mutation of aligned residues in
Saccharomyces cerevisiae and Escherichia coli inorganic
pyrophosphatases on enzymatic activity. Biochemistry 37, 1754–
1761.
35. Salminen, T., Ka
¨
pyla
¨

, J., Heikinheimo, P., Kankare, J., Goldman,
A., Heinonen, J., Baykov, A.A., Cooperman, B.S. & Lahti, R.
(1995) Structure and function analysis of Escherichia coli inorganic
pyrophosphatase: is a hydroxide ion the key to catalysis?
Biochemistry 34, 782–791.
36. Yoshida, C., Shah, H. & Weinhouse, S. (1982) Purification and
properties of inorganic pyrophosphatase of rat liver and hepatoma
3924A. Cancer Res. 42, 3526–3531.
37. Hakki, S. & Sitaramayya, A. (1990) Guanylate cyclase from
bovine rod outer segments: solubilization, partial purification, and
regulation by inorganic pyrophosphate. Biochemistry 29,
1088–1094.

38. Moe, O.A. & Butler, L.G. (1972) Yeast inorganic pyrophos-
phatase. II. Kinetics of Mg
2+
activation. J. Biol. Chem. 247,
7308–7314.
39. Baykov, A.A., Hyytia
¨
, T., Volk, S.E., Kasho, V.N., Vener, A.V.,
Goldman, A., Lahti, R. & Cooperman, B.S. (1996) Catalysis by
Escherichia coli inorganic pyrophosphatase: pH and Mg
2+
dependence. Biochemistry 35, 4655–4661.

40. Rodina, E.V., Vainonen, Y.P., Vorobyeva, N.N., Kurilova, S.A.,
Nazarova, T.I. & Avaeva, S.M. (2001) The role of Asp42 in
Escherichia coli inorganic pyrophosphatase functioning. Eur. J.
Biochem. 268, 3851–3857.
41. Davidson, A.M. & Halestrap, A.P. (1989) Inhibition of
mitochondrial-matrix inorganic pyrophosphatase by physiological
[Ca
2+
], and its role in the hormonal regulation of mitochondrial
matrix volume. Biochem. J. 258, 817–821.
42. Rea, P.A. & Poole, R.J. (1993) Vacuolar H
+

-translocating pyro-
phosphatase. Annu. Rev. Plant Physiol. Plant Mol. Biol. 44,
157–180.
43. Scott, D.A., de Souza, W., Benchimol, M., Zhong, L., Lu, H.G.,
Moreno, S.N.J. & Docampo, R. (1998) Presence of a plant-like
proton-pumping pyrophosphatase in acidocalcisomes of Trypa-
nosoma cruzi. J. Biol. Chem. 273, 22151–22158.
44.Rodrigues,C.O.,Scott,D.A.,Bailey,B.N.,DeSouza,W.,
Benchimol, M., Moreno, B., Urbina, J.A., Oldfield, E. & Moreno,
S.N.J. (2000) Vacuolar proton pyrophosphatase activity and
pyrophosphate (PP
i

)inToxoplasma gondii as possible chemo-
therapeutic targets. Biochem. J. 349, 737–745.
45. Marchesini, N., Luo, S., Rodrigues, C.O., Moreno, S.N.J. &
Docampo, R. (2000) Acidocalcisomes and a vacuolar H
+
-pyro-
phosphatase in malaria parasites. Biochem. J. 347, 243–253.
46. Nyre
´
n,P.,Nore,B.F.&Strid,A.(1991)Proton-pumping
N,N¢)dicyclohexylcarbodiimide-sensitive inorganic pyrophos-
phate synthase from Rhodospirillum rubrum: purification, charac-

terization, and reconstitution. Biochemistry 30, 2883–2887.
47. Hill, J.E., Scott, D.A., Luo, S. & Docampo, R. (2000) Cloning and
functional expression of a gene encoding a vacuolar-type proton-
translocating pyrophosphatase from Trypanosoma cruzi. Biochem.
J. 351, 281–288.
48. McIntosh, M.T., Drozdowicz, Y.M., Laroiya, K., Rea, P.A.
& Vaidya, A.B. (2001) Two classes of plant-like vacuolar-type
H
+
-pyrophosphatases in malaria parasites. Mol. Biochem.
Parasitol. 114, 183–195.
49. Nakanishi, Y. & Maeshima, M. (1998) Molecular cloning of

vacuolar H
+
-pyrophosphatase and its developmental expression
in growing hypocotyl of mung bean. Plant Physiol. 116, 589–597.
50. Josse, J. (1966) Constitutive inorganic pyrophosphatase of
Escherichia coli. I. Purification and catalytic properties. J. Biol.
Chem. 241, 1938–1947.
51. Ridlington, J.W., Yang, Y. & Butler, L.G. (1972) Yeast inorganic
pyrophosphatase. IV. Purification, quaternary structure, and evi-
dence for strongly bound Mg
2+
. Arch. Biochem. Biophys. 153,

714–725.
52. Visser, K., Heimovaara-Dijkstra, S., Kijne, J.W. & Wang, M.
(1998) Molecular cloning and characterization of an inorganic
pyrophosphatase from barley. Plant Mol. Biol. 37, 131–140.
53. Parfenyev, A.N., Salminen, A., Halonen, P., Hachimori, A.,
Baykov, A.A. & Lahti, R. (2001) Quaternary structure and metal
ion requirement of Family II pyrophosphatases from Bacillus
subtilis, Streptococcus gordonii,andStreptococcus mutans. J. Biol.
Chem. 276, 24511–24518.
54. Baykov, A.A., Volk, S.E. & Unguryte, A. (1989) Inhibition of
inorganic pyrophosphatase of animal mitochondria by calcium.
Arch. Biochem. Biophys. 273, 287–291.

55. Knight, W.B., Fitts, S.W. & Dunaway-Mariano, D. (1981)
Investigation of the catalytic mechanism of yeast inorganic pyro-
phosphatase. Biochemistry 20, 4079–4086.
56. Rhoads, M.L., Fetterer, R.H. & Urban, J.F. (2001) Release of
hyaluronidase during in vitro development of Ascaris suum from
the third to fourth larval stage. Parasitol. Res. 87, 693–697.
57. Lee, D.L. (2002) Cuticle, moulting and exsheathment. In The
Biology of Nematodes (Lee, D.L., ed.), pp. 171–209. Taylor &
Francis, London, UK.
58. Urbina, J.A., Moreno, B., Vierkotter, S., Oldfield, E., Payares, G.,
Sanoja, C., Bailey, B.N., Yan, W., Scott, D.A., Moreno, S.N.J. &
Docampo, R. (1999) Trypanosoma cruzi contains major

Ó FEBS 2003 Roundworm pyrophosphatase (Eur. J. Biochem. 270) 2825
pyrophosphate stores, and its growth in vitro and in vivo is blocked
by pyrophosphate analogs. J. Biol. Chem. 274, 33609–33615.
59. Sutton, S.V.W., Bender, G.R. & Marquis, R.E. (1987) Fluoride
inhibition of proton-translocating ATPases of oral bacteria. Infect.
Immun. 55, 2597–2603.
60.Meier,B.,Scherk,C.,Schmidt,M.&Parak,F.(1998)
pH-dependent inhibition by azide and fluoride of the iron
superoxide dismutase from Propionibacterium shermanii. Biochem.
J. 331, 403–407.
61. Thongboonkerd, V., Luengpailin, J., Cao, J., Pierce, W.M., Cai,
J., Klein, J.B. & Doyle, R.J. (2002) Fluoride exposure attenuates

expression of Streptococcus pyogenes virulence factors. J. Biol.
Chem. 277, 16599–16605.
2826 M. K. Islam et al. (Eur. J. Biochem. 270) Ó FEBS 2003