PolyphosphatesfromMycobacteriumbovis– potent
inhibitors ofclassIIIadenylate cyclases
Ying Lan Guo
1
, Hermann Mayer
2
, Waldemar Vollmer
3
, Dorothea Dittrich
4
, Peter Sander
4,5
,
Anita Schultz
1
and Joachim E. Schultz
1
1 Pharmazeutisches Institut, Universita
¨
tTu
¨
bingen, Germany
2 Institut fu
¨
r anorganische Chemie, Fakulta
¨
tfu
¨
r Chemie und Pharmazie, Universita
¨
tTu
¨
bingen, Germany
3 Institute for Cell and Molecular Biosciences, Newcastle University, UK
4 Institut fu
¨
r Medizinische Mikrobiologie, Universita
¨
tZu
¨
rich, Switzerland
5 Nationales Zentrum fu
¨
r Mykobakterien, Zurich, Switzerland
cAMP is a key signaling molecule in virtually all living
organisms. This ubiquity is mirrored by the abundance
and diversity of the synthetic enzymes, adenylate cyc-
lases (ACs). Currently, six classes of ACs exist, which
share no identifiable sequence similarities. Here we
deal with ACs grouped together in class III, which
contains by far the most AC isozymes. Among these
are all cyclasesfrom eukaryotes and the overwhelming
majority of those from bacteria [1]. In eukaryotic cells,
ACs are typically pseudoheterodimeric, i.e. the result
of a gene duplication [2]. Both pseudomonomers con-
tribute amino acids to a single catalytic center [3,4].
Bacterial classIII ACs are generally monomers that
must dimerize to form two catalytic centers that are
essentially identical to those of eukaryotic class III
ACs [5,6]. Usually, cAMP is generated intracellularly
in response to extracellular signals such as hormones,
changes in ion compositions, pH or nutrients, and a
variety of stress conditions. Although stimulatory
conditions often persist for considerable periods of
time, cAMP formed in vivo is mostly short-lived.
This requires activated ACs to be quickly returned to
a basal activity state [7–9]. In eukaryotic cells,
GTP hydrolysis and dissociation of the activated
Keywords
adenylate cyclase; cAMP; Mycobacterium;
polyphosphate; stress response
Correspondence
J. E. Schultz, Pharmazeutisches Institut,
Universita
¨
t, Tu
¨
bingen, Auf der Morgenstelle
8, 72076 Tu
¨
bingen, Germany
Fax: +49 7071 295952
Tel: +49 7071 2972475
E-mail: joachim.schultz@uni-tuebingen.de
(Received 7 October 2008, revised 7
November 2008, accepted 10 December
2008)
doi:10.1111/j.1742-4658.2008.06852.x
cAMP generation in bacteria is often stimulated by sudden, but lasting,
changes in extracellular conditions, whereas intracellular cAMP concen-
trations quickly settle at new levels. As bacteria lack G-proteins, it is
unknown how bacterial adenylate cyclase (AC) activities are modulated.
Mycobacterium tuberculosis has 15 classIII AC genes; therefore, we exam-
ined whether mycobacteria contain a factor that may regulate AC activi-
ties. We identified mycobacterial polyphosphates with a mean chain length
of 72 residues as highly potentinhibitorsof dimeric class IIIa, class IIIb
and class IIIc ACs from M. tuberculosis and other bacteria. The identity of
the inhibitor was established by phosphatase degradation,
31
P-NMR, acid
or base hydrolysis, PAGE and comparisons with commercial standards,
and functional substitution by several polyphosphates. The data indicate
that each AC dimer occupies 8–15 phosphate residues on a polyphosphate
strand. Other polyionic polymers such as polyglutamate, polylysine and
hyaluronic acid do not affect cyclase activity. Notably, the structurally
unrelated class I AC Cya from Escherichia coli is unaffected. Bacterial
polyphosphate metabolism is generally viewed in the context of stress-
related regulatory networks. Thus, regulation of bacterial classIII ACs by
polyphosphates could be a component of the bacterial stress response.
Abbreviations
AC, adenylate cyclase; poly-P, polyphosphate.
1094 FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS
AC–G-protein complex terminates signaling, possibly
also with the involvement of secondary modifications
such as phosphorylation [10,11]. Termination of acti-
vation of bacterial classIII ACs has not been investi-
gated. G-proteins or G-protein-like mediators of AC
stimulation are unknown in bacteria. For example, in
the cyanobacterium Anabaena, the ACs CyaB1 and
CyaB2 are stimulated by the enzymatic product cAMP
via an N-terminal tandem GAF domain [12,13].
Theoretically, this would result in perpetual self-activa-
tion until ATP is exhausted, an unlikely physiological
situation. Recently, it has been shown that sodium
ions may be involved in the process of autoinactivation
of CyaB1 AC [14].
In Mycobacterium tuberculosis, 15 classIII AC genes
have been identified [15,16], and for at least 10, AC
activity has been demonstrated [1,17,18]. Deletion of
single AC isoforms in M. tuberculosis did not result in
obvious phenotypes [17,19]. The stressor conditions
employed may have been insufficient and ⁄ or functional
replacement could compensate for the loss of an indi-
vidual AC isoform. Therefore, two major questions
exist: how are the bacterial ACs stimulated, and how
is stimulation terminated while stimulatory conditions
continue to prevail? To address these questions, we
examined whether mycobacteria contain endogenous
factors that modulate AC activities. We used a cell
homogenate fromMycobacteriumbovis BCG, the live
vaccine strain against tuberculosis, to search for the
presence of such modulators. We have isolated poly-
phosphate (poly-P) from M. bovis BCG, which is a
well-known bacterial constituent [20,21], and demon-
strated that it is a most powerful inhibitor ofclass III
ACs from M. tuberculosis and other bacteria. In test-
ing ACs from the class IIIa, IIIb and IIIc subfamilies,
we found that all were strongly inhibited by this cell
constituent, whereas a class I AC from Escherichia coli
was not affected.
Results
In an exploratory experiment, we homogenized 1 g
(wet weight) of M. bovis BCG cells (grown as a settling
culture under hypoxia) with a French press, and
prepared a 100 000 g supernatant (60 min). The super-
natant strongly inhibited the activity of recombinant
Rv1625c, a membrane-bound, mammalian-like AC
from M. tuberculosis [22]. The virtual IC
50
was 1.6 lg
of protein (data not shown). The suspended pellet
from the above centrifugation was inactive. To assess
the specificity of inhibition, several controls were used:
(a) suspended cell pellets and supernatants from Myco-
bacterium smegmatis or E. coli (BL-21) grown without
stress in well-oxygenated rich media had only very low
inhibitory potency; (b) M. bovis BCG cells were
washed extensively with 50 mm Tris ⁄ HCl buffer con-
taining 150 mm NaCl, yet this treatment did not
detach an AC inhibitor; and (c) we tested unused as
well as spent medium after harvesting of BCG. No
inhibition (or activation) was observed. This led us to
believe that M. bovis BCG produced a soluble intra-
cellular inhibitor of Rv1625c.
What is the chemical nature of the inhibitor? Boiling
removed 98% of protein, but AC inhibition was unim-
paired. Similarly, extended digestion with trypsin did
not abolish inhibition, virtually excluding a protein.
The inhibitory factor was not DNA or RNA. This was
verified by nuclease digestion and controlled by agarose
gel electrophoresis. After ether extraction, the inhibitor
remained in the aqueous phase, excluding lipids. Next,
we incubated the enriched inhibitory fraction with lyso-
zyme or cellosyl, a promiscuous bacterial muramidase
from Streptomyces coelicolor. The results were equivo-
cal. After digestion with cellosyl, AC inhibition was
retained, whereas incubation with lysozyme resulted in
loss of inhibition. An HPLC analysis failed to identify
muropeptides, the expected products of lysozyme or
cellosyl digestion (data not shown). Lysozyme has an
isoelectric point of 11. Therefore, we hypothesized that
the inhibitor might be an acidic compound such as
poly-P and bind to lysozyme and not to cellosyl. This
would explain the contradictory results with both
muraminidases. When we incubated a sample with
1.5 units of acid phosphatase at pH 5.5, AC inhibition
was lost. Similarly, inhibition was almost completely
lost upon treatment with HCl or NaOH at room
temperature, indicating hydrolysis of a poly-P.
For final inhibitor purification, the heat-treated sam-
ple was bound to DEAE–Sephacel, released with
400 mm NaCl, and fractionated by Superose 6 chro-
matography. Fractions were assayed for activity using
recombinant Rv1625c (Fig. 1A), and analyzed by elec-
trophoresis [23]. In active fractions, poly-Ps with a
chain length of about 70 residues were stained,
whereas inactive fractions did not stain (Fig. 1B). A
dose–response curve with the combined fractions
established the high inhibitory potency (Fig. 1C). This
indicated that the inhibitor was poly-P.
Final chemical identification was carried out by
31
P-
NMR spectroscopy of concentrated fractions 50–64
from the Superose 6 column (Fig. 2, trace E). The
intense resonance at )21.8 p.p.m. is characteristic for
interior phosphate residues, and the weak, broad peaks
at )6.2 and )20.2 p.p.m. are indicative of terminal
and penultimate phosphate groups, respectively. Thus,
the
31
P-NMR spectrum identified the sample as linear
Y. L. Guo et al. Polyphosphate inhibition ofadenylate cyclases
FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS 1095
poly-Ps [24–26]. This was confirmed by comparisons
with
31
P-NMR spectra of commercial polyphosphate
samples (Fig. 2 and Table 1). The cyclic triphosphate
(trimetaphosphate) showed a singlet at )21.5 p.p.m.,
due to exclusively interior residues (Fig. 2, trace A).
The two terminal groups and the interior residue of
the linear triphosphate gave two resonances at )7.1
and )21.4 p.p.m. at an integrated area ratio of 2 : 1
that were split by each other into a doublet and a
triplet (Fig. 2, trace B). Extending the chain length to
25 phosphate residues allowed detection of the
terminal groups (d = )5.8 p.p.m.), the penultimate
groups (d = )20.8 p.p.m.), and the interior residues
(d = )21.8 p.p.m.). The resonance of the terminal
groups displayed characteristic doublets and doublets
of doublets (Fig. 2, trace C). Further increases in chain
length led to broadening of terminal phosphate reso-
nances, which agreed with a reduced T
2
for polymeric
systems (Fig. 2, trace D, poly-P75). Moreover, calcula-
tion of the peak areas of the resonances allowed an
estimate of the chain length. Calculated chain lengths
for poly-P25 and poly-P75 were in agreement with
those given by the supplier (Table 1). The mycobacte-
rial sample yielded an average chain length of 72, in
agreement with the electrophoretic analysis (Fig. 1B).
Finally, commercial poly-Ps, which were further char-
acterized by
31
P-NMR and hydrolysis by acid or base,
inhibited classIII ACs identically to the material from
M. bovis BCG. With 167 nm Rv1625c in the assay,
200 nm poly-P75 inhibited enzyme activity completely.
Owing to the scarcity of poly-P isolated from
M. bovis BCG, we routinely used biological material
for initial studies and commercial poly-Ps for in-depth
45232
40
A
B
C
45 kDa
30
120
AC activity (%)
46
10
20
80
48
0
D 280 nm
40
020406080100
0
Fraction number
Fraction #
P75 P45 P25 5650 5852 60 6462
Poly P
46 48
100
80
60
20
40
AC activity (%)
0
Mycobacterial poly-P (M)
10
–9
10
–8
Fig. 1. Purification and analysis of an AC inhibitor from M. bovis
BCG. (A) Superose 6 gel filtration and inhibition of Rv1625c by indi-
vidual fractions. Solid line, D 280 nm; stippled lines (d), inhibition
of 167 n
M Rv1625c; arrows on top denote molecular mass mark-
ers. (B) Electrophoretic analysis (15% polyacrylamide gel containing
6
M urea) of inhibitory fractions from (A). Note almost empty
lanes 46 and 48, which are barely inhibitory. Controls of linear poly-
Ps with average chain lengths of 75 (14 lg), 45 (7 lg), and 25
(7 lg) residues are on the left. The gel was stained with toluidine
blue O to detect poly-Ps [23]. The wide bands indicate chain length
variations in the commercial standards and column fractions. One
hundred per cent activity corresponds to 360 nmol cAMPÆmg
)1
Æ
min
)1
. (C) Dose–response curve of Rv1625c inhibition of combined
fractions from the Superose 6 fractionation in (A).
A
B
C
D
E
p.p.m.
Fig. 2.
31
P-NMR spectra of the concentrated fractions (50–64) of
the Superose 6 gel filtration and of commercial poly-P standards:
(A) cyclic poly-P3; (B) linear poly-P3; (C) linear poly-P25; (D) linear
poly-P75; and (E) concentrated inhibitor. PP1, terminal phosphate
groups; PP2, penultimate groups; PPn, interior phosphate residues.
The concentrations of the poly-P standards were calculated to be
uniformly 100 m
M P
i
, i.e. 33.3 mM poly-P3 [(A) and (B)], 4 mM poly-
P25 (C), and 1.33 m
M poly-P75 (D).
Polyphosphate inhibition ofadenylatecyclases Y. L. Guo et al.
1096 FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS
characterizations. We never noted disparities in results
between the different poly-P sources. Rv1625 inhibi-
tion by poly-P75 was instantaneous, as demonstrated
experimentally (Fig. 3A). This established that inhibi-
tion was reversible [27]. This was confirmed in dilution
experiments in which the enzyme–inhibitor complex
was diluted eight-fold immediately prior to the start of
the reaction. Reaction velocities of Rv1625c were
determined in the presence of 10, 20 and 25 nm poly-
P75 as a function of the substrate Mn-ATP. A Linewe-
aver–Burk plot showed that K
m
values were unchanged
whereas V
max
decreased, indicating noncompetitive
inhibition (Fig. 3B). We excluded any chelating effect
of poly-Ps on the Mn
2+
concentration, because even if
the poly-P concentration was calculated in terms of
molarity of orthophosphate, it did not exceed low
micromolar values, whereas Mn
2+
was fixed at 2 mm;
that is, poly-P could not act as a chelating agent. Next,
we investigated whether phosphates of different chain
length had differing inhibitory potencies. Orthophos-
phate up to 11 mm had no effect on Rv1625c, pyro-
phosphate inhibited it with an IC
50
of 210 lm, the
linear triphosphate had an IC
50
of 20 lm, and the cyc-
lic trimetaphosphate an IC
50
of 2 mm; that is, these
compounds were poor inhibitors (Table 2). In contrast,
all IC
50
concentrations of poly-Ps with a chain length
of 16 or more were in the submicromolar range (Fig. 4
and Table 2). When we normalized the individual IC
50
concentrations of various poly-P compounds with
chain lengths > 16 to the concentration of orthophos-
phate, i.e. the poly-P IC
50
concentrations in Table 2
were multiplied by the average phosphate chain length,
we obtained a mean IC
50
concentration of
660 ± 62.8 nm phosphate (± SEM; range 561–
825 nm). This indicated that once a critical length of
the poly-P chain is reached, the increasing affinity
of poly-P is linearly related to the increase in the cal-
culated total phosphate concentration. For poly-P75
and cyclic poly-P17, the apparent K
i
values were
determined with 100 nm Rv1625c, 30 and 60 lm
Mn-ATP, and different inhibitor concentrations
Table 1. Chemical shifts and coupling constants of the purified inhibitor and the commercial poly-Ps. d, chemical shift;
2
J
pp
, coupling
constant of two adjacent phosphate groups; PP1, terminal phosphate groups; PP2, penultimate phosphate groups; PPn, interior phosphate
residues; s, singlet; d, doublet; t, triplet; br, broad; br s, broad singlet; br d, broad doublet.
Sample
d (p.p.m.)
Calculated chain
length (average)
PP1 PP2 PPn
2
J
PP
(Hz)
P3 cyclic )21.5 (s) Not applicable
P3 linear )7.1 (d) )21.4 (t) 20.77 Not applicable
P25 linear )5.8 (br d) )20.8 )21.8 (br s) 18.5; 17.4 27
P75 linear )5.7 (br) )20.9 (br) )21.9 (br s) 79
Inhibitor )6.2 (br) )20.2 (br) )21.8 (br s) 72
100
A
B
80
25 nM
40
60
40 nM
50 nM
20
AC activity (%)
60 nM
80 nM
0 40 80 120 160
0
7
Time of preincubation (s)
5
25 n
M
4
10
20
3
1/v (µmol·min*mg)
–1
0
2
1
0 0.02
1
–0.04 –0.02 0.04 0.06 0.08
1/S (µM)
–1
Fig. 3. Kinetics of the inhibition of Rv1625c by poly-P75. (A)
Rv1625c at 134 n
M was used in the assay. The concentrations of
poly-P75 added at the beginning are indicated. The first assay was
started after 7 s of preincubation of protein and poly-P75 by addition
of the substrate ATP. Assays were run for 4 min. One hundred per-
cent activity corresponds to 448 nmol cAMPÆmg
)1
Æmin
)1
. (B) Double
reciprocal plot (Lineweaver–Burk) of substrate kinetics of Rv1625c
in the presence of three concentrations of poly-P75 as an inhibitor.
Y. L. Guo et al. Polyphosphate inhibition ofadenylate cyclases
FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS 1097
(5–80 nm). Dixon diagrams gave apparent K
i
values of
14 nm for poly-P75 and 20 nm for poly-P17, i.e. lower
than the enzyme concentration. This indicated that a
single poly-P strand inhibited more than one AC mole-
cule. Next, we determined IC
50
concentrations of poly-
P25 at different concentrations of Rv1625c. Here, IC
50
values increased linearly with the increasing protein
concentration (Fig. 5). With a slope of the curve of
0.154 (R = 0.982; Fig. 5), and considering that at
IC
50
, 50% of the dimers are bound to poly-P, this indi-
cated a molar ratio between the Rv1625c dimer and
poly-P25 of 3 : 1, close to the results obtained with
poly-Ps with different chain lengths (Table 2).
Next, we examined whether poly-Ps inhibit different
bacterial AC isoforms. On the basis of systematic
differences in key amino acids and on small, strictly
localized length variations, classIII ACs have been
divided into four subfamilies, IIIa to IIId [1]. Rv1625c
is a class IIIa AC, as are all mammalian membrane-
bound ACs. A concatenated Rv1625c homodimer with
an identical domain sequence as the membranous
mammalian ACs, (Rv1625c)
2
, is active [17] and inhib-
ited by poly-P, just like Rv1625c (data not shown). Do
poly-Ps also inhibit AC isozymes from other class III
subfamilies? We examined the following ACs: cyaG
from Arthrospira platensis as another class IIIa isoform
containing a HAMP domain; as class IIIb ACs, myco-
bacterial Rv3645, which has a HAMP domain between
its membrane anchor and the catalytic domain, and
CyaB1 from Anabaena sp., which has an N-terminal
cAMP-binding tandem GAF domain; as a class IIIc
AC, the M. tuberculosis pH sensor Rv1264 [6]. In
general, all classIII ACs were potently inhibited by
poly-P75 (Fig. 6A). Although the IC
50
concentrations
differed slightly (11, 57, 315, 102 and 31 nm for
Rv1625c, CyaG, Rv3645, CyaB1 and Rv1264, respec-
tively), all were in the nanomolar range (Fig. 6A).
Because the catalytic centers ofclassIII ACs appear
to be highly similar [3,5,6], it is likely that poly-Ps gen-
erally inhibit classIII ACs. Rv1264 has been shown to
be a pH sensor that is strongly activated by pH values
around 5.5 [6]. This allowed testing of whether poly-Ps
affected the basal and the activated states of a clas-
s IIIc AC similarly. Poly-P75 inhibited the basal state
at pH 8 and the activated state at pH 5.5, but with
markedly different efficacies. The activated enzyme
was fully inhibited at 400 nm, whereas the basal state
was not yet fully inhibited at 30 lm (Fig. 6B). The
Table 2. IC
50
values of poly-Ps for the AC Rv1625c. The protein
concentration in the assays was 83 n
M.
Phosphates IC
50
values (nM)
Phosphate (P
i
)>11· 10
6
PP
i
21 · 10
4
Linear PPP
i
20 · 10
3
Cyclic PPP
i
20 · 10
5
Poly-P16 100
Cyclic poly-P17 33
Poly-P25 27
Poly-P30 18
Poly-P45 12
Poly-P68 12
Poly-P75 11
120
PPPi
Poly-P16
80
100
Poly-P30
Poly-P75
60
AC activity (%)
20
40
–9
0
–8 –7 –6 –5 –4 –3
10
Concentration (M)
10 10 10 10 10 10
Fig. 4. Inhibition of Rv1625c by poly-Ps of different strand lengths.
Rv1625c at 167 n
M was used in the assays. One hundred per cent
activity corresponds to 370 nmol cAMPÆmg
)1
min
)1
.
26
22
18
10
14
IC
50
(nM Poly-P25)
25 50 75 100 125 150
Rv1625c dimer concentration (nM)
Fig. 5. Poly-P25 IC
50
values increase with increasing AC concentra-
tions. Assays were carried out at different Rv1625c concentrations.
IC
50
values (y-axis) were plotted against the protein concentrations
at which only dimers exist [22] (regression coefficient r = 0.999).
Polyphosphate inhibition ofadenylatecyclases Y. L. Guo et al.
1098 FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS
reported structures of Rv1264 showed that a canonical
catalytic cleft ofclassIII ACs is formed at pH 5.5,
whereas at pH 8, the catalytic amino acids are far
apart [6]. The differences in the IC
50
concentrations
may indicate that poly-P binds in the catalytic crevice.
Another question was whether poly-P specifically
inhibits classIII ACs or also acts on class I ACs. We
expressed Cya, the class I AC from E. coli, and purified
it to homogeneity by Ni
2+
–nitrilotriacetic acid chro-
matography. The specific activity of the purified protein
was 27.2 nmolÆmg
)1
Æmin
)1
with Mg-ATP as a sub-
strate. This class I AC was unaffected by up to 30 lm
poly-P75 in the assay (Fig. 6A, stippled line). Finally,
the specificity of poly-P inhibition was investigated
using other ionic polymers. We employed anionic poly-
glutamate (M
r
‡ 15 000, Sigma, Munich, Germany),
cationic polylysine (M
r
4000–15 000, Sigma), and acidic
hyaluronic acid (from Streptococcus equi, Fluka,
Munich, Germany). We tested these compounds at 0.2
and 2 lm with Rv1625c. None inhibited AC activity,
demonstrating that the effect of poly-P was specific.
Discussion
cAMP in bacteria is discussed in numerous publica-
tions as a second messenger involved in regulatory
pathways. However, reliable studies in which intracel-
lular cAMP concentrations were determined and regu-
lation of cAMP biosynthesis in vivo was examined are
rare. This is due to its low intracellular concentrations
and secretion of up to 95% of total cAMP into the
medium, which causes unusual experimental difficulties
and ambiguities [28–35]. Generally, conditions that
stimulate cAMP formation in bacteria, e.g. changes in
pH, ion or nutrient concentrations, are related to
stress conditions [28,36]. In this study, we could not
remedy the lack of knowledge of bacterial cAMP
metabolism, and the physiological relevance of bacte-
rial classIII AC inhibition by poly-P merits further
experiments.
At the outset, we asked whether M. bovis BCG con-
tains endogenous factors that regulate AC activities.
The isolated AC inhibitor was unequivocally identified
by chemical means (
31
P-NMR, acid and base hydro-
lysis, SDS ⁄ PAGE), biochemical means (phosphatase
degradation), and full functional substitution by com-
mercial poly-P. In addition, other bacterial constitu-
ents were excluded, such as DNA, RNA, proteins and
peptides, and peptidoglycans of the cell wall. Although
poly-P is a metabolic staple that is present in cells
from all the kingdoms of life – bacteria, fungi, plants
and animals – it has never been studied in conjunction
with regulation of ACs [20,37]. Mycobacteria produce
poly-Ps under a variety of stress conditions, and pos-
sess two poly-P kinases [21,38]. Bacterial poly-Ps vary
in size and solubility, and intracellular concentrations
of poly-P fluctuate considerably (2–15 ngÆmg
)1
protein)
[20,37,39,40]. Depending on the organism, growth and
100
A
B
80
40
60
AC activity (%)
20
0
–8 –7 –6 –5
Poly-P75 (M)
10 10 10 10
100
pH 8.0
80
pH 5.5
40
60
20
AC activity (%)
–8
0
–7 –6 –5
10
Poly-P75 (
M)
10 10 10
Fig. 6. Inhibition of bacterial ACs fromclassIII and class I by
poly-P75. (A) The following amounts of protein were used:
Rv1264(1–397), 660 n
M (d); CyaG(370–672), an N-terminal truncated
version with only HAMP and catalytic domains, 640 n
M (s);
CyaB1(1–859), 22 n
M ( ); Rv3645(1–549), 153 lg of total membrane
proteins (h); Cya(1–446) from E. coli, 876 n
M ( , stippled line). (B)
Inhibition of basal and activated states of 660 n
M Rv1264 by poly-
P75. The protein concentrations used were based on the linear sec-
tions of the protein dependency of the respective AC reactions. This
ensured that dimerization of respective monomers was complete.
Y. L. Guo et al. Polyphosphate inhibition ofadenylate cyclases
FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS 1099
physiological conditions, poly-P may amount to up to
20% of bacterial dry weight [40,41].
For poly-P in bacteria, several physiological
functions in many locations have been proposed, e.g.
poly-P accumulation in stress sensing [37,39,42–46].
Furthermore, poly-P is discussed as a primordial pre-
cursor of ATP, a flexible scaffold for the assembly of
macromolecules, a cellular phosphate store, a buffer
system in pH regulation, being involved in chelation of
cations such as Ca
2+
,Mn
2+
and Mg
2+
[20,37,39,47].
Our data suggest a new potential role for poly-P as an
inhibitor of bacterial classIII ACs. Because cAMP as
a bacterial alarmone is a global signaling molecule, the
reported actions of poly-P on bacterial classIII ACs
may affect several cellular functions simultaneously. It
was most interesting to note that the class I AC from
E. coli was not inhibited by poly-P. This may indicate
that the sequence dissimilarities between different clas-
ses of ACs reflect functional differences. The presence
in some bacteria of ACs from different classes would
then enable different modes and levels of regulations.
Actually, it may be useful for certain bacteria to con-
tain AC isoforms from different classes (such as Pseu-
domonas aeroginosa, which has AC isoforms from
classes I, II and III), because this would broaden the
modes of cellular regulation and pathogenicity [48].
Because the catalytic folds of a bacterial class IIIc
and a mammalian class IIIa AC are superimposable
[3,6], it is reasonable to assume that mammalian ACs
will be inhibited by poly-P as well. In preliminary
experiments using membranes prepared from a rat
brain homogenate, we found that brain AC activity
was inhibited by poly-P with an IC
50
of 10 lm (data
not shown). As far as the mechanism of inhibition is
concerned, the data obtained with Rv1264 indicate
binding in the catalytic fold. Possibly, poly-P bridges
and occludes the substrate-determining lysine (Lys296
in Rv1625c) and the arginine (Arg376), which stabilizes
the transition state. Therefore, poly-P may be helpful
in attempts to crystallize and characterize other bacte-
rial ACs.
To the best of our knowledge, poly-P metabolism
has never been studied in conjunction with cAMP
metabolism. The concentration of poly-P in stressed
bacteria appears to be higher than is needed for AC
inhibition. Under the hypoxic growth conditions of
M. bovis BCG used in this study, the concentration of
poly-P would probably silence all classIII ACs.
Several possibilities exist to explain this fact. One is
that at a low level of stress conditions, such as modest
oxygen deprivation or nutrient depletion, cAMP
formation is elicited as an initial response. Poly-P bio-
synthesis is then initiated, and this turns off cAMP
production ofclassIII ACs. Another possibility is that
the availability of poly-P is locally restricted. The neg-
atively charged polyanionic compound may bind to
positively charged carrier molecules or it may be neu-
tralized by inorganic cations. The availability of poly-P
for termination ofclassIII activation would then be
tied to competition for different intracellular binding
sites. Finally, the possibility exists that poly-P accumu-
lation is controlled locally, such that ACs are partially
maintained in an inhibited, inactive state. Release from
inhibition could occur by poly-P degradation by tightly
regulated phosphatases. The latter would thus attain
regulatory significance.
Experimental procedures
Mycobacterial strain and growth
M. bovis BCG 1721, a streptomycin-resistant derivative of
BCG Pasteur, carrying a non-restrictive rpsL mutation
(K42R) [49], was grown as a settling culture in tissue culture
flasks in Middlebrook 7H9 medium supplemented with oleic
acid, albumin, dextrose, catalase (Difco, Heidelberg,
Germany) and Tween-80 (0.05%) [50]. Flasks were shaken
once daily by hand; that is, cells were grown under hypoxic
stress. E. coli was grown in LB medium, and M. smegmatis
in LB + 0.05% Tween-80 under constant shaking for oxy-
genation (210 r.p.m. shaking speed). Bacteria were harvested
at an attenuance (D
600 nm
) of 0.5–0.7 by centrifugation
(10 min, 4400 g at 4 °C) and stored at )80 °C until use.
Expression and purification of Rv1625c
Rv1625c(1–443) in pQE60 was expressed in E. coli BL-
21(DE3)(pRep4) and purified to homogeneity using 0.6%
n-dodecyl-b-d-maltoside for solubilization as previously
described [17]. The purified protein could be stored at
)80 °C without loss of activity for at least 6 months. The
catalytic domain Rv1625c(204–443) and other bacterial
ACs used were expressed and purified to homogeneity as
previously reported [12,17,22,51,52].
AC assay
AC activity was measured for 10 min in a volume of
100 lLat30°C [53]. The reactions contained 22% glycerol,
50 mm Tris ⁄ HCl (pH 7.5), 2 mm MnCl
2
or 2 mm MgCl
2
,
the indicated concentrations of ATP with 25 kBq of
[
32
P]ATP[aP] and 2 mm cAMP with 150 Bq of
[2,8-
3
H]cAMP to monitor yield during product isolation.
For determination of kinetic constants, ATP was varied
from 14 lm to 100 lm, with constant 2 mm Mn
2+
. The
reaction was started by addition of enzyme. ATP conver-
sion was limited to < 10%, to guarantee linearity. ATP
Polyphosphate inhibition ofadenylatecyclases Y. L. Guo et al.
1100 FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS
was separated from product cAMP by sequential chroma-
tography [53].
Purification and analysis of poly-Ps from
M. bovis BCG
Cells were suspended in 50 mm Tris ⁄ HCl (pH 7.5) and
broken with a French press. The homogenate was centri-
fuged (100 000 g for 1 h), and the supernatant was heated
at 95 °C for 30 min. Coagulated protein was removed
(100 000 g for 1 h). The supernatant was centrifuged
through Sephadex G50 spin columns to remove small con-
taminants; the inhibitory capacity was in the eluate. Next,
DEAE–Sephacel was added, and the material bound to
the anion exchanger. The matrix was poured into a col-
umn, washed, and eluted with 400 mm NaCl. The eluate
was applied to a Superose 6 column (30 · 1 cm), and frac-
tions were examined by electrophoresis and for inhibitory
activity (Fig. 1). Electrophoresis of poly-Ps was carried out
as previously described [23]. A 15% acrylamide ⁄ bisacryla-
mide gel with 6 m urea was prepared with TBE buffer
(89 mm Tris ⁄ borate, 2 mm EDTA, pH 8.3). The gel was
prerun at 200 V for 60 min. Poly-Ps with average chain
lengths of 25, 45 and 75 residues were used as markers
(Sigma). Probes that contained 25% of sample buffer
(50% sucrose, 0.125% bromophenol blue and 450 mm
Tris ⁄ borate at pH 8.3, 13.5 mm EDTA) were loaded and
electrophoresed for 25 min. Gels were stained with 0.05%
toluidine blue O in 25% methanol and 5% glycerol for
20 min. Destaining was performed with the same solvent,
lacking the dye. The concentration of poly-P in mycobac-
terial preparations were assessed using standard curves
with poly-P75.
31
P-NMR measurement
The
31
P-NMR spectra were obtained on Bruker
Avance 400 and Bruker Avance 500 spectrometers operat-
ing at 161.98 and 202.45 MHz, respectively. The spectra
were recorded by applying 30° pulses with a repetition time
of 1 s at 21 °C, and referenced against external 85%
H
3
PO
4
. Multipuls decoupling sequences were applied to
remove any proton phosphorus interactions.
31
P peak areas
were obtained by fitting the spectra to a set of Lorenzian
line shapes using the Bruker topspin 2.0 software package.
The pH of all samples was adjusted to 7.5.
The fractions of Superose 6 that showed 95% inhibitory
activity against Rv1625c in 50 mm Tris ⁄ HCl and 100 mm
NaCl were concentrated with an Amicon Ultra-4 5K cen-
trifugal filter device, and subsequently diluted with D
2
Oto
reduce noise (50% final D
2
O in the water). A total volume
of 0.5 mL was transferred into a Norell 507-HP sample
tube; 50 mm linear poly-P3, poly-P25 and poly-P75, and
cyclic poly-P3 with sodium cations in 50% D
2
O, were used
as standards.
Muropeptide determination by HPLC
The concentrated Superose 6 fractions (see above) were
analyzed for the presence of muropeptides according to
Glauner [54]. Briefly, the sample was incubated with cello-
syl or lysozyme at pH 4.8 at 37 °C overnight, deactivated
(100 °C, 10 min), and centrifuged (14 000 g, 8 min). Eighty
microliters of the supernatant was mixed with 80 lLof
sodium borate (0.5 m, pH 9.0) and 1–2 mg of sodium boro-
hydride. The excess of borohydride was destroyed after
incubation at room temperature for 30 min. Separation of
muropeptides occurred on a 250 · 4.6 mm 3 lm Hypersil
ODS column at 55 °C using a 135 min gradient from
buffer A (50 mm sodium phosphate, pH 4.3) to buffer B
(75 mm sodium phosphate, pH 4.9, 15% methanol) at a
flow rate of 0.5 mLÆmin
)1
. Detection of muropeptides
occurred at 205 nm. We confirmed that cellosyl degraded
mycobacterial cell wall material in respective controls with
mycobacterial homogenates.
Acknowledgements
This work was supported by the Deutsche Forschungs-
gemeinschaft (SFB 766). P. Sander is in part supported
by the Swiss National Science Foundation (contract:
3100A0_120326) and the European Union (LSHP-CT-
2006-037217).
References
1 Linder JU & Schultz JE (2003) The classIII adenylyl
cyclases: multi-purpose signalling modules. Cell Signal
15, 1081–1089.
2 Krupinski J, Coussen F, Bakalyar HA, Tang WJ, Fein-
stein PG, Orth K, Slaughter C, Reed RR & Gilman
AG (1989) Adenylyl cyclase amino acid sequence: possi-
ble channel- or transporter-like structure. Science 244,
1558–1564.
3 Tesmer JJ, Sunahara RK, Gilman AG & Sprang SR
(1997) Crystal structure of the catalytic domains of ade-
nylyl cyclase in a complex with Gsalpha.GTPgammaS.
Science 278, 1907–1916.
4 Tesmer JJ, Sunahara RK, Johnson RA, Gosselin G,
Gilman AG & Sprang SR (1999) Two-metal-ion
catalysis in adenylyl cyclase. Science 285, 756–
760.
5 Sinha SC, Wetterer M, Sprang SR, Schultz JE & Linder
JU (2005) Origin of asymmetry in adenylyl cyclases:
structures ofMycobacterium tuberculosis Rv1900c.
EMBO J 24, 663–673.
6 Tews I, Findeisen F, Sinning I, Schultz A, Schultz JE &
Linder JU (2005) The structure of a pH-sensing myco-
bacterial adenylyl cyclase holoenzyme. Science 308,
1020–1023.
Y. L. Guo et al. Polyphosphate inhibition ofadenylate cyclases
FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS 1101
7 Schultz J & Daly JW (1973) Cyclic adenosine 3¢,5¢-
monophosphate in guinea pig cerebral cortical slices. 3.
Formation, degradation, and reformation of cyclic
adenosine 3¢,5¢-monophosphate during sequential stimu-
lations by biogenic amines and adenosine. J Biol Chem
248, 860–866.
8 Imashimizu M, Yoshimura H, Katoh H, Ehira S &
Ohmori M (2005) NaCl enhances cellular cAMP and
upregulates genes related to heterocyst development in
the cyanobacterium, Anabaena sp. strain PCC 7120.
FEMS Microbiol Lett 252, 97–103.
9 Schultz JE, Klumpp S, Benz R, Schurhoff-Goeters WJ
& Schmid A (1992) Regulation of adenylyl cyclase from
Paramecium by an intrinsic potassium conductance.
Science 255, 600–603.
10 Romero-Avila MT, Flores-Jasso CF & Garcia-Sainz JA
(2002) alpha1B-Adrenergic receptor phosphorylation
and desensitization induced by transforming growth
factor-beta. Biochem J 368, 581–587.
11 Rondard P, Iiri T, Srinivasan S, Meng E, Fujita T &
Bourne HR (2001) Mutant G protein alpha subunit
activated by Gbeta gamma: a model for receptor activa-
tion? Proc Natl Acad Sci USA 98, 6150–6155.
12 Kanacher T, Schultz A, Linder JU & Schultz JE (2002)
A GAF-domain-regulated adenylyl cyclase from Anaba-
ena is a self-activating cAMP switch. EMBO J 21,
3672–3680.
13 Bruder S, Linder JU, Martinez SE, Zheng N, Beavo JA
& Schultz JE (2005) The cyanobacterial tandem GAF
domains from the cyaB2 adenylyl cyclase signal via
both cAMP-binding sites. Proc Natl Acad Sci USA 102,
3088–3092.
14 Cann MJ (2007) Sodium regulation of GAF domain
function. Biochem Soc Trans 35, 1032–1034.
15 Cole ST, Brosch R, Parkhill J, Garnier T, Churcher C,
Harris D, Gordon SV, Eiglmeier K, Gas S, Barry CE
III et al. (1998) Deciphering the biology of Mycobacte-
rium tuberculosis from the complete genome sequence.
Nature 393, 537–544.
16 McCue LA, McDonough KA & Lawrence CE (2000)
Functional classification of cNMP-binding proteins and
nucleotide cyclases with implications for novel regula-
tory pathways in Mycobacterium tuberculosis. Genome
Res 10, 204–219.
17 Guo YL, Kurz U, Schultz A, Linder JU, Dittrich D,
Keller C, Ehlers S, Sander P & Schultz JE (2005) Inter-
action of Rv1625c, a mycobacterial class IIIa adenylyl
cyclase, with a mammalian congener. Mol Microbiol 57,
667–677.
18 Shenoy AR & Visweswariah SS (2006) New messages
from old messengers: cAMP and mycobacteria. Trends
Microbiol 14, 543–550.
19 Dittrich D, Keller C, Ehlers S, Schultz JE & Sander P
(2006) Characterization of a Mycobacterium tuberculosis
mutant deficient in pH-sensing adenylate cyclase
Rv1264. Int J Med Microbiol 296, 563–566.
20 Kornberg A (1995) Inorganic polyphosphate: toward
making a forgotten polymer unforgettable. J Bacteriol
177
, 491–496.
21 Winder FG & Denneny JM (1957) The metabolism of
inorganic polyphosphate in mycobacteria. J Gen Micro-
biol 17, 573–585.
22 Guo YL, Seebacher T, Kurz U, Linder JU & Schultz
JE (2001) Adenylyl cyclase Rv1625c of Mycobacterium
tuberculosis: a progenitor of mammalian adenylyl
cyclases. EMBO J 20, 3667–3675.
23 Robinson NA & Wood HG (1986) Polyphosphate kinase
from Propionibacterium shermanii. Demonstration that
the synthesis and utilization of polyphosphate is by a
processive mechanism. J Biol Chem 261, 4481–4485.
24 MacDonald JC & Mazurek M (1987) Phosphorus
magnetic resonance spectra of open-chain linear poly-
phosphates. J Magn Reson 72, 48–60.
25 Bental M, Pick U, Avron M & Degani H (1991) Poly-
phosphate metabolism in the alga Dunaliella salina stud-
ied by 31P-NMR. Biochim Biophys Acta 1092, 21–28.
26 Castrol CD, Koretsky AP & Domach MM (1999)
NMR-observed phosphate trafficking and polyphos-
phate dynamics in wild-type and vph1-1 mutant Saccha-
romyces cerevisae in response to stresses. Biotechnol
Prog 15, 65–73.
27 Bisswanger H (2002) Enzyme Kinetics Principles and
Methods, 3rd edn. Wiley-VCH, Weinheim.
28 Makman RS & Sutherland EW (1965) Adenosine 3¢,5¢-
phosphate in Escherichia coli. J Biol Chem 240, 1309–
1314.
29 Lee CH (1977) Identification of adenosine 3¢,5¢-mono-
phosphate in Mycobacterium smegmatis. J Bacteriol
132, 1031–1033.
30 Padh H & Venkitasubramanian TA (1977) Adenosine
3¢,5¢-monophosphate in mycobacteria. Life Sci 20,
1273–1280.
31 Padh H & Venkitasubramanian TA (1976) Adenosine
3¢,5¢-monophosphate in Mycobacterium phlei and Myco-
bacterium tuberculosis H37Ra. Microbios 16, 183–189.
32 Pastan I & Perlman R (1970) Cyclic adenosine mono-
phosphate in bacteria. Science 169
, 339–344.
33 Peterkofsky A & Gazdar C (1971) Glucose and the
metabolism of adenosine 3¢:5¢-cyclic monophosphate in
Escherichia coli . Proc Natl Acad Sci USA 68, 2794–
2798.
34 Peterkofsky A & Gazdar C (1973) Measurements of
rates of adenosine 3 ¢:5 ¢-cyclic monophosphate synthesis
in intact Escherichia coli B. Proc Natl Acad Sci USA
70, 2149–2152.
35 Bettenbrock K, Sauter T, Jahreis K, Kremling A, Leng-
eler JW & Gilles ED (2007) Correlation between growth
rates, EIIACrr phosphorylation, and intracellular cyclic
Polyphosphate inhibition ofadenylatecyclases Y. L. Guo et al.
1102 FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS
AMP levels in Escherichia coli K-12. J Bacteriol 189,
6891–6900.
36 Maeda M, Lu S, Shaulsky G, Miyazaki Y, Kuwayama
H, Tanaka Y, Kuspa A & Loomis WF (2004) Periodic
signaling controlled by an oscillatory circuit that
includes protein kinases ERK2 and PKA. Science 304,
875–878.
37 Brown MR & Kornberg A (2004) Inorganic polyphos-
phate in the origin and survival of species. Proc Natl
Acad Sci USA 101, 16085–16087.
38 Suzuki H, Kaneko T & Ikeda Y (1972) Properties of
polyphosphate kinase prepared from mycobacterium
smegmatis. Biochim Biophys Acta 268, 381–390.
39 Kornberg A, Rao NN & Ault-Riche D (1999) Inorganic
polyphosphate: a molecule of many functions. Annu
Rev Biochem 68, 89–125.
40 Rao NN & Kornberg A (1996) Inorganic polyphos-
phate supports resistance and survival of stationary-
phase Escherichia coli. J Bacteriol 178, 1394–1400.
41 Rao NN, Roberts MF & Torriani A (1985) Amount
and chain length ofpolyphosphates in Escherichia coli
depend on cell growth conditions. J Bacteriol 162, 242–
247.
42 Sureka K, Dey S, Datta P, Singh AK, Dasgupta A,
Rodrigue S, Basu J & Kundu M (2007) Polyphosphate
kinase is involved in stress-induced mprAB-sigE-rel
signalling in mycobacteria. Mol Microbiol 65, 261–
276.
43 Reusch RN, Huang R & Bramble LL (1995) Poly-3-hy-
droxybutyrate ⁄ polyphosphate complexes form voltage-
activated Ca2+ channels in the plasma membranes of
Escherichia coli. Biophys J 69, 754–766.
44 Kim KS, Rao NN, Fraley CD & Kornberg A (2002)
Inorganic polyphosphate is essential for long-term
survival and virulence factors in Shigella and Salmonella
spp. Proc Natl Acad Sci USA 99, 7675–7680.
45 Jahid IK, Silva AJ & Benitez JA (2006) Polyphosphate
stores enhance the ability of Vibrio cholerae to
overcome environmental stresses in a low-phosphate
environment. Appl Environ Microbiol 72, 7043–7049.
46 Fraley CD, Rashid MH, Lee SS, Gottschalk R, Harri-
son J, Wood PJ, Brown MR & Kornberg A (2007) A
polyphosphate kinase 1 (ppk1) mutant of Pseudomonas
aeruginosa exhibits multiple ultrastructural and func-
tional defects. Proc Natl Acad Sci USA 104, 3526–3531.
47 Tanaka S, Lee SO, Hamaoka K, Kato J, Takiguchi N,
Nakamura K, Ohtake H & Kuroda A (2003) Strictly
polyphosphate-dependent glucokinase in a polyphos-
phate-accumulating bacterium, Microlunatus phosphovo-
rus. J Bacteriol 185, 5654–5656.
48 Smith RS, Wolfgang MC & Lory S (2004) An adenylate
cyclase-controlled signaling network regulates Pseudo-
monas aeruginosa virulence in a mouse model of acute
pneumonia. Infect Immun 72, 1677–1684.
49 Sander P, Meier A & Bottger EC (1995) rpsL+: a dom-
inant selectable marker for gene replacement in myco-
bacteria. Mol Microbiol 16, 991–1000.
50 Sander P, Rezwan M, Walker B, Rampini SK, Krop-
penstedt RM, Ehlers S, Keller C, Keeble JR, Hagemeier
M, Colston MJ et al. (2004) Lipoprotein processing is
required for virulence ofMycobacterium tuberculosis.
Mol Microbiol 52, 1543–1552.
51 Linder JU, Schultz A & Schultz JE (2002) Adenylyl
cyclase Rv1264 fromMycobacterium tuberculosis has an
autoinhibitory N-terminal domain. J Biol Chem 277,
15271–15276.
52 Linder JU, Hammer A & Schultz JE (2004) The effect
of HAMP domains on class IIIb adenylyl cyclases from
Mycobacterium tuberculosis . Eur J Biochem 271,
2446–2451.
53 Salomon Y, Londos C & Rodbell M (1974) A highly
sensitive adenylate cyclase assay. Anal Biochem 58,
541–548.
54 Glauner B (1988) Separation and quantification of
muropeptides with high-performance liquid chromatog-
raphy. Anal Biochem 172, 451–464.
Y. L. Guo et al. Polyphosphate inhibition ofadenylate cyclases
FEBS Journal 276 (2009) 1094–1103 ª 2009 The Authors Journal compilation ª 2009 FEBS 1103
. polyphosphates with a mean chain length of 72 residues as highly potent inhibitors of dimeric class IIIa, class IIIb and class IIIc ACs from M. tuberculosis and other bacteria. The identity of the. Polyphosphates from Mycobacterium bovis – potent inhibitors of class III adenylate cyclases Ying Lan Guo 1 , Hermann Mayer 2 , Waldemar Vollmer 3 ,. activity (%) –8 0 –7 –6 –5 10 Poly-P75 ( M) 10 10 10 Fig. 6. Inhibition of bacterial ACs from class III and class I by poly-P75. (A) The following amounts of protein were used: Rv1264( 1–3 97), 660