Department of Biochemistry, Stanford
University School of Medicine, Stanford, California 94305-5307
A major impediment to understanding the biological roles of
inorganic polyphosphate (polyP) has been the lack of sensitive definitive methods to extract and quantitate cellular polyP. We show
that polyP recovered in extracts from cells lysed with guanidinium isothiocynate can be bound to silicate glass and quantitatively measured by a two-enzyme assay: polyP is first converted to ATP by
polyP kinase, and the ATP is hydrolyzed by luciferase to generate light. This nonradioactive method can detect picomolar amounts of
phosphate residues in polyP per milligram of extracted protein. A
simplified procedure for preparing polyP synthesized by polyP kinase is
also described. Using the new assay, we found that bacteria subjected
to nutritional or osmotic stress in a rich medium or to nitrogen
exhaustion had large and dynamic accumulations of polyP. By contrast,
carbon exhaustion, changes in pH, temperature upshifts, and oxidative
stress had no effect on polyP levels. Analysis of Escherichia
coli mutants revealed that polyP accumulation depends on several
regulatory genes, glnD (NtrC), rpoS,
relA, and phoB.
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INTRODUCTION |
Inorganic polyphosphates (polyP) are
linear polymers of orthophosphate residues linked by high-energy
phosphoanhydride bonds. These polymers can vary in size from 3 to over
1,000 phosphate residues. PolyP is ubiquitous, having been found in
archaea, bacteria, fungi, plants, insects, and mammals (11,
25). To determine the physiological role(s) of polyP, several
polyP-metabolizing enzymes have been purified and their genes have been
cloned. The enzyme primarily responsible for polyP synthesis in
Escherichia coli is polyP kinase (PPK), which uses the gamma
phosphate of ATP to make the polymer (1, 9, 10). PolyP can
also be hydrolyzed to Pi either by
exopolyphosphatases (PPX) (2, 26) or by
endopolyphosphatases (PPN) (12). In E. coli
the genes for ppk and ppx are under the control
of a common promoter (2). The inability to produce
normal amounts of polyP upon deletion of the ppk ppx operon
has produced several striking phenotypes: (i) decreased long-term
survival in the stationary phase of growth; (ii) increased sensitivity
to oxidative, osmotic, and thermal stresses; and (iii) the appearance
of morphological variants (19).
Research on polyP has been hampered by the lack of adequate analytic
methods. Previous methods were tedious and inexact. With purified
enzymes as analytic reagents, analysis of polyP has been made specific,
but these methods are still slow, depend on radioactive labeling with
32Pi, and can be used only with a limited range
of media.
Here we describe methods for rapid extraction of polyP and a rapid and
sensitive two-step conversion of polyP into ATP by PPK and hydrolysis
of ATP by luciferase to generate light. With this assay, dynamic
accumulations of polyP in bacterial cells have been observed in
response to nutrient limitation or osmotic stress. Various E. coli mutants revealed multiple mechanisms for polyP
accumulation through NtrC-, RpoS-, RelA-, and PhoB-dependent pathways.
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MATERIALS AND METHODS |
Sources were ADP, ATP, and the ATP Bioluminescence Assay Kit CLS
II from Boehringer Mannheim; [
-32P]ATP from Amersham;
cesium chloride from U.S. Biochemicals; and Glassmilk and New Wash from
Bio101. PolyP markers of defined chain lengths were gifts from the late
Harland Wood. Purified recombinant E. coli PPK
(1) and recombinant S. cerevisiae PPX
(26) were prepared as described previously. Strains are
described in Table 1. Plasmids containing
histidine-tagged ppk or ppx genes and protocols
detailing the overexpression and purification of the enzymes are
available upon request.
In vitro preparation of [32P]polyP.
A 0.7-ml
reaction mixture contained 50 mM Tris-HCl (pH 7.4), 40 mM ammonium
sulfate, 4 mM MgCl2, 40 mM creatine phosphate, 20 ng of
creatine kinase per ml, 1 mM ATP (pH 7.2), 100 µCi of [-32P]ATP, and 72,000 U of PPK. After 30 min at
37°C, the reaction was stopped with 70 µl of 0.5 M EDTA. Conversion
of ATP to polyP was determined by ascending thin-layer chromatography
(TLC) with polyethyleneimine-cellulose in a solvent of 1 M formic
acid-0.75 M LiCl. The TLC was dried and cut, and the polyP was
determined by liquid scintillation counting. Yields were about 50% of
the [32P]ATP. The polyP reaction mixture was divided into
five equal volumes of 154 µl, and each was loaded over a 6×-volume
cushion (924 µl) of 2.5 M CsCl-50 mM Tris-HCl (pH 7.4)-10 mM EDTA.
After centrifugation at 95,000 rpm for 30 min at 20°C in a TLA 100.2 rotor (Beckman TL100 ultracentrifuge), the bottom 100 µl from each
was pooled. With addition of a 0.7× volume of isopropanol, polyP
precipitated at room temperature within 30 min and was recovered by
centrifugation at 15,000 rpm in a microcentrifuge for 15 min; the
pellet was washed with 0.3 ml of 70% ethanol, dried briefly under
vacuum, resuspended in 100 µl of distilled water, and stored at
4°C. Recoveries were typically greater than 90%. The purity of the
polyP (99%) was determined by susceptibility to hydrolysis in a
reaction mixture containing 20 mM Tris-HCl (pH 7.5), 5 mM magnesium
acetate, 50 mM ammonium acetate, and 48,000 U of PPX incubated at
37°C for 15 min (26). Pi released was
separated by TLC and quantified by liquid scintillation counting (as
above). The polyP product contained chains of approximately 750 Pi residues. Ladders of shorter lengths were generated by
mild acid hydrolysis (12) as follows: 20 µl of 2 mM
[32P]polyP750 prewarmed to 70°C was added
to 20 µl of 20 mM HCl prewarmed to 70°C; the mixtures were
maintained at 70°C from 0.5 to 1.5 min, after which the reaction was
stopped in an ice bath and by the addition of 2 µl of 1 M Tris-HCl
(pH 7.4). PolyP hydrolysis products were analyzed on a 6%
polyacrylamide-urea gel (12) and compared with polyP
markers.
Extraction of polyP with Glassmilk.
E. coli cultures
(1 ml if optical density at 600 nm [OD600] <1.5 or 0.5 ml if >1.5) were pelleted in a 1.5-ml tube for 2 min in a tabletop
microcentrifuge. The pellet was processed directly or frozen with
crushed dry ice and stored at 
80°C. To the pellet was added 0.5 ml
of 4 M guanidine isothiocyanate (GITC)-50 mM Tris-HCl, pH 7.0 (GITC
lysis buffer), prewarmed to 95°C. Tubes were vortexed, incubated for
2 to 5 min in a sand bath at 95°C, and sonicated briefly; a 10-µl
sample was removed for protein estimation, with Coomassie Plus Protein
Assay Reagent (Pierce) with bovine serum albumin as the standard
resuspended in the same buffer as the sample. To each tube was added 30 µl of 10% sodium dodecyl sulfate (SDS), 0.5 ml of 95% ethanol, and
5 µl of Glassmilk. After being vortexed, the tube was centrifuged
briefly to pellet the glass, which was then suspended in 0.5 ml of cold
New Wash buffer (5 mM Tris-HCl [pH 7.5], 50 mM NaCl, 5 mM EDTA, 50%
ethanol) by sonication in a Fisher FS30 ultrasonic cleaning bath and
repelleted; washing was repeated twice. The washed pellet was then
resuspended in 50 µl of 50 mM Tris-HCl (pH 7.4)-10 mM
MgCl2-20 µg each of DNase and RNase per ml and incubated
at 37°C for 10 min. The pellet was washed first with 150 µl of 4 M
GITC lysis buffer and 150 µl of 95% ethanol and then twice in New
Wash buffer. PolyP was eluted from the pellet with 50 µl of 50 mM
Tris-HCl (pH 8.0) at 95°C for 2 min; recovery of polyP was complete
with two additional elutions.
Assay of polyP.
PolyP was assayed in a 0.1-ml reaction
mixture (50 mM Tris-HCl [pH 7.4], 40 mM ammonium sulfate, 4 mM
MgCl2, 5 µM ADP, 24,000 U of PPK) incubated at 37°C for
40 min and then at 90°C for 2 min. The reaction mixture was diluted
1:100 in 100 mM Tris-HCl (pH 8.0)-4 mM EDTA, of which 0.1 ml was added
to 0.1 ml of luciferase reaction mixture. Luminescence was measured by
using a luminometer (Monolight 2010 [Analytical Luminescence
Laboratory] or Topcount [Packard Instruments]). A standard curve for
ATP (0, 0.55, 1.1, 1.65, and 3.3 pmol) in 100 mM Tris-HCl (pH 8.0)-4
mM EDTA was used for comparison. Concentration of polyP is given in
terms of Pi residues. [32P]polyP values are
based on specific radioactivity and susceptibility to hydrolysis with
PPX.
| |
RESULTS |
Estimation of PolyP.
Separation of polyP from ATP was achieved
by differential binding to Glassmilk (Table
2). ATP failed to bind to the glass (7%
bound), and the remaining ATP was removed in subsequent wash steps with
minimal loss of polyP (2 to 3% loss). Addition of SDS minimized the
binding of protein to the Glassmilk and enabled polyP to be completely
recovered. Greater than 98% conversion of polyP into ATP resulted when
the ADP in the reaction mixture was greater than 13-fold in excess of
the polyP, while the conversion of polyP into ATP was less than 30%
when the ADP was in twofold excess of the polyP (Fig.
1A).
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FIG. 1.
PolyP assay. (A) Conversion of [32P]polyP
to ATP. Various concentrations of [32P]polyP were reacted
with 24,000 U of PPK and 200 µM ADP in a 0.1-ml reaction mixture for
40 min at 37°C. [32P]polyP and [32P]ATP
were separated by TLC and quantitated by scintillation counting. Values
are averages of triplicate reactions. (B) ATP estimation with
luciferase. A standard curve for ATP (0, 0.55, 1.1, 1.65, 3.3 pmol of
ATP) was prepared in 100 mM Tris-HCl (pH 8.0)-4 mM EDTA and added to
an equal volume of luciferase reaction mixture, and the luminescence
was measured. A linear relationship with a correlation coefficient of
1.00 was obtained. (C) Quantitation of [32P]polyP.
[32P]polyP was added to a 0.1-ml reaction mixture
containing 12,000 U of PPK-5 µM ADP and incubated at 37°C for 40 min and then 90°C for 5 min and diluted 100-fold with 100 mM Tris-HCl
(pH 8.0)-4 mM EDTA. Then 0.1 ml of the diluted reaction mixture was
added to 0.1 ml of luciferase reaction mixture and the luminescence was
measured. A linear relationship with a correlation coefficient of 1.00 was obtained. (D) Recovery of polyP from Glassmilk.
[32P]polyP was added at various concentrations to GITC
lysates of ppk ppx cells from 0.5-ml cultures
(OD600 of 1.1), and polyP was extracted. Radioactivity
eluted and remaining on the Glassmilk pellet was used to estimate
recovery. Results are averages of triplicate assays. (E) Variation of
polyP assay. PolyP extracts as for panel D were analyzed by conversion
with PPK and ADP followed by ATP estimation with luciferase. Results
are averages of triplicate assays and include error bars. (F)
Quantitation of polyP extracts. ppk ppx cells complemented
with a plasmid overexpressing the ppk gene (pGexPPK) were
grown to an OD600 of 0.7 and then induced to express PPK by
the addition of isopropyl- -D-thiogalactopyranoside to 50 µM and grown at 30°C for 2 h. PolyP was extracted and analyzed
from 1-ml cultures as described in Materials and Methods. Serial
dilutions were initially assayed to identify an appropriate dilution of
polyP to be examined. A second round of analysis gave a linear
correlation over six determinations with a correlation coefficient of
1.00.
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ATP was determined in a linear range between 0.55 and 3.3 pmol in a
100-µl sample (Fig. 1B). All commercial samples of ADP contained
trace amounts of ATP when tested with the luciferase assay (the purest
source [Boehringer] contained 0.2%). To minimize the signal due to
ATP contamination, the polyP was converted in a 100-µl reaction
mixture containing 5 µM ADP, producing a background signal of
1.04 pmol in the reaction mixture which was further reduced by a
100-fold dilution after incubation with PPK. In vitro-synthesized [32P]polyP could be accurately quantitated with
linearity over the concentration range tested (1.4 to 13.6 pmol of
polyP in a 100-µl reaction mixture) with a correlation coefficient of
1.00. In this reaction ADP was at least 36-fold in excess of the polyP
(Fig. 1C). The estimated concentration (1.36 mM) agrees with the
concentration judged by specific radioactivity and PPX hydrolysis (1.37 mM).
Recovery of polyP from Glassmilk decreased at very low polyP
concentrations due to a relative increase in the percentage of sites of
irreversible binding to the Glassmilk (Fig. 1D). Variation in the
analysis comes from variability in the extraction (Fig. 1D)
rather than variability in the enzymatic conversion (Fig. 1E). This
level of sensitivity (500 pmol/mg of protein) is comparable to the
sensitivity of the radioactive methods. Analysis of ppk ppx mutant cells complemented with a plasmid overexpressing the ppk gene (pGexPPK) and induced to express PPK gave a linear
correlation over six determinations with a correlation coefficient of
1.00 (Fig. 1F). The linearity implies a complete conversion of polyP into ATP with an eightfold excess of ADP. The signal was
completely removed by exposure to PPX followed by heat inactivation
prior to analysis with PPK.
Nutrient limitation.
In cells grown overnight in
3-[N-morpholino]propanesulfonic acid (MOPS) medium
containing 2 mM Pi and 4 mg of glucose per ml and then
reinoculated into the same medium with limited phosphate (0.1 mM) and
amino acids (2 µg/ml), a rapid and massive transient accumulation of
polyP was observed (Fig. 2). Mutant
ppk ppx cells failed to accumulate polyP (<200 pmol/mg of
protein). PolyP values measured with the nonradioactive assay for
E. coli are in close agreement with those obtained with
radiolabeled phosphate, enrichment of [32P]polyP on
ion-exchange filters, and quantitation by susceptibility to PPX (within
95%).
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FIG. 2.
Phosphate and amino acid limitation. Cells were grown
overnight in MOPS medium and then reinoculated into the same medium
with limited amounts of phosphate (0.1 mM) and amino acids (2 µg/ml).
PolyP was assayed as described in Materials and Methods. E. coli is strain MG1655.
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The new assay applied to Acinetobacter johnsonii and
Pseudomonas aeruginosa revealed even greater
polyP accumulations (Fig. 2). E. coli grown to mid-log phase
(OD600 of 0.9) in Luria-Bertani broth (LB) and transferred
to MOPS medium with sufficient phosphate (2 mM) but without amino acids
or a carbon source accumulated polyP within the first 15 min and
continued to do so for the next 2 h before levels returned to
background over the next 4 h (Fig. 3A). Cells pelleted and resuspended in
fresh LB failed to accumulate polyP, nor did ppk ppx cells
subjected to the nutrient downshift (<200 pmol/mg protein). The
accumulations in response to the downshift could be reversed by an
upshift back into LB (Fig. 3B). This resulted in an extremely rapid
decrease in the levels of polyP (within 15 min). This pattern could be
repeated with a subsequent round of downshifting and upshifting. In the
second downshift, the rate of polyP accumulation was less rapid, while
the decrease in polyP levels to background in response to an upshift
remained immediate.
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FIG. 3.
Nutrient downshifts. (A) E. coli MG1655 cells
grown to mid-log phase in LB were downshifted by resuspension in MOPS
medium without a carbon source or amino acids. (B) E. coli
MG1655 cells grown to mid-log phase in LB were downshifted as above and
then upshifted by resuspension in LB. Growth in rich medium (LB) or SM
(MOPS medium without a carbon source or amino acids) is indicated by
dark or light bars, respectively. (C) E. coli MG1655 cells
grown to mid-log phase in LB were downshifted to MOPS medium with low
phosphate (0.1 mM) and without amino acids. PolyP was assayed as
described in Materials and Methods.
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Much greater accumulations of polyP were observed when cells were grown
in LB and then downshifted to MOPS medium with a low concentration of
Pi (0.1 mM) and 4 mg of glucose per ml and without amino
acids (Fig. 3C). After an initial hour lag, the accumulation was
extremely rapid (from less than 0.5 to 200 nmol/mg of protein within an
hour) and continued to more than double over the next hour. E. coli CF1652, which contains a deletion of the relA
gene, failed to accumulate polyP under the same conditions (Table
3).
Osmotic stress.
Cells at mid-log phase in LB, pelleted and
resuspended in LB containing 1.17 M NaCl, stopped growing immediately
and polyP levels rapidly increased (Fig.
4, closed circles). Shifts into LB of the
same osmolarity (0.17 M NaCl) did not lead to accumulations of polyP
(<200 pmol/mg of protein). A shift to LB with 0.85 M NaCl produced a
smaller transient accumulation of polyP (Fig. 4, open circles), while
shifts into LB containing 10 or 50 mM procaine, an anesthetic which is
known to stimulate the osmotic response protein EnvZ had no effect on
polyP levels (<200 pmol/mg of protein). Significant stimulation of
EnvZ by procaine is reported to occur at concentrations of 10 mM
(17). The large differences in polyP accumulation resulting
from only a 27% change in osmolarity (1.17 versus 0.85 M) may indicate
a threshold response.
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FIG. 4.
Osmotic stress. PolyP was measured for E. coli cultures grown to mid-log phase in LB and then resuspended in
LB containing 1.17 M NaCl (closed symbols) or 0.85 M NaCl (open
circles). PolyP was assayed as described in Materials and Methods.
Strains are described in Table 1.
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Osmotic stress of strains with deletions in relA or
rpoN led to significant polyP accumulations (10 to 20 nmol/mg of protein) despite the increased sensitivity of the strains to
the osmotic stress (indicated by decreases in the ODs during the stress
due to lysis or hypertonic dehydration). Strains with deletions in rpoS or phoB failed to accumulate polyP in
response to osmotic stress (Table 3). Accumulations of polyP did not
occur in response to several other stress conditions: shifts in LB
from pH 7.0 to pH 4.5, 7.0, or 9.0 or shifts from 37 to 42°C (Table
3).
Nitrogen limitation.
In cells grown in starvation medium (SM)
(6) containing a limited amount of nitrogen (2 mM), stoppage
of growth due to exhaustion of the nitrogen source coincided with a
rapid accumulation of polyP (Fig. 5A).
PolyP levels fell and growth resumed immediately after addition of
nitrogen to the medium (Fig. 5B). A similar response occurred in
strains containing deletions in relA or rpoN (Fig. 5B). In contrast, strains with deletions in
rpoS, phoB, glnD (Utase/UR), or
glnG (NtrC) did not accumulate polyP (Table 3). PolyP
accumulated in cells grown in SM to mid-log phase and then shifted to
SM lacking nitrogen and amino acids (Fig. 5C). In contrast to nitrogen
exhaustion, RpoS
strains could accumulate polyP under
this condition.
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FIG. 5.
Nitrogen limitation. (A) E. coli MG1655 cells
were inoculated into SM containing limited nitrogen. (B) Cells were
grown as for panel A, and nitrogen was added to the culture at 9.5 h. (C) Cells were grown in SM to mid-log phase and then shifted to SM
lacking nitrogen and amino acids. All cultures stopped growing after
the shift.
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DISCUSSION |
A major impediment to understanding the physiological role of
polyP has been the inadequacy of quantitative methods. The availability of purified polyP-specific enzymes (PPX and PPK) increased the specificity of assays but required extensive enrichment of polyP from
the crude extract before enzymatic conversion became feasible. The new
method involves a rapid and simple extraction and does not require
prior radioactive labeling. A limitation of the new method is its
requirement for polyP chains of at least 60 residues. It is possible
that smaller, but physiologically relevant, polyP chains escape
detection with this method.
The accumulations of polyP in response to nutritional stringencies or
osmotic shock link for the first time changes in polyP levels with
phenotypes observed in the ppk ppx mutants. Mutants which
fail to produce polyP have a decreased ability to survive in the
stationary phase and are more susceptible to osmotic stress (19). The phenotypic effects of the ppk ppx
deletion on adaptive responses to nitrogen exhaustion require further
study.
The analyses of mutant strains demonstrate multiple pathways leading to
polyP accumulation. Guanosine tetraphosphate (ppGpp) was previously
shown to inhibit PPX activity (13), thus providing a
mechanism for polyP accumulation independent of stimulated PPK activity
(Fig. 6). Accumulation of ppGpp occurs
through the actions of RelA and/or SpoT in response to a variety of
stringencies and leads to up regulation of biosynthetic operons. The
failure of relA mutants to accumulate polyP points to some
mechanism for accumulation involving ppGpp (4). Conditions
in which relA mutants accumulate polyP may also involve
ppGpp synthesized by SpoT.
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FIG. 6.
Model for stress-induced polyP accumulation. Nitrogen is
regulated by a signal cascade involving Utase/UR, NtrB, NtrC, and RpoN.
NtrC, together with RpoS and PhoB, is needed for polyP accumulation in
response to nitrogen limitation. Involvement of a sigma factor (RpoS)
implies activation of an additional factor ("X") which could lead
to polyP accumulation by direct interaction with polyP, inhibition of
PPX, stimulation of PPK, or a combination of all three. Under nutrient
limitation, ppGpp accumulates by RelA and SpoT actions, which can lead
to polyP accumulation by PPX inhibition and/or RpoS activation. Failure
to accumulate polyP when ppGpp and RpoS levels are high (such as under
carbon starvation) implies the presence of additional regulator(s).
Osmotic stress triggers polyP accumulation through a mechanism that
does not involve EnvZ, the osmotic sensor.
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Factors in addition to ppGpp are necessary for accumulation under
different stress conditions. The rpoS gene of E. coli encodes a sigma factor required for the expression
of over 50 genes induced in the stationary phase, involved in stress
resistance and long-term survival (16), and known to be
partially regulated by ppGpp (4). rpoS mutants
failed to accumulate polyP in response to nitrogen exhaustion and
osmotic stress. Dependence on a sigma factor implies a requirement for
an additional factor(s) for polyP accumulation, designated "X" in
Fig. 6. In contrast to the rpoS-dependent polyP accumulation
reported here, it has been reported that polyP-depleted cells were
unable to induce transcription of both rpoS and
katE (an rpoS-dependent gene) (22). An
interdependent regulatory mechanism could account for this apparent
codependence for stimulation between polyP and rpoS. An
inhibition of rpoS expression in the ppk ppx
mutant is consistent with the observed phenotypes (19).
Under some conditions in which ppGpp and RpoS accumulations are known
to occur, such as carbon starvation, acid or alkaline stress,
temperature stress, and oxidative stress (7, 23), polyP
failed to accumulate (Table 3), indicating that an additional factor(s)
is involved. Clearly then, polyP accumulation is not a general response
to stress but occurs as a result of specific stresses. With regard to
salt stress, the failure of procaine to trigger polyP accumulation
demonstrates that the EnvZ-mediated pathway is not involved. PolyP
accumulation in response to salt stress could be triggered by secondary
effects of the hyperosmolarity, such as nutrient or nitrogen depletion.
Accumulation of polyP in response to nitrogen exhaustion directly links
the nitrogen regulation signaling cascade with rpoS and
changes in phosphate metabolism. RpoN mutants were able to accumulate
polyP in response to nitrogen exhaustion, but mutations in Utase/UR and
NtrC, the regulator, abolished this ability (Table 3), indicating a
signaling mechanism in which NtrC can act as the response regulator for
RpoS to activate a factor(s), "X", which could then trigger polyP
accumulation by mechanisms involving the stimulation of PPK activity,
the inhibition of PPX activity, or both (Fig. 6).
In adaptations to stress, cells must coordinate major changes in the
rates of transcription, translation, and replication as well as choices
in the genes expressed (8). PolyP could provide activated
phosphates or coordinate an adaptive response by binding metals and/or
specific proteins. The transient nature of the accumulations and the
rapid turnover rate of ATP in the cell (the entire pool of ATP is
typically turned over in 0.2 s [5]) argues
against polyP simply as an ATP store. As a polyanionic polymer, polyP
has chemical similarities to DNA and RNA in interactions with basic
domains of proteins that interact with DNA and RNA. PolyP was reported
to be associated with RNA polymerase isolated from stationary-phase
E. coli (15). Further investigation of the
cellular location of polyP, its state of metabolic availability, and
identification of its binding partners should help to clarify this
issue.
We thank the late Harland Wood for his generous gift of polyP
markers; Michael Cashel, Larry Reitzer, and Anand Chakrabarty for their
generous gifts of E. coli strains and helpful discussions; Christian Itin for numerous helpful discussions during the course of
this study and preparation of the manuscript; and Wayne Fiori for help
with the computers.
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Abstract
Introduction
