(Received for publication, July 13, 1995; and in revised form, October 6, 1995)
From the
In an earlier study a mutant Dictyostelium cell-line (plc) was constructed in which all
phospholipase C activity was disrupted and nonfunctional, yet these
cells had nearly normal Ins(1,4,5)P
levels (Drayer, A. L.,
Van Der Kaay, J., Mayr, G. W, Van Haastert, P. J. M. (1990) EMBO J. 13, 1601-1609). We have now investigated if these cells have
a phospholipase C-independent de novo pathway of
Ins(1,4,5)P
synthesis. We found that homogenates of plc
cells produce Ins(1,4,5)P
from endogenous precursors. The enzyme activities that performed
these reactions were located in the particulate cell fraction, whereas
the endogenous substrate was soluble and could be degraded by phytase.
We tested various potential inositol polyphosphate precursors and found
that the most efficient were Ins(1,3,4,5,6)P
,
Ins(1,3,4,5)P
, and Ins(1,4,5,6)P
. The
utilization of Ins(1,3,4,5,6)P
, which can be formed
independently of phospholipase C by direct phosphorylation of inositol
(Stephens, L. R. and Irvine, R. F.(1990) Nature 346, 580-582),
provides Dictyostelium with an alternative and novel pathway
of de novo Ins(1,4,5)P
synthesis. We further
discovered that Ins(1,3,4,5,6)P
was converted to
Ins(1,4,5)P
via both Ins(1,3,4,5)P
and
Ins(1,4,5,6)P
. In the absence of calcium no
Ins(1,4,5)P
formation could be observed; half-maximal
activity was observed at low micromolar calcium concentrations. These
reaction steps could also be performed by a single enzyme purified from
rat liver, namely, the multiple inositol polyphosphate phosphatase.
These data indicate that organisms as diverse as rat and Dictyostelium possess enzyme activities capable of
synthesizing the second messengers Ins(1,4,5)P
and Ins(1,
3, 4, 5)P
via a novel phospholipase C-independent pathway.
It is now well established that the occupation of cell-surface
receptors on many mammalian cells leads to an activation of
phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P) (
)hydrolysis by phospholipase C (PLC), (
)and the
concomitant release of inositol 1,4,5-trisphosphate
(Ins(1,4,5)P
), which mobilizes cellular Ca
stores (Berridge and Irvine, 1989). Termination of the signaling
activities of Ins(1,4,5)P
is accomplished by a
5-phosphatase (reviewed by Verjans et al.(1994)) to form
Ins(1,4)P
. In addition an Ins(1,4,5)P
3-kinase
(reviewed by Shears(1992)) yields the putative second messenger
Ins(1,3,4,5)P
(see Irvine(1992)). Several aspects of this
signal transduction process are present in the cellular slime mold Dictyostelium discoideum, which, due to its genetic
tractability, is therefore a useful model system. For example, Dictyostelium cells possess cell-surface receptors for the
chemoattractant cAMP that are coupled via G-proteins to a phospholipase
C (Bominaar and Van Haastert, 1994); thus, extracellular stimulation of
cells with cAMP elevates intracellular Ins(1,4,5)P
levels
(Van Haastert, 1989). There is also evidence that in Dictyostelium Ins(1,4,5)P
releases Ca
from
nonmitochondrial stores (Europe-Finner and Newell, 1986; Flaadt et
al., 1993).
The Dictyostelium gene for phospholipase C
has been cloned, and the enzyme appeared to be structurally similar to
PLC- found in mammalian cells (Drayer and Van Haastert, 1992).
Subsequently, a Dictyostelium cell line with a disrupted
phospholipase C gene was constructed (Drayer et al., 1994).
These plc
cells had no measurable
phospholipase C activity, but they expressed a normal phenotype. More
surprisingly, the basal levels of Ins(1,4,5)P
were only 20%
lower than that of wild-type cells (Drayer et al., 1994). This
observation led to the suggestion that there might be a phospholipase
C-independent source of Ins(1,4,5)P
. This suggestion is
without precedent, and even the demonstration that Dictyostelium can convert Ins(1,3,4,5)P
to Ins(1,4,5)P
by a 3-phosphatase activity (Van Dijken et al., 1995)
does not resolve this problem, since the only known source for
Ins(1,3,4,5)P
is Ins(1,4,5)P
itself.
These
results raise fundamental questions concerning the sources of both
Ins(1,4,5)P and Ins(1,3,4,5)P
in both wild-type
and plc
Dictyostelium cells. We now
describe a novel metabolic pathway by which these two inositol
phosphates can be formed by dephosphorylation of
Ins(1,3,4,5,6)P
. Moreover, we further discovered that these
enzyme activities were similar to those that can also be catalyzed by a
single mammalian enzyme that was recently renamed as a multiple
inositol polyphosphate phosphatase (MIPP) (Craxton et al.,
1995). These observations indicate that we may have to reevaluate the
mechanisms by which cellular Ins(1,4,5)P
is generated and
utilized.
[3-P]Ins(1,3,4,5)P
was prepared
from Ins(1,4,5)P
and [
-
P]ATP as
described in Van Dijken et al.(1994).
[
H]Ins(1,3,4,5,6)P
and DL-[
C]Ins(1,4,5,6)P
were isolated, respectively, from
[
H]inositol-labeled or
[
C]inositol-labeled turkey erythrocytes
(Stephens and Downes, 1990).
[
H]Ins(1,4,5,6)P
was synthesized as
described by Craxton et al.(1994). Ins(1,2,4,5,6)P
was prepared as described by Ye et al.(1995).
A phytase-treated soluble fraction was prepared by
overnight incubation of 250 µl of soluble fraction in a 285-µl
incubation with 0.2 units/ml A. ficuum phytase at 55 °C in
the presence of 50 mM sodium acetate, pH 5.15. The reaction
was stopped by boiling for 60 min to totally inactivate the phytase. In
a control experiment, added [H]InsP
(15,000 dpm) was totally degraded by the phytase to
[
H]InsP and [
H]inositol.
Identification
with Type I Ins(1,4,5)P/Ins(1,3,4,5)P
5-phosphatase was carried out similarly except that the reaction
mixture contained 100 mM Hepes, pH 7.5, 1 mg/ml BSA, 2 mM MgCl
, 5 mM 2-mercaptoethanol, and the sample
was incubated for 10 min. at 37 °C.
Figure 1:
Formation of Ins(1,4,5)P cross-reactivity by Dictyostelium cellular fractions
from endogenous and exogenous substrates. A, Dictyostelium particulate and soluble fraction were obtained by centrifugation
of a lysate. Some fractions were boiled or treated with phytase before
they were incubated as indicated in the figure. Samples were
incubated in the presence of 10 mM CaCl
for 0 and
30 min and then quenched and neutralized as described under
``Experimental Procedures.'' 20-µl aliquots of the final,
quenched extracts were analyzed for Ins(1,4,5)P
cross-reactivity by the isotope dilution assay using
Ins(1,4,5)P
binding protein. The data in the figure were obtained by subtracting the cross-reactivity at zero time
from that at 30 min. Data shown are the means and standard errors of
mean of at least three experiments. Note that the samples soluble
+ phytase + particulate and soluble + boiled
phytase + particulate (which serves as a control for the
former) are 5% more diluted than the other samples. Bars indicated with a star indicate Ins(1,4,5)P
cross-reactivity accumulation during the incubation time, that is
significantly different from zero as assessed by Student's t test (p
0.05). B, unlabeled inositol
phosphates were incubated for 0 and 30 min with Dictyostelium particulate fraction in the presence of 10 mM CaCl
. Ins(1,4,5)P
cross-reactivity
accumulating per 10 pmol of substrate is shown as mean and standard
error of the mean. The experiments were performed at least in
triplicate. The amount of Ins(1,4,5)P
cross-reactivity
accumulating during the incubation period is for all values
significantly different from zero as assessed by Student's t test (p
0.05).
Ins(1,4,5)P/Ins(1,3,4,5)P
5-phosphatase and
Ins(1,4,5)P
3-kinase were used to investigate the extent to
which the material that expressed Ins(1,4,5)P
cross-reactivity was genuine Ins(1,4,5)P
, as follows.
Samples with known amounts of Ins(1,4,5)P
cross-reactivity
were prepared from incubations containing cell lysate as described in
the legend to Fig. 1A.
[
H]Ins(1,4,5)P
was added to these
samples, which were then incubated with either 5-phosphatase or
3-kinase. After such incubations, the residual Ins(1,4,5)P
cross-reactivity was determined and compared with the amount of
metabolism of [
H]Ins(1,4,5)P
. Both
with 5-phosphatase and 3-kinase, the decrease in Ins(1,4,5)P
cross-reactivity amounted to 75% of the amount of
[
H]Ins(1,4,5)P
metabolized (data not
shown). Thus 75% of the material that expressed Ins(1,4,5)P
cross-reactivity represented authentic Ins(1,4,5)P
.
The remaining 25% of the Ins(1,4,5)P
cross-reactivity was
not further investigated but could have been due to the presence of
other inositolphosphates. For example
Ins(1,2,4)P
/Ins(2,3,6)P
has previously been
shown to be a potent ligand of the Ins(1,4,5)P
receptor
(Freund et al., 1992).
The Ins(1,4,5)P-forming
activity present in the homogenate required both soluble and
particulate components, as these separate fractions did not form
Ins(1,4,5)P
unless mixed (Fig. 1A). The
factor in the particulate fraction was heat-labile, whereas the soluble
factor was heat stable. Preincubation of the soluble fraction with the
inositol polyphosphate-degrading enzyme, phytase, abolished
Ins(1,4,5)P
formation (Fig. 1A). These data
suggest that the particulate cell fraction contains enzyme activities
that form Ins(1,4,5)P
, whereas the soluble fraction
contains endogenous substrates that are presumably inositol
polyphosphates, because they are degraded by phytase.
Figure 2:
Calcium dependence of the accumulation of
Ins(1,4,5)P cross-reactivity. The accumulation of
Ins(1,4,5)P
cross-reactivity was determined at various free
CaCl
concentrations, under conditions described in Fig. 1B, with Ins(1,3,4,5)P
(
) or
Ins(1,3,4,5,6)P
(
) as substrates.
Ins(1,3,4,5,6)P
was also used as substrate (
) for
Ins(1,4,5)P
in the presence of 1 mM MgCl
and 25 mM LiCl, and 0.5 mM
2,3-diphospho-D-glycerate as inhibitors of Ins(1,4,5)P
degradation. The accumulation of Ins(1,4,5)P
cross-reactivity is expressed as a percentage of maximal
Ins(1,4,5)P
accumulation. The data presented in this figure were fitted with the Hill coefficient set at 1. Free
Ca
concentrations were set using
Ca
/EGTA buffers as described by
Bartfai(1979).
The Ca dependence was also investigated in the
presence of 1 mM MgCl
. As MgCl
strongly enhances the degradation of Ins(1,4,5)P
, two
inhibitors of Ins(1,4,5)P
phosphatases were included:
lithium to inhibit Ins(1,4,5)P
1-phosphatase and
2,3-diphospho-D-glycerate to inhibit Ins(1,4,5)P
5-phosphatase (Van Lookeren Campagne et al., 1988). In
the absence of CaCl
no accumulation of Ins(1,4,5)P
cross-reactivity was observed. However, at submicromolar
Ca
concentration Ins(1,4,5)P
formation
was detectable; half-maximal activity was observed at 1.3 µM Ca
(Fig. 2).
In the remaining part of
this study we use conditions in which no MgCl but high
concentrations of total CaCl
(10 mM) are present.
These conditions are used as a tool to further explore the formation of
Ins(1,4,5)P
as under these conditions Ins(1,4,5)P
phosphatase activity is greatly reduced, which precludes the need
of phosphatase inhibitors that might interfere with the phosphatases
under investigation.
Figure 3:
Identification of InsP isomers
formed from Ins(1,3,4,5,6)P
degradation by Dictyostelium particulate fraction. A,
[
H]Ins(1,3,4,5,6)P
(about 4000 dpm)
was incubated with Dictyostelium particulate fraction (5
10
lysed cells/ml) for 30 min as described under
``Experimental Procedures.'' The sample was analyzed on a
Zorbax SAX column eluted with a gradient that does not separate
InsP
isomers. The gradient used was 0 min, 0% B; 10 min,
100% B. Fractions of 0.75 ml were collected. The InsP
fractions obtained in an experiment similar to that described in panel A were desalted and used in the experiments presented in panels B and C. B, an aliquot of the
InsP
fraction (see panel A) was incubated in the
absence (
, 600 dpm) or presence (
, 950 dpm) of Ins(1,4,5,
6)P
3-kinase as described under ``Experimental
Procedures,'' and chromatographed such that InsP
isomers were separated on an Adsorbosphere SAX column as
described by Menniti et al.(1990). The second InsP
peak, co-eluting with the internal standard of DL-[
C]Ins(1,4,5,6)P
,
was converted by the 3-kinase, identifying it as
Ins(1,4,5,6)P
. C, an aliquot of the InsP
fraction (see panel A) was incubated in the presence
(
, 500 dpm) or absence (
, 400 dpm) of purified
Ins(1,4,5)P
/Ins(1,3,4,5)P
5-phosphatase as
described under ``Experimental Procedures'' and analyzed on a
Zorbax SAX column that was eluted with a gradient such that InsP
isomers were separated. The gradient used was 0 min, 0% B; 5 min,
30% B; 35 min, 46% B; 36 min, 100% B; 1-ml fractions were collected.
The first InsP
peak, co-eluting with an internal
[
P]Ins(1,3,4,5)P
standard, was
degraded by the 5-phosphatase, identifying it as
Ins(1,3,4,5)P
.
The total
[H]InsP
fraction formed following
[
H]Ins(1,3,4,5,6)P
degradation by Dictyostelium particulate fraction (see Fig. 3A) was isolated, desalted, and rechromatographed
using a different HPLC system that resolved this material into two
InsP
peaks. Fig. 3B shows that the
InsP
fraction contained two main InsP
isomers.
Incubation of this material with purified Ins(1,4,5,6)P
3-kinase revealed that 88% of the second InsP
peak
was converted to Ins(1,3,4,5,6)P
(Fig. 3B).
This identifies the second peak as minimally 88% Ins(1,4,5,
6)P
. The first InsP
peak co-eluted with an
internal standard of
[
C]Ins(1,3,4,5)P
. Upon incubation of
the total [
H]InsP
fraction with
Ins(1,4,5)P
/Ins(1,3,4,5)P
5-phosphatase the
first [
H]InsP
peak was totally
degraded (Fig. 3C) to a compound with the same
retention time as Ins(1,3,4)P
(data not shown) identifying
the first [
H]InsP
peak as
Ins(1,3,4,5)P
. From these data we concluded that
Ins(1,3,4,5,6)P
is mainly metabolized to
Ins(1,3,4,5)P
and Ins(1,4,5,6)P
. This is a
significant conclusion because data summarized in Fig. 1B indicate that both Ins(1,3,4,5)P
and
Ins(1,4,5,6)P
are precursors for material that expressed
Ins(1,4,5)P
cross-reactivity. It was therefore important to
prove that authentic Ins(1,4,5)P
was formed from these two
InsP
isomers.
The metabolism of Ins(1,3,4,5)P was investigated using a mixture of
[3-
P]Ins(1,3,4,5)P
and
[
H]Ins(1,3,4,5)P
. Incubation of this
mixture with particulate fraction resulted in the formation of
InsP
, which had a
P:
H ratio that
was 25% that of the original Ins(1,3,4,5)P
. Since
P is removed from the 3-position of
Ins(1,3,4,5)P
, these data demonstrate that 75% of the
InsP
was Ins(1,4,5)P
. The remainder of the
InsP
fraction still contained the
P label at
the 3-position and was not further identified. In a previous study,
using different conditions, the isomers of InsP
that were
formed were identified as Ins(3,4,5)P
and Ins(1,4,5)P
(Van Dijken et al., 1995). The metabolism of
[
H]Ins(1,4,5,6)P
by particulate
fraction was also investigated. The resultant
[
H]InsP
fraction was isolated by
HPLC, desalted, and incubated with
Ins(1,4,5)P
/Ins(1,3,4,5)P
5-phosphatase,
whereupon 72% of the [
H]InsP
was
degraded to InsP
(Fig. 4B). Under identical
conditions authentic [
H]Ins(1,4,5)P
was completely degraded. This means that 72% of the InsP
formed from Ins(1,4,5,6)P
is Ins(1,4,5)P
.
Figure 4:
Identification of Ins(1,4,5)P formed from Ins(1,3,4,5)P
and Ins(1,4,5,6)P
degradation by Dictyostelium particulate fraction. A, [3-
P]Ins(1,3,4,5)P
(3000
dpm,
) and [
H]Ins(1,3,4,5)P
(7000 dpm,
) were mixed and degraded for 30 min with Dictyostelium particulate fraction (5
10
lysed cells/ml) as described under ``Experimental
Procedures.'' The reaction products were separated on a Zorbax SAX
column. The gradient used was 0 min, 0% B; 10 min, 10% B. The
P:
H ratio in the formed InsP
fraction is 0.25 relative to this ratio in the
Ins(1,3,4,5)P
fraction. B,
[
H]Ins(1,4,5,6)P
was incubated for 30
min with Dictyostelium particulate fraction (5
10
lysed cells/ml) as described under ``Experimental
Procedures.'' The InsP
fraction was isolated,
desalted, and incubated for 10 min at 37 °C with (350 dpm,
)
or without (
, 300 dpm) 10 µl of purified
Ins(1,4,5)P
/Ins(1,3,4,5)P
5-phosphatase in a
150-µl incubation containing 50 mM Hepes, pH 7.5, 1 mg/ml
BSA, 5 mM MgCl
, 5 mM 2-mercaptoethanol.
The sample was analyzed on a Zorbax SAX column. The gradient used was 0
min, 0% B; 3 min, 20% B; 12 min, 100% B.
Together these data indicate that Ins(1,3,4,5,6)P is
degraded to two InsP
isomers, Ins(1,3,4,5)P
and
Ins(1,4,5,6)P
. These InsP
isomers are degraded
to predominantly Ins(1,4,5)P
.
Figure 5:
Identification of the InsP
product of MIPP-catalyzed hydrolysis of Ins(1,4,5,6)P
.
[
H]Ins(1,4,5,6)P
was hydrolyzed by
MIPP as described in the legend to Fig. 6, and the
[
H]InsP
(1500 dpm,
) was
isolated by HPLC, desalted, and incubated after the addition of
[
C]Ins(1,4,5)P
(200 dpm,
)
for 10 min at 37 °C with a 2500-fold dilution of recombinant rat
brain Ins(1,4,5)P
3-kinase exactly as described by Craxton et al.(1994). Reactions were quenched, neutralized, and
chromatographed by HPLC as described by Menniti et al.(1990).
The 50-110-min portion of the HPLC gradient depicts co-elution of
both the unknown [
H]InsP
with the
[
C]Ins(1,4,5)P
standard and
co-chromatography of the [
H]InsP
product with the internally derived
[
C]Ins(1,3,4,5)P
standard. The
relative position of a
[
H]Ins(1,3,4,6)P
standard in a
subsequent HPLC elution profile is indicated by the arrow.
Further assays (see inset) contained various dilutions of
recombinant Ins(1,4,5)P
3-kinase (100-10,000-fold).
Data are expressed as the percentage phosphorylation of either
[
C]Ins(1,4,5)P
(closed
squares) or [
H]InsP
(open
squares). No significant phosphorylation of either
[
C]Ins(1,4,5)P
or the unknown
[
H]InsP
was observed with control E. coli lysates (100-fold
dilution).
Figure 6:
Dephosphorylation of Ins(1,4,5,6)P and Ins(1,3,4,5,6)P
by purified MIPP. Trace
quantities of either [
H]Ins(1,4,5,6)P
(4000 dpm,
) or
[
H]Ins(1,3,4,5,6)P
(4000 dpm,
)
were incubated with purified MIPP (0.95 ng) for the indicated times.
The [
H]Ins(1,3,4,5,6)P
used in this
experiment was also the precursor of this batch of
[
H]Ins(1,4,5,6)P
; therefore, the
specific activities of these polyphosphates were identical.
[
H]inositol phosphates were separated on Bio-Rad
AG1-X8 columns. Data represent averages of duplicate incubations. Inset, inhibition of MIPP-catalyzed dephosphorylation of
[
H]Ins(1,4,5,6)P
(4000 dpm,
)
and [
H]Ins(1,3,4,5,6)P
(4000 dpm,
) by Ins(1,3,4,5)P
(0-10 µM).
[
H]inositol phosphates were resolved as described
above, and data were expressed as a percentage of MIPP activity in the
absence of Ins(1, 3, 4, 5)P
. The conditions used were such
that less than 20% of the substrate was
metabolized.
The unexpected demonstration that MIPP
could catalyze Ins(1,4,5,6)P 6-phosphatase activity now led
us to reconsider the specificity of this enzyme's attack upon
Ins(1,3,4,5,6)P
. Previous experiments only identified an
Ins(1,3,4,5,6)P
3-phosphatase activity (Nogimori et
al., 1991), but we have now greatly increased the sensitivity of
such an assay by incubating MIPP with a large amount (750,000 dpm) of
[
H]Ins(1,3,4,5,6)P
. Fig. 7shows that in addition to the large amount of
[
H]Ins(1,4,5,6)P
that we anticipated
would be formed (97% of total
[
H]InsP
), small amounts of two
additional InsP
isomers were detected. One of these (2% of
total [
H]InsP
) eluted just before the
[
C]Ins(1,3,4,5)P
standard and was
therefore identified as
[
H]Ins(1,3,4,6)P
. Another
[
H]InsP
peak (1% of total
[
H]InsP
) co-eluted with an internal
[
C]Ins(1,3,4,5)P
standard and must
therefore be Ins(1,3,4,5)P
and/or
Ins(1,3,5,6)P
. These data provide only a minimum estimate
of flux through Ins(1,3,4,5)P
, since this InsP
isomer will be further metabolized to Ins(1,4,5)P
by
MIPP.
Figure 7:
InsP isomers formed from
MIPP-mediated hydrolysis of [
H]
Ins(1,3,4,5,6)P
. MIPP (0.4 µg) was incubated for 3 h at
37 °C in 1.33 ml of medium containing 100 mM KCl, 50
mM Bis-Tris, pH 6.1, 1 mM EDTA, 0.5 mM EGTA,
2 mM CHAPS, 0.05% (w/v) BSA, and approximately 750,000 dpm
[
H]Ins(1,3,4,5,6)P
(
).
Incubations were quenched with perchloric acid and neutralized with
freon/octylamine as described under ``Experimental
Procedures.'' Samples were spiked with an internal standard of
approximately 400 dpm [
C]Ins(1,3,4,5)P
(
) and chromatographed by HPLC on an Adsorbosphere SAX
column as described by Menniti et al.(1990). The inset shows a magnification of the area indicated with a box.
Prior to these studies, the only known de novo source for Ins(1,4,5)Pin vivo was
phospholipase C-mediated hydrolysis of PtdIns(4,5)P
. Even
though cells contain a variety of highly phosphorylated inositol
polyphosphates, there has been general acceptance of the axiom that
these compounds ``must be synthesised and degraded without
inadvertently generating significant quantities of potent signals such
as Ins(1,4,5)P
and Ins(1,3,4,5)P
'' (Downes
and McPhee, 1990). Thus, it was completely unexpected to discover that
a Dictyostelium mutant in which the gene for phospholipase C
was disrupted and rendered nonfunctional was nevertheless capable of
maintaining near normal unstimulated levels of both Ins(1,4,5)P
and Ins(1,3,4,5)P
(Drayer et al., 1994).
This led to the conclusion that there may be a phospholipase
C-independent source for these second messengers and therefore
initiated an unprecedented search for an alternative pathway by which
these compounds could be synthesized. Here we demonstrate that
Ins(1,3,4,5,6)P
is a physiologically relevant precursor of
both Ins(1,4,5)P
and Ins(1,3,4,5)P
in Dictyostelium. Moreover, we have further demonstrated that the
same enzymatic reactions can also occur in rat liver by, somewhat
surprisingly, the action of a single enzyme.
The mammalian enzyme
that degrades Ins(1,3,4,5,6)P to Ins(1,4,5)P
was originally identified as an Ins(1,3,4,5)P
3-phosphatase (Cunha-Melo et al., 1988; Doughney et
al., 1988; Höer et al., 1988).
Recently, it was suggested that the enzyme be renamed a multiple
inositol polyphosphate phosphatase (MIPP) since it also removed the
3-phosphate from Ins(1,3,4,5,6)P
and could nonspecifically
dephosphorylate InsP
(Nogimori et al., 1991;
Craxton et al., 1995). Here we show that this enzyme is also
capable of hydrolyzing the 6-phosphate of both Ins(1,3,4,5,6)P
and Ins(1,4,5,6)P
. This new observation can be
rationalized by noting the similarities of the structures of
``inverted'' Ins(1,4,5,6)P
and
Ins(1,3,4,5)P
; the inositol ring pucker is similar in both
cases, and the 3-hydroxy, 4-phosphate, 5-phosphate, and 6-phosphate of
``inverted'' Ins(1,4,5,6)P
, respectively, mimic
the configuration of the 6-hydroxy, 5-phosphate, 4-phosphate, and
3-phosphate of Ins(1,3,4,5)P
(Fig. 8). Provided
these similarities satisfy the requirements for substrate recognition
and that the nature and orientation of the C-1 and C-2 substituents are
not critical determinants of substrate recognition, the 6-phosphate of
Ins(1,4,5,6)P
can be considered to mimic the 3-phosphate of
Ins(1,3,4,5)P
. An attack upon the 6-phosphate of
Ins(1,3,4,5,6)P
can be explained similarly since the
3-phosphate of Ins(1, 3,4,5,6)P
is mimicked by the
6-phosphate once the Ins(1,3,4,5,6)P
is
``inverted.''
Figure 8:
Comparison of the structures of
Ins(1,3,4,5)P and ``inverted''
Ins(1,4,5,6)P
.
The exact relationship between the enzyme
activities in Dictyostelium particulate fraction and rat MIPP
is still unclear. Despite the striking similarities of the net
dephosphorylation of Ins(1,3,4,5,6)P to Ins(1,4,5)P
and the similar localization of the enzyme activities to the
particulate portion of the cells, major differences between the enzyme
activities in Dictyostelium particulate fraction and rat MIPP
exist. For example, Ins(1,3,4,5)P
is, at best, a relatively
minor product to accumulate when MIPP hydrolyzes
Ins(1,3,4,5,6)P
, whereas in Dictyostelium particulate fraction comparable amounts of Ins(1,3,4,5)P
and Ins(1,4,5,6)P
accumulate. This difference may
reflect a similarity in the rates of Dictyostelium 3- and
6-phosphatase activities toward Ins(1,3,4,5,6)P
, in sharp
contrast to the substantial preference of MIPP for the 3-phosphate of
Ins(1,3,4,5,6)P
. The most striking difference lay in the
effect of CaCl
on the enzyme reaction; the net conversion
of Ins(1,3,4,5,6)P
to Ins(1,4,5)P
by the Dictyostelium phosphatase activities is only detectable in the
presence of Ca
, but when we investigated the effect
of up to 1 mM Ca
upon MIPP, the activity was
increased by no more than 20% (data not shown).
In order for hepatic
MIPP (or any other mammalian species of this enzyme) to produce
significant amounts of Ins(1,4,5)P from
Ins(1,3,4,5,6)P
in vivo, both the enzyme and the
Ins(1,3,4,5,6)P
would have to be in a compartment that did
not have access to even the tiny amounts of InsP
that
competitively inhibit this enzyme activity (Nogimori et al.,
1991). As it happens, in rat liver at least, MIPP resides inside the
endoplasmic reticulum (Ali et al., 1993) but we have no
evidence that Ins(1,3,4,5,6)P
has specific access to the
enzyme in vivo. Nevertheless, we should still pursue this
possibility, since if it is correct that there is a cellular
subcompartment in which a pool of Ins(1,4,5)P
can be
generated independently of PtdIns(4,5)P
, this could finally
address the longstanding enigma that a number of unstimulated mammalian
cell types contain rather more Ins(1,4,5)P
than is
necessary to fully mobilize Ca
stores (see
Shears(1992)). We should also consider that there might be
physiological relevance to the demonstration that MIPP produced small
amounts of Ins(1,3,4,6)P
from Ins(1,3,4,5,6)P
,
since just such a reaction was hypothesized to occur in 3T3 fibroblasts
(Balla et al., 1994). The evidence for Ins(1,3,4,5,6)P
being a physiologically relevant precursor of Ins(1,4,5)P
is much stronger in the case of Dictyostelium. For
example, there are kinetic differences between the
Ins(1,4,5)P
-forming activities of MIPP as compared with the Dictyostelium enzyme (see above), and it seems from our data
that the latter does not need to be spatially separated from InsP
in order to function. More importantly, total steady-state levels
of Ins(1,4,5)P
in vivo were only slightly
decreased by rendering inactive all phospholipase C activity in this
organism, indicating that another de novo pathway was present
(Drayer et al., 1994). Moreover, previously published data on
the inositol phosphates present in intact Dictyostelium cells
substantiate the idea that in this organism the conversion of
Ins(1,3,4,5,6)P
to Ins(1,4,5)P
is of
physiological relevance. Drayer et al.(1994) showed that, in plc
cells that contained near normal levels
of Ins(1,4,5)P
and Ins(1,3,4,5)P
, the levels of
Ins(1,3,4,5,6)P
were specifically reduced by around 70%.
Indeed, this reduction in levels of Ins(1,3,4,5,6)P
is
indicative of an adaptive response in order to sustain the steady-state
levels of important signaling molecules such as Ins(1,4,5)P
and Ins(1,3,4,5)P
. Although wild-type Dictyostelium cells contain large quantities of several
InsP
isomers (Stephens et al., 1991), it is
Ins(1,3,4,5,6)P
that is formed as an intermediate in the
stepwise phosphorylation of inositol to InsP
(Stephens and
Irvine, 1990). Together these results suggest an independent de
novo link between Ins(1,4,5)P
and inositol (Fig. 9). This conclusion should also prompt research into the
possibility that mammalian tissues might also synthesize some
Ins(1,3,4,5,6)P
and InsP
by this pathway.
Figure 9: The inositol cycle in Dictyostelium discoideum. The metabolism of inositol in the slime mold Dictyostelium is shown. The metabolic steps presented in this study are indicated with broad arrows. For simplicity not all metabolic pathways of PIP and PI are indicated in the figure.
Since the role of Ins(1,4,5)P is itself to mobilize
Ca
, the observation that the formation of
Ins(1,4,5)P
from Ins(1,3,4,5,6)P
is
Ca
dependent, raise some hitherto unexpected and
complex questions concerning the interactions between levels of
Ca
and Ins(1,4,5)P
. It is possible that
the calcium-mediated activation of the Dictyostelium enzyme is
indicative of an ongoing regulatory process in the cytosol. Certainly
the observation that organisms as diverse as Dictyostelium and
the rat both possess the enzymatic capacity to synthesize
Ins(1,4,5)P
from Ins(1,3,4,5,6)P
, essentially
independently of short term PtdIns(4,5)P
turnover, should
provoke us to reevaluate the cellular control processes that regulate
Ca
mobilization.