(Received for publication, November 19, 1996, and in revised form, February 19, 1997)
From the Institute of Biochemistry, University of
Genova, 16132 Genova, Italy and the ¶ Department of Physiology,
University of Minnesota, Minneapolis, Minnesota 55455
CD38, a lymphocyte differentiation antigen, is
also a bifunctional enzyme catalyzing the synthesis of cyclic
ADP-ribose (cADPR) from NAD+ and its hydrolysis to
ADP-ribose (ADPR). An additional enzymatic activity of CD38 shared by
monofunctional ADP-ribosyl cyclase from Aplysia californica
is the exchange of the base group of NAD+ (nicotinamide)
with various nucleophiles. Both human CD38 (either recombinant or
purified from erythrocyte membranes) and Aplysia cyclase
were found to catalyze the exchange of ADPR with the nicotinamide group
of NAD+ leading to the formation of a dimeric ADPR
((ADPR)2). The dimeric structure of the enzymatic product,
which was generated by recombinant CD38 and by CD38+
Namalwa cells from as low as 10 µM NAD+, was
demonstrated using specific enzyme treatments (dinucleotide pyrophosphatase and 5-nucleotidase) and mass spectrometry analyses of
the resulting products. The linkage between the two ADPR units of
(ADPR)2 was identified as that between the N1
of the adenine nucleus of one ADPR unit and the anomeric carbon of the
terminal ribose of the second ADPR molecule by enzymatic analyses and
by comparison with patterns of cADPR cleavage with Me2SO:tert-butoxide. Although
(ADPR)2 itself did not release Ca2+ from sea
urchin egg microsomal vesicles, it specifically potentiated the
Ca2+-releasing activity of subthreshold concentrations of
cADPR. Therefore, (ADPR)2 is a new product of CD38 that
amplifies the Ca2+-mobilizing activity of cADPR.
NAD+ can be metabolized by either symmetrical
hydrolysis of the pyrophosphate bond by a number of pyrophosphatases
(1, 2) or by cleavage of the -glycosyl linkage between nicotinamide and ribose (3). The latter linkage has been shown to be cleaved by an
increasing number of enzymes belonging to diverse classes and having
widely varying cellular distribution. For example, the classical
NAD+ glycohydrolases, which catalyze the hydrolysis of
NAD+ to nicotinamide and adenosine diphosphate ribose
(ADPR),1 are mostly ectoenzymes in
mammalian cells (4-7). Endogenous ADP-ribosyl transferases responsible
for either mono-ADP-ribosylation (8-12) or poly(ADP-ribosylation) (13)
reactions are localized at the plasma membrane or in the nucleus,
respectively. Finally, ADP-ribosyl cyclase, which converts
NAD+ to nicotinamide and cyclic ADP-ribose (cADPR)
(14-16), was first demonstrated as a membrane-bound enzyme in sea
urchin egg extracts (14). Various bifunctional enzymes in many
mammalian tissues can also catalyze both the synthesis and the
hydrolysis of cADPR. The first such enzyme reported was purified from
dog spleen (17). The ectocellular domain of CD38, a lymphocyte antigen
(18, 19), can also catalyze a two-step reaction involving transient
formation of cADPR (cyclase) followed by its hydrolysis to ADPR
(hydrolase), eventually generating nicotinamide and ADPR (20-24). The
overall reaction catalyzed by CD38 is thus identical to that catalyzed by the classical NAD+ glycohydrolases. The confusion
between CD38-like enzymes and the NAD+ glycohydrolases has
been recently resolved by the use of NGD+, an
NAD+ analog, as an alternative substrate. Only CD38-like
bifunctional enzymes possessing a cyclase activity can convert
NGD+ to cyclic GDP-ribose, a fluorescent product.
NAD+ glycohydrolases, on the other hand, can only convert
NGD+ to GDP-ribose, a non-fluorescent product (25).
In addition to CD38, another family of NAD+-utilizing enzymes has been recently identified that includes BST-1 (26) and BP-3 (27). These enzymes are glycosyl-phosphatidylinositol-anchored ectoproteins having considerable sequence homology to CD38 and may be involved in the ectocellular metabolism of NAD+, cADPR, and ADPR as well.
Various models have been proposed for the catalytic mechanism of cleavage of the nicotinamide-ribosyl linkage by NAD+-utilizing enzymes. A common feature among these models is the postulated presence of an enzyme-ADP-ribose intermediate (28-31). Depending on whether water is accessible to the active site, the intermediate can dissociate to yield either cADPR or ADPR, respectively, thus accounting for either a cyclase or an NAD+ glycohydrolase reaction (31). This catalytic scheme can also account for the base-exchange reaction actually observed. Thus, other nucleophiles including nicotinic acid can interact with the intermediate and generate products with the nicotinamide base exchanged for the nucleophiles. A physiologically relevant example of this type of base-exchange reaction is the synthesis of the new calcium-mobilizing molecule NAADP+ (32) from NADP+ and nicotinic acid, which is catalyzed by CD38 and ADP-ribosyl cyclase (31-34).
In this paper, we report another example of the exchange reaction that can lead to the formation of a hitherto unknown biologically active product. Using CD38 purified from human erythrocyte membranes, we show that ADP-ribose itself can function as a nucleophile and can interact with the enzyme intermediate. The result is the formation of a dimeric ADPR molecule. Of particular relevance is the fact that this ADPR dimer possesses functional activity, as it potentiates in a specific way the calcium-releasing activity of cADPR from sea urchin egg microsomal vesicles.
CD38 was purified to homogeneity from human erythrocyte membranes as described (21). Its cyclase specific activity, which was measured with NGD+ as a substrate (35), was 2.8 µmol cyclic GDP-ribose/min/mg. Its cADPR hydrolase-specific activity (35) was 2.8 µmol ADPR/min/mg. Production and purification of recombinant human CD38 was carried out as described previously (31, 36). Its cyclase specific activity on NGD+ was 18 µmol cyclic GDP-ribose/min/mg. ADP-ribosyl cyclase purified from Aplysia californica ovotestes as previously reported (31) had a specific activity that was measured using NAD+ as a substrate (35) of 20 µmol cADPR/min/mg. [3H]NAD+ (30.7 Ci/mmol) was obtained from DuPont NEN (Florence, Italy). All other chemicals were from Sigma and were of the highest purity grade available.
Analytical HPLCThe conditions for HPLC analyses using a C18 reverse phase column on a Hewlett-Packard HP 1090 instrument were as described previously (35). The nucleotide peaks were identified by comparing their retention times and UV spectra with highly pure standard compounds.
Synthesis and Purification of Dimeric ADPRPurified CD38
(0.75 µg) was incubated for 15 h at 37 °C with 1 mM NAD+ and 20 mM ADPR
(HPLC-purified) in 5 mM Tris-HCl, pH 6.5, containing bovine
serum albumin (0.1 mg/ml) and 0.05% Triton X-100. The final volume was
5.0 ml. The reaction was terminated by addition of trichloroacetic acid
(5% final concentration), the mixture was centrifuged, and the excess
trichloroacetic acid was removed with three successive extractions with
diethylether. The excess diethylether was evaporated under an
N2 stream, and the solution was vacuum-dried. The resulting
powder was dissolved in 0.5 ml of H2O, and 2 µl of the
solution was submitted to analytical HPLC (35). The yield of the
dimeric ADPR ((ADPR)2) eluted at 34 min was around 1% of the total area of all peaks. Therefore, to obtain sufficient amounts of
(ADPR)2 for analyses the incubation conditions described
above were repeated nine times. Each of the ten 0.5-ml samples thus obtained was then submitted separately to preparative HPLC. This was
carried out on a System Gold HPLC (Beckman Instruments) equipped with a
diode-array spectrophotometric detector set at 260 nm. An anion
exchange perfusion chromatography column (100 × 4.6 mm, Poros
QE/M, PerSeptive Biosystems, Cambridge, MA) was used. Solvent A was
H2O, and solvent B was 0.15 M trifluoroacetic
acid. The solvent program was a linear gradient starting at 100%
solvent A and increasing to 100% solvent B in 15 min. The flow rate
was 6 ml/min. (ADPR)2 eluted at 7 min, well separated from
all other nucleotides (NAD+, ADPR, and cADPR) and
nicotinamide, which eluted in the same peak between 2 and 5 min. The
7-min peak was lyophilized and stored at 20 °C until used.
Labeled (ADPR)2
was obtained by incubating purified CD38 (30 ng) at 37 °C with 20 mM ADPR, 1 mM
[3H]NAD+ (5000 dpm/nmol) in 5 mM
Tris-HCl, pH 6.5, with 0.05% Triton X-100 in a final volume of 0.2 ml.
After an 18-h incubation period, aliquots of 100,000 dpm were submitted
to analytical HPLC (35), fractions were collected every minute, and the
radioactivity values were determined in a -counter. The
HPLC-purified labeled (ADPR)2 (25 µg, corresponding to
5,000 dpm) was then incubated for 18 h at 37 °C with each of
the following enzymes separately: (i) 5 µg NAD+
glycohydrolase from Neurospora crassa in 65 µl of
phosphate-buffered saline; (ii) 32 ng of ADP-ribosyl cyclase purified
from A. californica (31) in 65 µl of Tris-HCl, pH 6.5, containing 0.05% Triton X-100; (iii) 30 ng of purified CD38 (21) in 65 µl of Tris-HCl, pH 6.5, containing 0.05% Triton X-100. Residual
(ADPR)2 and neo-synthesized ADPR were evaluated by
analytical HPLC (35).
Purified (ADPR)2 (0.25 mg)
was dissolved in H2O and incubated for 18 h at
37 °C with 30 µg of commercial dinucleotide pyrophosphatase (EC
3.6.1.9) from Crotalus adamanteus venom in 0.2 ml of 20 mM Tris-HCl, pH 8.3, containing 2 mM
MgCl2. The reaction was terminated with trichloroacetic
acid (5% final concentration), and the trichloroacetic acid was
removed as described above. Part of the deproteinized mixture (2%) was
analyzed by HPLC (35) to monitor the reaction products. Two products
were formed, AMP (retention time, 15 min) and an unknown product that
eluted at 25 min (hence designated P25). The remaining 98% of the
deproteinized mixture was submitted to anion exchange perfusion
chromatography as described above except that a flow rate of 4 ml/min
was used instead. The unknown compound (P25) that eluted at 7 min was
well separated from AMP, which had a retention time of 3 min in this
system. Purified P25 was lyophilized and stored at 20 °C.
Lyophilized P25 (0.1 mg) was dissolved in H2O and incubated
for 15 h at 37 °C with 15 µg of 5-nucleotidase (EC 3.1.3.5) from C. adamanteus venom in 0.25 ml of phosphate-buffered
saline. The reaction was terminated with trichloroacetic acid (5%
final concentration), the trichloroacetic acid was removed as described above, and the resulting solution was analyzed by HPLC (35), which
showed only one peak with a retention time of 12 min (P12). This peak
was purified by HPLC using C18 reverse phase chromatography (ODS
Hypersil 3 µm, 60 × 4.6 mm) at a flow rate of 0.5 ml/min. Solvent A was deionized water, and solvent B was methanol (HPLC grade).
The solvent program was a gradient starting at 100% Solvent A for 5 min, then linearly increasing to 100% Solvent B in 15 min. Under these
HPLC conditions, the P12 peak showed a retention time of 12 min. This
peak was collected, lyophilized, and stored at
20 °C before
HPLC/MS analyses.
Various purified samples were resuspended at 5-50 µg/ml in H2O:methanol:trifluoroacetic acid (49.5:49.5:1). The sample was pumped by an HPLC pump (Kontron Instruments, Milan, Italy) through a Valco valve into the mass spectrometer (Hewlett Packard 5989A Engine, Palo Alto, CA) electrospray source. Background spectra were obtained by injecting the solvent without the samples and were subtracted from the averaged spectra of various samples. Data were collected in the positive ion mode in a range including the expected molecular masses to avoid the influence of the sodium and potassium adducts usually found with phosphate-containing molecules.
Conversion of cADPR to N1-(5The procedure was as
described by Gu and Sih (37) with minor modifications. Briefly, sodium
tert-butoxide was prepared by dissolving 0.3 g of
sodium in 13 ml of tert-butyl alcohol overnight at room
temperature. The solution was vacuum-dried, and the resulting white
powder was dissolved in an Me2SO:water (998:2) solution to
give a final concentration of 0.4 mg/ml. cADPR (30 µg) was then
incubated for 60 min at 35 °C in 1.0 ml of the
Me2SO:tert-butoxide mixture. Under these
conditions, the main product observed by analytical HPLC (35) had a
retention time of 25 min, identical to that obtained upon incubation of
(ADPR)2 with dinucleotide pyrophosphatase (P25, see above).
This product was purified by anion exchange perfusion chromatography
(see above), lyophilized, and stored at 20 °C before being
submitted to MS analysis. Prolonging the incubation of cADPR in
Me2SO:tert-butoxide to 2 h resulted in
greatly reduced formation of the peak that eluted at 25 min and in the
concomitant appearance of another compound that had a retention time of
12 min. This is identical to the retention time of the product obtained
by sequential digestion of (ADPR)2 with dinucleotide
pyrophosphatase and 5
-nucleotidase (P12, see above).
Fractionation of sea urchin egg (Strongylocentrotus purpuratus) microsomes was performed as described in Ref. 38 except that calmodulin (10 µg/ml) was added to the dilution medium. Ca2+ release assays were performed at 17 °C under constant stirring.
Fig. 1 shows that native CD38 purified
from human erythrocyte membranes catalyzes the base-exchange reaction
resulting in the synthesis of NAAD+ from NAD+
and nicotinic acid. The reaction occurs even at pH 6.5, which is
significantly higher than that previously demonstrated to be optimal
for the base-exchange reaction catalyzed by a recombinant CD38
containing only the extracellular domain (31). The extent of
NAAD+ formation was comparable with that catalyzed by
Aplysia cyclase also shown in Fig. 1. In addition to
NAAD+, erythrocyte CD38 also results in the synthesis of a
modest amount of cADPR and of a substantial amount of ADPR. In
contrast, the Aplysia cyclase mainly catalyzes the formation
of cADPR with minor production of NAAD+ and ADPR (Fig.
1).
ADP-ribosyl Cyclase and CD38 Catalyze the Formation of an Unknown Product from NAD+ and ADPR
Whether ADPR can serve as
a nucleophile in the base-exchange reaction was investigated by
incubating the native CD38 with [adenine-3H]NAD+ in the presence
of excess ADPR. Fig. 2 shows that the majority of the
3H label was converted to ADPR (eluted at 23 min) with a
small portion of it going to cADPR (eluted at 5.5 min). In addition, a
detectable amount of the label (around 1%) was present in an unknown
peak eluted at 34 min (P34). This peak was also labeled when
[-32P]NAD+ was used as traced instead of
[3H]NAD+ (not shown). The UV spectrum of the
unknown peak shown in the inset of Fig. 2 is clearly
different from that of ADPR, with a maximum at 270 nm rather than at
260 nm.
Fig. 3 shows that the formation of P34 from
[3H]NAD+ and ADPR is catalyzed not only by
CD38 but also by ADP-ribosyl cyclase from A. californica.
The time course of P34 production was similar with the two enzymes. The
amount of P34 formed by CD38 at 18 h was only slightly lower than
that of cADPR, indicating that it is a major product of CD38 (Fig.
3A). The long incubation time was chosen in these
experiments because CD38 is known to be substantially inactivated due
to the NAD+-induced self-aggregation of the protein (35,
39, 40). This inactivating effect as well as the attendant
oligomerization was also observed with a recombinant form of human CD38
corresponding to its ectocellular C-terminal region (35, 36). The
NAD+-dependent inactivation of CD38 is
different from that recently observed with NAD+
glycohydrolase purified from rabbit erythrocytes (41), which is due to
auto-ADP-ribosylation and is reversible.
Since the two-step activity of CD38 leads to accumulation of ADPR as
the major end product (Fig. 3A), we investigated whether P34
production also occurs without any initial addition of ADPR as
co-substrate. As shown in Fig. 4, this proved to be the
case, although formation of P34 became detectable only at 8 h of
incubation, when production of ADPR peaked at approximately 5 mM and was comparably lower in extent than that recorded
with both NAD+ and ADPR added. In these experiments, the
NAD+-induced inactivation of CD38 was counteracted by
repeated additions of the soluble recombinant protein. At the same
times, the incubation mixtures were also regularly supplemented with
NAD+ to provide enough substrate for the reactions
catalyzed by CD38 to take place. Indeed, under these conditions the
disappearance of NAD+ over the extended incubation times
went to completion differently from the experiment shown in Fig.
3A in which NAD+ accumulated because of enzyme
inactivation (not shown).
Generation of P34 and cADPR was also observed at much lower initial
concentrations of NAD+. In these experiments also, no ADPR
was added, and the starting concentration of NAD+ was 10 µM (Fig. 5). The amount of CD38 was
remarkably lower than in the experimental conditions of Fig. 4 to avoid
fast exhaustion of NAD+. Again, both 10 µM
NAD+ and fresh recombinant CD38 were added to the
incubation mixtures at sequential time intervals to allow the enzymatic
reactions to proceed continuously. As shown in Fig. 5, even at these
physiological concentrations of NAD+, the formation of P34
progressed slowly and was only slightly lower than that of cADPR.
In an attempt to address the physiological occurrence of P34 formation from NAD+, experiments were carried out on intact Namalwa cells (a continuous B-cell-derived line from Burkitt's lymphoma), which have both enzymatic activities of CD38 at their outer surface (42) rather than on purified CD38. Using these cells at the same activity levels as purified recombinant CD38 on 10 µM NAD+ (Fig. 5), disappearance of NAD+ and concomitant progressive formation of ADPR, cADPR, and P34 were apparently observed at short time intervals as low as 5-30 min (not shown). In these experiments, however, unequivocal identification of the nucleotide peak eluted at 34 min with authentic P34 was not possible on the basis of UV spectrum because amounts of this peak were too limited. Moreover, the picture of NAD+ metabolism in Namalwa cells was remarkably complicated by additional products of ADPR catabolism including AMP, adenosine, and hypoxanthine, which were generated partly at the cell surface and partly intracellularly.
Attempts were then made to establish optimal conditions of P34 formation. In these experiments we used the general setting shown in Fig. 3, i.e. a single addition of CD38 (either purified from erythrocyte membranes or recombinant) and one of NAD+. However, even in these simplified conditions, variations of substrate concentrations (both NAD+ and ADPR) as well as of the pH of the reaction produced no further improvement. Likewise, P34 formation was not enhanced upon performing the incubations in more apolar media such as 6 M methanol to minimize hydrolytic reactions. Control experiments showed that the formation of P34 requires catalysis since it was undetectable in the absence of enzyme.
Structural Analysis of P34Various enzymes were used to convert P34 into known products (see "Experimental Procedures"). P34 proved not to be a substrate of commercial NAD+ glycohydrolase or Aplysia ADP-ribosyl cyclase. On the contrary, P34 was quantitatively converted to ADPR by purified CD38, and no other UV-absorbing or labeled compounds were produced in these conditions. This finding indicates P34 is likely a dimer or an oligomer of ADPR.
Mass spectrometry measurements showed that the molecular ion of P34,
(M+H)+, has a mass of 1,101. This value corresponds to two
individual ADPR molecules joined together with the loss of a water
molecule (Fig. 6A). P34 can, therefore, be
designated as (ADPR)2. Incubation of (ADPR)2
with dinucleotide pyrophosphatase cleaved the pyrophosphate bond of
ADPR, yielding two degradation products. One was identified as AMP by
its retention time of 15 min. The other product was designated as P25
since it eluted at 25 min. The m/z value of P25
was 560, which is consistent with an adenine moiety linked to two
ribose-phosphate units, i.e. A(R-5-P)2 as shown
in Fig. 6B. Further incubation of A(R-5-P)2 with
5-nucleotidase led to the formation of a single compound with a
retention time of 12 min (indicated as P12 in Fig. 6C). P12
had an m/z value of 400 corresponding to an
adenine bonded to two ribose moieties (AR2).
Further insight into the structure of (ADPR)2 was provided
by parallel experiments in which the pyrophosphate linkage of authentic cADPR was cleaved under conditions not perturbing the integrity of its
C-N1 glycosyl bond. The pyrophosphate linkage of cADPR is
markedly resistant to a number of pyrophosphatases, both from
commercial sources and present in human plasma (not shown). However, it
can be chemically broken by exposure to tert-butoxide in
Me2SO. The resulting product is
N1-(5-phosphoribosyl)-AMP, a compound in which both
N1 and N9 are bound to two anomeric carbons of
ribose 5
-P (37). If the C-N1 glycosyl bond were involved
in joining the two ADPR moieties of (ADPR)2 together, then
the same molecule of N1-(5
-phosphoribosyl)-AMP
(i.e. the compound referred to as A(R-5-P)2 in
Fig. 6B) should arise from the digestion of
(ADPR)2 by dinucleotide pyrophosphatase. Indeed, the
degradation procedures employed for cADPR and (ADPR)2 both
yielded a compound with identical retention time (25 min using a
reverse phase column as described under "Experimental Procedures"),
the same UV spectrum, and also the same mass of 560 for its molecular
ion (M+H)+.
Prolonging the incubation of cADPR in
Me2SO:tert-butoxide for 2 h instead of
1 h further degraded the molecule into a product having a
retention time of 12 min on a reverse phase column and a mass of 400 for its molecular ion, (M+H)+. These properties coincide
with those of the product obtained upon digesting P34 sequentially with
dinucleotide pyrophosphatase and 5-nucleotidase (Fig. 6C).
These results suggest the same type of N1-glycosyl linkage
that occurs in cADPR is present also in (ADPR)2. Taken
together, these data are therefore consistent for the molecular structure shown in Fig. 7, in which the same adenine
moiety binds two anomeric carbons of the two ADP-ribosyl units at
N1 and N9, respectively.
Functional Activity of (ADPR)2
Fig.
8A shows that addition of (ADPR)2
to reach a final concentration as high as 50 µM produces
no detectable Ca2+ release from sea urchin egg microsomes.
For comparison, as low as 20 nM cADPR elicits detectable
Ca2+ release in the same microsomal preparation (not
shown). However, when increasing amounts of (ADPR)2 were
added to microsomes pretreated with 15 nM cADPR, which was
too low to produce Ca2+ release by itself, a
concentration-dependent release of Ca2+ was
observed reaching its maximum at about 50 µM. This
synergistic effect was half-maximal at 25-30 µM
(ADPR)2 (Fig. 8A). Ca2+ stores were
not emptied by the combination of 50 µM
(ADPR)2 and 15 nM cADPR. Indeed, higher
concentrations of cADPR alone (up to 200 nM) could induce a
further Ca2+ release from the microsomes.
Fig. 8B shows the converse experiment when the egg microsomes were pretreated with a 50 µM concentration of (ADPR)2, followed by various concentrations of cADPR. The pretreatment sensitized the microsomes such that only 18 nM of cADPR was sufficient to induce near-maximal Ca2+ release from the microsomes. When this experiment was performed at low constant concentrations of (ADPR)2 (10 and 25 µM, respectively), a potentiating effect on subthreshold concentrations of cADPR was still observed, although the effect was less evident than with 50 µM (ADPR)2. These results indicate that (ADPR)2 can function as a sensitizer of cADPR.
The amplifying effect of (ADPR)2 on cADPR activity proved to be specific. Thus, ADPR and its potential metabolites AMP (43) and adenosine, each at concentrations up to 100 µM, failed to release Ca2+ from sea urchin egg microsomes and to display any synergistic effect on cADPR (not shown).
The involvement of an enzyme-ADP-ribose intermediate in the
catalytic mechanism of monofunctional ADP-ribosyl cyclase, bifunctional cyclases/hydrolases, or NAD+ glycohydrolases has been
suggested by several lines of evidence including: (i) methanolysis of
NAD+ (29, 30, 44), (ii) base-exchange reaction between
nicotinamide and nicotinic acid (31), (iii) NAD+ synthesis
from cADPR and nicotinamide (17), and (iv) the conformation of the
N1-glycosyl linkage of cADPR being in the configuration
in both its crystal form (45) and in aqueous solution (37, 46). The
data reported in this paper are also consonant with this mechanism because the dimeric structure of (ADPR)2 implies the
transfer of an enzyme-activated ADP-ribosyl moiety to a pre-existing
ADPR molecule behaving as an acceptor substrate.
Fig. 9 illustrates a working model accounting for the
versatility of CD38 in terms of catalyzing the formation of multiple products. As already reported (17, 31, 47), the ADP-ribosyl-enzyme intermediate can react either with water, resulting in generation of
ADPR, or with the N1 of the adenine moiety, leading to
production and release of cADPR. The third possible pathway is closely
reminiscent of the previously reported mechanism of base-exchange
leading to the synthesis of NAAD+ or of NAADP+
in the presence of nicotinic acid (31, 34). In the scheme shown in Fig.
9, the exchange occurs between nicotinamide and the preformed ADPR,
thereby producing the dimeric ADPR molecule identified in this study.
(ADPR)2 itself is a good substrate of the hydrolase
activity of CD38. This fact may be responsible for the low amounts of
(ADPR)2 detected in our experiments. Also, it is possible
that exceedingly high concentrations of ADPR in the proximity of the
active site are needed to compete with the intramolecular cyclization
that results in cADPR synthesis. This can account for the similarly low
production of (ADPR)2 by the Aplysia cyclase
despite its weak hydrolase activity as compared with CD38. Although
less extensive, formation of (ADPR)2 was also observed upon
incubating CD38 with NAD+ in the absence of added ADPR
(Figs. 4 and 5). Nevertheless, since (ADPR)2 became
detectable only when ADPR reached its maximal concentrations, this
finding provides further confirmation of the exchange mechanism shown
in Fig. 9. Moreover, it demonstrates that the reaction of CD38 on
NAD+ can ultimately generate cADPR, ADPR, and
(ADPR)2 and that the two last compounds can be reciprocally
interconvertible.
The formation of (ADPR)2 requires enzymatic catalysis since it is not detectable under our experimental conditions without enzyme. Nevertheless, a compound sharing the same mass as the enzymatically prepared (ADPR)2 was observed as a 0.1% impurity of commercial ADPR preparations. This impurity also co-eluted with 3H-labeled (ADPR)2 that was synthesized by CD38. In all experiments reported in this study, the impurity of the commercial ADPR was removed by repurification using HPLC. A similar situation has been described for NAADP+, which has also been shown to be present as an impurity in commercial NADP+ preparations. Also similar to (ADPR)2 is that NAADP+ can be synthesized either chemically by alkaline treatment of NADP+ (32) or, as recently reported, by a base-exchange reaction catalyzed by CD38 using NADP+ and nicotinic acid as substrates (31).
In this study, specific patterns of enzymatic degradation combined with
MS analyses of the products were used to unequivocally establish the
dimeric structure of (ADPR)2. Evidence for the
N1-ribosyl bond as the linkage responsible for joining the
two ADPR units together is taken from the following two findings: (i)
the susceptibility of (ADPR)2 to be hydrolyzed by CD38;
(ii) the analyses of the degradation products of the digestion of
(ADPR)2 with dinucleotide pyrophosphatase either alone or
in combination with 5-nucleotidase that show that they are the same as
those produced following chemical cleavage of the pyrophosphate bond of
cADPR. Since the cyclizing linkage in cADPR has been unambiguously
shown to be the N1-ribosyl bond by x-ray crystallography
(45), the identity of the degradation products of (ADPR)2
and cADPR provides convincing evidence that the linkage in
(ADPR)2 is also the N1-ribosyl bond.
(ADPR)2 itself is devoid of Ca2+-releasing activity on sea urchin egg microsomal vesicles. However, it synergizes with cADPR in producing an enhancement of the Ca2+-releasing activity of subthreshold concentrations of cADPR (Fig. 8B). This effect is specific for (ADPR)2 as it was not observed with ADPR or with known metabolites of ADPR including AMP (43) and adenosine. The Ca2+-releasing activity of cADPR has been previously shown to be potentiated by calmodulin and divalent cations (38, 48) and also by sensitizers of the Ca2+-induced Ca2+ release such as caffeine (49). The potentiation effect of (ADPR)2 represents yet another way of modulating the Ca2+ release mechanism activated by cADPR.
The amount of (ADPR)2 produced by incubation of CD38 with NAD+, either with or without added ADPR, is low. It can be, nevertheless, physiologically significant since it is comparable with the amounts of cADPR produced under the same conditions (Figs. 3, 4, 5). Moreover, generation of (ADPR)2 was also detectable starting from the reaction of recombinant CD38 on NAD+ concentrations as low as 10 µM (Fig. 5). Use of intact Namalwa B cells as a source of ectocellular CD38 (42) instead of purified CD38 also supported the notion that 10 µM NAD+ acts as a precursor of (ADPR)2 as well as of cADPR and ADPR.
Previously unrecognized formation of (ADPR)2 might account for regulation of Ca2+-releasing activity of cADPR in biological systems. Recently, ADPR has been reported to potentiate cADPR synthesis from NAD+ in sea urchin egg homogenates (50), and this effect was ascribed to an inhibitory mechanism afforded by ADPR on the cADPR hydrolase activity resulting in elevated cADPR levels. Since cADPR concentrations were evaluated using a bioassay based on Ca2+-releasing activity from sea urchin egg homogenates, an alternative explanation could be the formation of (ADPR)2 thus potentiating the Ca2+ release induced by cADPR. Indeed both mechanisms, i.e. inhibition of cADPR hydrolase and formation of (ADPR)2, could result in the observed effect of potentiation of Ca2+-releasing activity in sea urchin egg homogenates (50). Accordingly, (ADPR)2 might be involved in up-modulating the biological effect of cADPR in this system.
The ectocellular localization of the catalytic domain of CD38 suggests that its enzymatic products may have extracellular functions. It has been previously shown that cADPR applied extracellularly to B lymphocytes can enhance the cell proliferation stimulated by co-agonists, while ADPR has the opposite effect (20). Recently, transient exposure of rat cerebellar granule cells to extracellular cADPR has been observed to enhance the Ca2+-induced Ca2+ release activity of these cells in approximately 50% of the experiments (51). Conversely, this functional effect was an almost constant finding when the same CD38+ cerebellar granule neurons were exposed to extracellular NAD+ as a precursor of cADPR. Whether this difference is due to (ADPR)2 formation from NAD+ potentiating the effect of cADPR is an attractive possibility that is currently under study.
(ADPR)2 adds to a growing number of metabolites generated
by the enzymes involved in the metabolism of cADPR. Depending on the
presence of NAD+ or NADP+ as a substrate,
cADPR, NAADP+, and 2-P-cADPR can be produced under
different experimental conditions. All of these have been shown to have
Ca2+-releasing activity (14, 32, 33, 52). Degradation
pathways for these active metabolites have also been described. Thus,
cADPR is known to be hydrolyzed by CD38-like enzymes to ADPR, while alkaline phosphatase has been shown to be effective in hydrolyzing 2
-P-cADPR to cADPR (31) and NAADP+ to the functionally
inactive NAAD+ (53). Importantly, we observed that CD38 is
able to not only synthesize (ADPR)2 but also to hydrolyze
it, suggesting that the concentration of this nucleotide in cells may
be regulated. It is conceivable that a small augmentation of the
synthesizing activity of CD38 coupled with a corresponding decrease in
the hydrolyzing activity can result in a larger increase of the steady
state concentration of (ADPR)2. Indeed, it has been shown
that Zn2+ can modulate the cADPR synthesizing activity of
CD38 in precisely this manner (21, 54).
This proliferation of interacting metabolites of this signaling pathway is reminiscent of the phosphatidylinositide pathway, which began with the discovery of inositol trisphosphate but soon grew with the multiplicity of interacting inositol phosphates (reviewed in Ref. 55). In this context, (ADPR)2 can be viewed as a product of the signaling pathway designed to amplify the physiological effects of cADPR. In any case, discovery of (ADPR)2 as an end product of CD38 enzymatic activities does raise many intriguing possibilities demanding further investigation.
We thank Dr. R. M. Graeff for critical reading of this manuscript.