From the Department of Biochemistry, Shimane Medical University, 89-1, Izumo, Shimane 693-8501, Japan
Received for publication, September 9, 2002, and in revised form, December 5, 2002
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ABSTRACT |
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NAD synthetase catalyzes the final step in the
biosynthesis of NAD. In the present study, we obtained cDNAs for
two types of human NAD synthetase (referred as NADsyn1 and NADsyn2).
Structural analysis revealed in both NADsyn1 and NADsyn2 a domain
required for NAD synthesis from ammonia and in only NADsyn1 an
additional carbon-nitrogen hydrolase domain shared with enzymes of the
nitrilase family that cleave nitriles as well as amides to produce the
corresponding acids and ammonia. Consistent with the domain
structures, biochemical assays indicated (i) that both NADsyn1 and
NADsyn2 have NAD synthetase activity, (ii) that NADsyn1 uses
glutamine as well as ammonia as an amide donor, whereas NADsyn2
catalyzes only ammonia-dependent NAD synthesis, and (iii)
that mutant NADsyn1 in which Cys-175 corresponding to the catalytic
cysteine residue in nitrilases was replaced with Ser does not use
glutamine. Kinetic studies suggested that glutamine and ammonia serve
as physiological amide donors for NADsyn1 and NADsyn2, respectively.
Both synthetases exerted catalytic activity in a multimeric form. In
the mouse, NADsyn1 was seen to be abundantly expressed in the small
intestine, liver, kidney, and testis but very weakly in the skeletal
muscle and heart. In contrast, expression of NADsyn2 was observed in all tissues tested. Therefore, we conclude that humans have two types
of NAD synthetase exhibiting different amide donor specificity and
tissue distributions. The ammonia-dependent synthetase has not been found in eucaryotes until this study. Our results also indicate that the carbon-nitrogen hydrolase domain is the functional domain of NAD synthetase to make use of glutamine as an amide donor in
NAD synthesis. Thus, glutamine-dependent NAD synthetase may
be classified as a possible glutamine amidase in the nitrilase family.
Our molecular identification of NAD synthetases may prove useful to
learn more of mechanisms regulating cellular NAD metabolism.
The coenzyme NAD has a role in the majority of metabolic redox
reactions and represents an essential component of metabolic pathways
in all living cells. In a number of signaling pathways, NAD also serves
as a precursor of potent calcium-mobilizing agents such as cyclic
ADP-ribose and nicotinic acid adenine dinucleotide phosphate (1) and
serves as a substrate for post-translational modifications of protein,
mono- (2-4) and poly(ADP-ribosyl)ations (5). Depletion of cellular NAD
by poly(ADP-ribosyl)transferase activation in response to DNA damage
results in cell death (6). Increased NAD synthesis has been shown to
extend life span in yeast (7) and in Caenorhabditis elegans
(8) via activation of an NAD-dependent histone deacetylase,
silent information regulator 2 (Sir2) (9). The cellular level of NAD
may modulate the sensitivity of cells to apoptotic responses through
deacetylation of the p53 tumor suppressor by a human homologue of Sir2
(10). Recent publications have demonstrated that fluctuation of the NAD
level in cells seems to have significant impact on their physiology.
Despite these significant effects of NAD levels on cellular functions,
mechanisms regulating cellular contents of NAD through metabolic events
remain to be established.
NAD biosynthesis is accomplished through either de novo or
salvage pathways (11, 12). These two pathways converge at the level of
an intermediate nicotinic acid mononucleotide
(NaMN),1 which is then
converted into nicotinic acid adenine dinucleotide (NaAD) through the
action of NaMN adenylyltransferase and, lastly, into NAD by NAD
synthetase (Fig. 1). Although most of the
genes involved in both pathways have been identified in procaryotes (13), little is known of those genes, including that of NAD synthetase
in eucaryotes, except for nicotinamide mononucleotide adenylyltransferase (14) and quinolinic acid phosphoribosyltransferase (15) genes.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Metabolic pathways of NAD biosynthesis.
Nam, nicotinamide; NMN, nicotinamide
mononucleotide; NA, nicotinic acid; QA,
quinolinic acid; NAPRT, nicotinic acid
phosphoribosyltransferase; NaMNAT, nicotinic acid
mononucleotide adenylyltransferase; NADsyn, NAD synthetase;
QPRT, quinolinic acid phosphoribosyltransferase.
NAD synthetase catalyzes the conversion of NaAD into NAD, and
NH3 or glutamine is used as an amide donor in the following reactions.
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We now report molecular identification of two human NAD synthetases, a
synthetase that can use not only ammonia but also glutamine and the
other synthetase with strictly ammonia-dependent activity (referred as NADsyn1 and NADsyn2, respectively). To our knowledge, this
is the first report demonstrating the presence of the strictly ammonia-dependent NAD synthetase in eucaryotes. We also
describe the structural basis underlying the potential of NADsyn1 to
use glutamine as an amide donor as well as the distinct distribution of
NADsyn1 and NADsyn2 in animal tissues.
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EXPERIMENTAL PROCEDURES |
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Materials--
[-32P]dCTP (6000 Ci/mmol) was
purchased from Amersham Biosciences. NaAD, AMP and inorganic
pyrophosphatase were from Sigma. ATP, NAD, L-glutamine, and
ammonium chloride were from Oriental Yeast (Tokyo, Japan), Roche
Molecular Biochemicals (Basel, Switzerland), Nacalai Tesque (Kyoto,
Japan), and Wako Pure Chemical Industries (Osaka, Japan), respectively.
COS-7 cells and a human promyelocytic leukemia cell line HL60 were
obtained from Riken Cell Bank (Tsukuba Science City, Japan). Human
glioma cell line LN229 and human hepatocyte cell lines HepG2 and Huh7
were from American Type Culture Collection (Manassas, VA).
Expression of NADsyn1 and NADsyn2 in COS-7 Cells-- To express NADsyn1 and NADsyn2 as C-terminal-His6-tagged proteins in COS-7 cells, a His6 tag sequence followed by a TGA termination codon was introduced into the pcDNA3 vector (Invitrogen) between XbaI and ApaI cloning sites to obtain the pcDNA3His6 vector. Segments of human NADsyn1 cDNA were PCR-amplified from fetal human brain cDNA (Clontech, Palo Alto, CA) using two sets of primers, 5'-ATG GGC CGG AAG GTG ACC-3' (sense) and 5'-CAG ACC TGG CAG CAC ATG-3' (antisense) and 5'-AAG CCT TGG ACC TGC CTG-3' (sense) and 5'-GAA GGA ACC GGC CTC A-3' (antisense). The PCR products were gel-purified, combined, and amplified using primers (the underlined regions correspond to cloning sites) 5'-AAG CTT GGT ACC ATG GGC CGG AAG GTG ACC-3' (sense) and 5'-AAG CTT TCT AGA GTC CAC GCC GTC CAG GGA-3' (antisense), yielding full-length NADsyn1 cDNA. Human NADsyn2 cDNA was amplified from LN229 total RNA treated with deoxyribonuclease I (Nippon Gene, Tokyo, Japan) by reverse transcription-PCR using primers 5'-AAG CTT GGA TCC ATG CAA GCC GTA CAG CGC-3' (sense) and 5'-AAG CTT TCT AGA GGG TGC AAA CGG CAT CAC-3' (antisense). Amplified cDNAs were digested with KpnI and XbaI for NADsyn1 and BamHI and XbaI for NADsyn2 and ligated, respectively, into KpnI- and XbaI-digested and BamHI- and XbaI-digested pcDNA3His6. The vector used to express the mutant NADsyn1, in which Cys-175 was replaced with serine (C175S-NADsyn1), was made using a QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) using the pcDNA3His6 plasmid vector carrying wild-type NADsyn1 and the oligonucleotide primers 5'-TT GGA AGT GAG ATC agt GAG GAG CTC TGG-3' and 5'-CCA GAG CTC CTC act GAT CTC ACT TCC AA-3', where the altered codon is indicated in lowercase italics. The resultant expression plasmids, purified using a Qiagen plasmid kit (Hilden, Germany), were transfected into COS-7 cells (4.5-6.5 × 105 cells/100-mm dish) using an activated-dendrimer PolyFect Transfection Reagent (Qiagen) according to the manufacturer's instructions. Forty-eight hours after transfection, the COS-7 cells were washed twice with phosphate-buffered saline and collected by scraping. After the cells were lysed by sonication and centrifugation, recombinant NAD synthetases were purified from the supernatants with His-Bind Resin (Novagen, Madison, WI) according to the manufacturer's protocol. A cDNA fragment of mouse glutamine-dependent NAD synthetase was amplified from Balb/c mouse kidney total RNA treated with deoxyribonuclease I by reverse transcription-PCR using primers based on the sequence of mouse homologue of human NADsyn1 (see Fig. 7),5'-ATG GGC CGG AAA GTG ACC-3' (sense) and 5'-CAG ACC AGG CAG CAC ATG-3' (antisense). Sequences of expression plasmids or PCR fragments were confirmed by entire sequencing in both directions.
5'-Rapid Amplification of cDNA Ends-- Adaptor-ligated double strand cDNA was synthesized using Marathon cDNA amplification kit (Clontech) from poly(A)+ RNA isolated from HL60 cells with QuickPrep micro mRNA purification kit (Amersham Biosciences). The 5' parts of the NADsyn1 and NADsyn2 cDNAs were amplified with Advantage 2 polymerase mix (Clontech) using sense adaptor primer 5'-CCA TCC TAA TAC GAC TCA CTA TAG GGC-3' and gene-specific antisense primers 5'-CAG CGC AGC TCG CGG TAG TTG CCT TC-3' (NADsyn1) and 5'-CCC AGC ACG AAG TCC CGG GAG ACT GC-3' (NADsyn2). After nested PCR, the products were subcloned into pcDNA3, and positive clones were isolated and sequenced.
Enzyme Assays--
Unless otherwise stated, NAD synthetase
activity of the recombinant protein was based on fluorometric
measurements of the NAD formed, as described below. Recombinant
NADsyn1, the mutant NADsyn1, and NADsyn2 were incubated with glutamine
or NH4Cl as indicated in the reaction mixture (50 µl)
containing 50 mM HEPES (pH 8.8), 2 mM ATP, 1 mM NaAD, 56 mM KCl, 5 mM
MgCl2, and 10 µg of bovine serum albumin. For the assay
of NADsyn1 activity, 2 mM dithiothreitol was included in
the reaction mixture. When glutamine was used as a substrate for
NADsyn1, 50 mM Tris-Cl (pH 7.5) was included
instead of 50 mM HEPES (pH 8.8). The reactions were
terminated by adding 0.4 ml of 7 N NaOH and then incubated at 37 °C for 30 min to obtain the fluorescent product (21). The
fluorescence was measured using 380 nm for excitation and 460 nm for
emission by Fluoroskan Ascent FL (Labsystems, Helsinki, Finland). The
fluorescence intensity of standard NAD solutions at known
concentrations was used to calculate the amount of NAD. NAD synthetase
activity was calculated by subtracting the NAD content of
enzyme-deficient blanks from the NAD content of the complete reaction mixture.
In some cases NAD synthetase activity was determined by HPLC analysis. After the NAD synthetase reactions had been terminated by a 10-fold dilution with 0.1% trifluoroacetic acid, the reaction products (NAD, NaAD, and AMP) were separated on a reversed phase Cosmosil 5C-18MS column (4.6 × 150 mm, Nacalai Tesque) with 0.1% trifluoroacetic acid as the mobile phase and detected by measuring absorbance at 254 nm.
Kinetic parameters for NAD synthetase reaction were determined as follows by analysis of a Lineweaver-Burk plot of the initial rates of NAD synthesis. In the reaction mixture at fixed concentrations of two of the three substrates concentrations of the third substrate varied from 0.1 to 2 mM for NaAD, from 0.05 to 1 mM for ATP, from 1 to 40 mM for glutamine, and from 1.5 to 30 mM (NADsyn1 and C175S-NADsyn1) or from 0.01 to 0.2 mM (NADsyn2) for NH4Cl. In the reaction catalyzed by NADsyn1, Km values for NaAD and ATP were determined using glutamine as an amide donor, whereas in reactions catalyzed by NADsyn2 and C175S-NADsyn1, values were determined using NH4Cl. Amounts of NAD formed were determined by the fluorometric method.
Determination of PPi--
NAD synthetase reactions
(50 µl) were terminated by adding 8 µl of 10% trifluoroacetic
acid. After standing on ice for 15 min and then neutralizing by 1 M Tris-Cl (pH 9.0), the reaction mixtures
were incubated with or without 2.5 milliunits/µl of pyrophosphatase
at 25 °C for 30 min. The reactions were terminated by 10%
perchloric acid, and the inorganic phosphate formed was determined as
described by Yu and Dietrich (19).
Determination of Molecular Masses of Catalytically Active NAD Synthetases-- Purified recombinant NADsyn1 and NADsyn2 were electrophoresed on 7% and 12.5% non-denaturing polyacrylamide gels, respectively, in Tris-glycine buffer (pH 8.3) (25 mM Tris, 192 mM glycine) in the presence of 0.2% 2-mercaptoethanol. Gels were sliced into 2-3-mm pieces then incubated in the reaction mixture (100 µl) containing either 20 mM glutamine for NADsyn1 or 1 mM NH4Cl for NADsyn2 at 37 °C for 2 h. NAD formed in the reaction mixture was determined using the fluorometric method, as described above.
Northern Blot Analysis--
Total RNAs (20 µg) prepared from
Balb/c mouse tissues and human cell lines were fractionated on a 1%
agarose-formaldehyde gel, transferred to Hybond-N+ nylon membrane
(Amersham Biosciences), and UV-cross-linked. The blot was prehybridized
at 43 °C for 5 h in hybridization solution containing 5× SSPE
(1× SSPE: 0.15 M NaCl, 8.65 mM sodium
dihydrogen phosphate, 1.25 mM EDTA), 50% formamide, 5×
Denhardt's solution (1× Denhardt's solution: 0.02% Ficoll, 0.02%
polyvinylpyrrolidone, 0.02% bovine serum albumin), 0.5% SDS, and 100 µg/ml heat-denatured salmon sperm DNA. The blots were then hybridized
at 43 °C for 18 h in the same solution containing heat-denatured cDNA probes (corresponding to amino acids 1-377 of
mouse glutamine-dependent NAD synthetase in Fig. 7 or the
entire coding region of human NADsyn2 cDNA) labeled by PCR with
[-32P]dCTP and an antisense primer. The membranes were
once washed in 2 × SSC (1× SSC: 0.15 M NaCl, 0.015 M sodium citrate), 0.1% SDS at 25 °C for 15 min, then
twice with 0.1× SSC, 0.1% SDS at 65 °C for hybridization with the
mouse glutamine-dependent NAD synthetase cDNA or 0.5×
SSC, 0.1% SDS at 50 °C for with NADsyn2 cDNA and exposed to
x-ray film at
80 °C with an intensifying screen. Under these
conditions, NADsyn2 probe did not detect the 3.1-kb message (see Fig.
6B). RNA integrity and loading were assessed using a
glyceraldehyde-3-phosphate dehydrogenase probe.
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RESULTS |
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cDNA Cloning and Structural Analysis of Human NAD Synthetases-- Using the deduced amino acid sequence of B. subtilis NAD synthetase (16) (SWISS-PROT accession number P08164) as a probe, we found two candidate sequences encoding human NAD synthetase, GenBankTM accession numbers AK001493 and HSA236685 in a homology search analysis. Using primers corresponding to the presumed 5'- and 3'-terminal sequences of the candidates, we carried out reverse transcription-PCR as described under "Experimental Procedures" and obtained two human cDNA clones NADsyn1 and NADsyn2, encoding proteins of 706 and 275 amino acids, respectively. To ensure the correct prediction of the open reading frames, we amplified sequences containing 5' upstream regions of the cDNAs using 5'-rapid amplification of cDNA ends. In the 5'-rapid amplification of cDNA ends products, we confirmed sequences corresponding to the sense primers used for amplification of the cDNAs. In 5' upstream region of NADsyn1 cDNA (92 bp) we amplified we did not observe in-frame stop codon but found that the presumed start methionine codon was in a favorable Kozak initiation sequence AGGATGG (22), and the codon was the first in-frame methionine codon. For NADsyn2, we obtained an in-frame stop codon 6 bp upstream of the presumed initiation codon. Thus, we concluded that NADsyn1 and NADsyn2 encode complete coding regions of the respective proteins.
In a search of known functional domains and motifs in protein
structures, we identified the NAD_synthase domain (Pfam
accession number PF02540 http://www.sanger.ac.uk/Software/Pfam/) and an ATP binding site (SGGXDS, P-loop), a conserved sequence fingerprint of
a family of ATP-pyrophosphatases, including NAD synthetase (23), in
both NADsyn2 and the C-terminal half of NADsyn1 (Fig. 2, A and B). A
careful sequence comparison showed that most of the residues forming
ATP binding (Ser-46, Asp-50, and Ser-51) and NaAD binding sites
(Arg-137, Leu-153, Glu-162, Thr-169, Asp-173, Lys-186, Leu-211, His-257
and Lys-258) in B. subtilis synthetase (16) are strictly
conserved in both NADsyn1 and NADsyn2 (Fig. 2B). These
observations suggest that cDNAs for NADsyn1 and NADsyn2 encode NAD
synthetases. We designated the NAD_synthase domain and the P-loop motif
as the synthetase domain.
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In addition to the synthetase domain, NADsyn1 only had the carbon-nitrogen hydrolase (CN_hydrolase) domain (Pfam accession number PF00795) at the N-terminal half (Fig. 2A). The CN_hydrolase domain, with a cysteine residue essential for nitrilase activity (24, 25), is shared with enzymes belonging to the nitrilase family that cleave nitriles as well as amides to produce the corresponding acids and ammonia (Fig. 2C) (26, 27). Because B. subtilis synthetase, which lacks the CN_hydrolase domain (Fig. 2A), utilizes ammonia but not glutamine (16), the presence of the CN_hydrolase domain in NADsyn1 but not in NADsyn2 suggested that the former enzyme uses both glutamine and ammonia in NAD synthesis, whereas the latter is strictly ammonia-dependent. Furthermore, because the critical cysteine residue in the CN_hydrolase domain was also conserved in NADsyn1 at a position of 175 (Fig. 2C), the cysteine (Cys-175) in NADsyn1 was considered to be essential for the use of glutamine. From these sequence analyses, we speculated (i) that both NADsyn1 and NDsyn2 have NAD synthetase activity, (ii) that NADsyn1 uses not only ammonia but also glutamine, whereas NADsyn2 uses only ammonia, and (iii) that the site-directed mutagenesis of Cys-175 in NADsyn1 eliminates glutamine-dependent NAD synthetase activity with the ammonia-dependent activity intact.
Expression and Functional Characterization of Human NAD
Synthetases--
We next expressed NADsyn1, NADsyn2, and a mutant
NADsyn1 in which Cys-175 was replaced with serine (C175S-NADsyn1) in
COS-7 cells as His6-tagged recombinant proteins, and we
purified these proteins on nickel chelate resin. SDS-PAGE analysis
indicated that the purified wild-type (Fig.
3A, inset) and
mutant NADsyn1 (data not shown) have a molecular mass of 80 kDa, in
accordance with the value calculated from the deduced sequences, 80.3 kDa. The purified recombinant NADsyn2 appeared as a single band with a
molecular mass of 34 kDa (Fig. 3B, inset),
slightly larger than the value calculated from the deduced sequence,
30.8 kDa. The difference may depend on the low pI of the recombinant
protein (pI 5.9).
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To investigate whether NADsyn1, NADsyn2, and C175S-NADsyn1 have the
predicted enzymatic activities, we incubated the purified recombinant
proteins with glutamine or NH4Cl in the presence of NaAD
and ATP then determined the NAD synthetase activities of the proteins
using a fluorometric method. As shown in Fig.
4A, the recombinant wild-type
NADsyn1 exhibited almost the same activity with either glutamine or
NH4Cl. On the other hand, the recombinant NADsyn2 catalyzed
NAD synthesis primarily with NH4Cl (Fig. 4C). In
marked contrast with the wild-type NADsyn1, the activity of the mutant
NADsyn1 (C175S-NADsyn1) was not detected when glutamine was used as a
substrate, whereas the activity remained unaltered with
NH4Cl (Fig. 4B). Kinetic analysis indicated that
NADsyn1 shows a lower Km for glutamine (1.44 mM) than for NH4Cl (13.1 mM) (Table
I). Compared with NADsyn1, NADsyn2 showed
a much lower Km for NH4Cl (34 µM) but a much higher Km for glutamine
(103 mM) (Table I). NADsyn1 and NADsyn2 showed essentially
the same Km values for ATP and NaAD, in the range of
those reported for native synthetases (16, 19). Kinetic parameters of
the mutant NADsyn1 obtained with NH4Cl did not differ from
those of the wild-type NADsyn1, which suggests that disappearance of
the glutamine dependence was not because of a drastic change in the
tertiary structure of NADsyn1. Compared with NADsyn2 (34 µM), the mutant synthetase exhibited a much higher
Km value for NH4Cl (23.9 mM). With the glutamine preparation, which contained up to
0.38% ammonia in itself, the concentration of ammonia in the assay
solution in Fig. 4 would be up to 76 µM. Thus, under
these conditions NADsyn2 but not the mutant synthetase could synthesize
NAD without exogenously added ammonia (Fig. 4), as was seen with the
B. subtilis synthetase (16).
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All these results are consistent with our predictions (i) that both NADsyn1 and NADsyn2 have NAD synthetase activity, (ii) that NADsyn1 utilizes not only ammonia but also glutamine as an amide donor, whereas NADsyn2 is primarily an ammonia-dependent NAD synthetase, and (iii) that Cys-175 in NADsyn1 is essential for the ability to use glutamine as an amide donor. In agreement with our sequence analyses noted above, we therefore conclude that the CN_hydrolase domain in the N-terminal half of NADsyn1, in particular Cys-175, is responsible for utilization of glutamine as an amide donor and, thus, confers glutamine dependence on the synthetase, whereas the synthetase domain in NADsyn1 and NADsyn2 participates solely in NAD synthesis from ammonia. To further confirm the role of the CN_hydrolase domain in using glutamine, we made a chimera consisting of the N-terminal region of NADsyn1 and NADsyn2 as well as a construct containing solely the N- or C-terminal half of NADsyn1. However, because of their insufficient expressions, we could not characterize these constructs (data not shown).
We next examined the stoichiometry of the reactions catalyzed by the
recombinant synthetases. The purified recombinant NADsyn1 and NADsyn2
were incubated with glutamine and NH4Cl, respectively, in
the presence of NaAD and ATP and the reaction products were analyzed by
reversed phase HPLC. As shown in Fig. 5,
1 nmol of AMP and PPi was produced per 1 nmol of NAD
synthesized during the reaction catalyzed by each synthetase. These
results indicate that amidation of NaAD by the recombinant enzymes is
associated with ATP cleavage to AMP and PPi, as noted for
the native enzyme (28). Omission of either ATP, Mg2+, NaAD,
or amide donors from the reaction mixture resulted in a complete loss
of NAD synthesis by each enzyme (data not shown).
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To examine whether catalytically active forms of human NADsyn1 and NADsyn2 are multimers as native synthetases (16, 19, 20), we fractionated the purified recombinant synthetases by non-denaturing PAGE and determined NAD synthetase activity in gel slices. As shown in Fig. 3, activities of NADsyn1 and NADsyn2 had mobilities consistent with proteins of 500 and 70 kDa, respectively, suggesting that NADsyn1 and NADsyn2 may exist as a homohexamer and a homodimer, respectively.
Tissue Distribution of NAD Synthetases--
To evaluate the tissue
distribution of NADsyn1 and NADsyn2, Northern blot analyses were done
with total RNA from various mouse tissues and several human cell lines.
As shown in Fig. 6A, a message of 3.1 kb was detected for NADsyn1, with various intensities. The major
sites of NADsyn1 gene expression were the small intestine, kidney,
liver, and testis, whereas the skeletal muscle, spleen, lung, heart,
and brain showed a weak signal. In the liver and small intestine, an
additional signal was observed at 2.1 kb. The NADsyn1 gene was also
expressed in human glioma (LN229) and promyelocytic leukemia (HL60)
cell lines (data not shown). Although an EST data base homology search
did not reveal a human clone with a high degree of similarity to
NADsyn2, Northern blot analysis clearly showed that an
mRNA species of 1.4 kb is expressed in human cells LN229, HL60, and
HepG2 and Huh7 (hepatocyte cell lines) (Fig. 6B). In the
mouse, all of the tissues tested expressed the 1.4-kb mRNA,
probably representing a mouse homologue of NADsyn2. In the lung and in
skeletal muscle, the same 1.4-kb signal was observed after long
exposure of the blot (data not shown). In mouse brain and kidney and
human cell lines an additional 2.6-kb mRNA species was
observed.
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Sequence Alignment of NADsyn1 Homologues--
The deduced amino
acid sequences of NADsyn1 and NADsyn2 showed mismatches of three and
four amino acids, respectively, compared with those deposited in
GenBankTM data base (AK001493 and HSA236685, respectively)
(Figs. 7 and 2B). A homology
search in a protein data base revealed that NADsyn1 exhibits
significant amino acid identity to several eucaryotic putative proteins
from the mouse (83%), Saccharomyces cerevisiae (58%),
Drosophila melanogaster (53%) and C. elegans
(46%) (Fig. 7). Alignment of the NADsyn1 sequence with these proteins
showed that there are highly conserved regions among the five proteins, including ATP and NaAD binding sites in the C-terminal regions and the
essential cysteine residues for use of glutamine in the N-terminal
halves. The CN_hydrolase domain of NADsyn1 also showed sequence
similarity to NAD synthetases from M. tuberculosis (18) and
Rhodobacter capsulatus (29) (Fig. 2C), and in
particular, the critical cysteine residues were strictly conserved in
the two bacterial synthetases. NADsyn2 did not exhibit significant sequence similarity to any other eucaryotic proteins over the entire
length.
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DISCUSSION |
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The present study is the first identification of open reading frames encoding two NAD synthetases, NADsyn1 and NADsyn2, in humans. Heterologous expression of the synthetases indicated that although NADsyn1 utilizes both glutamine and ammonia as amide donors, ammonia may not serve as a physiological amide donor for NADsyn1 in vivo (Km for NH4Cl >10 mM). In marked contrast, NADsyn2 uses ammonia more efficiently than does NADsyn1 (Km for NH4Cl = 34 µM) but appears to be unable to use glutamine as a physiological amide donor (Km for glutamine >100 mM). Thus, we conclude that NADsyn1 is a glutamine-dependent NAD synthetase, whereas NADsyn2 is a strictly ammonia-dependent synthetase. To our knowledge, this is the first evidence for the presence of an ammonia-dependent NAD synthetase in eucaryotes. Furthermore, by comparing the catalytic activity of NADsyn2, which lacks CN_hydrolase domain, with the mutant NADsyn1, in which Cys-175 corresponding to the catalytic cysteine residue in nitrilases (24, 25) was replaced with Ser, we identified the CN_hydrolase domain as the functional domain of NAD synthetase to abstract nitrogen from the amide of glutamine and, thus, to use glutamine as an amide donor.
Identification of the CN_hydrolase domain as the determinant of glutamine dependence means that the homologues of human NADsyn1 found in different species are also glutamine-dependent NAD synthetases. The domain occurring in the NAD synthetase from M. tuberculosis, known to catalyze NAD synthesis with glutamine (18), now provides the previously unrecognized structural basis underlying the glutamine dependence of the synthetase. These results suggest that the glutamine-dependent NAD synthetases can be classified as a possible glutamine amidase into the nitrilase family. In the nitrilases, an invariant cysteine residue has been proposed to act as a nucleophile in the catalytic mechanism, where a nitrile carbon is subjected to a nucleophilic attack by sulfhydryl group in the active site of the enzyme (24, 25). Because the critical cysteine residue in the nitrilases is conserved in these glutamine-dependent NAD synthetases, the cysteine residues in the synthetases probably carry out a nucleophilic attack on a carbonyl carbon of glutamine, abstracting ammonia from glutamine. In the synthetase domains of these synthetases, after adenylation of NaAD in the presence of ATP, the ammonia thus abstracted in the CN_hydrolase domain attacks the adenylated NaAD, resulting in NAD, as proposed (28). X-ray diffraction analysis and a detailed structure-functional analysis of NADsyn1 will give a better understanding of the catalytic mechanism of these glutamine-dependent NAD synthetases.
We showed that NADsyn1 exerts catalytic activity in a multimeric form. NAD synthetases purified from human erythrocytes and yeast, glutamine-dependent and, thus, expected to possess CN_hydrolase domain, are also multimeric enzymes (19, 20). It has been reported that nitrilase family members form a multimer, probably by subunit contact through highly hydrophobic regions conserved in the CN_hydrolase domain (26) (Fig. 2C). Thus, it appears that the CN_hydrolase domain, including hydrophobic regions, participates in multimer formation of NADsyn1.
Glutamine-dependent yeast NAD synthetase has been reported to have two components, an 80-kDa ammonia-dependent NAD synthetase subunit and an additional 65-kDa subunit, and the latter has been hypothesized to use glutamine as amide donor (19, 30). However, because the yeast homologue of human NADsyn1 with a calculated molecular mass of 80.7 kDa has the CN_hydrolase domain, it seems that the 80-kDa subunit solely represents the yeast synthetase, and it is unlikely that the 65-kDa subunit is required for glutamine-dependent NAD synthetase activity.
The wide variability of NADsyn1 expression revealed by Northern blot analysis may reflect differences in NAD demand among animal tissues. Abundant expression of NADsyn1 was observed in the small intestine, liver, kidney, and testis, whereas skeletal muscle and the heart showed very weak signals. However, nicotinamide mononucleotide adenylyltransferase, catalyzing the formation of the substrate of NAD synthetase NaAD (Fig. 1), has been reported to be expressed mainly in skeletal muscle and the heart (14), thus being inconsistent with NADsyn1 expression. This raises the question of how NAD synthesis occurs in these tissues. NADsyn2 is expressed in skeletal muscle and in the heart. In these tissues, gene expression of glutaminase catalyzing the formation of ammonia from glutamine has also been demonstrated (31). Taken together the finding that NADsyn2 could catalyze NAD synthesis using ammonia as an amide donor, NADsyn2 may largely mediate NAD synthesis in these tissues. Alternatively, based on a somewhat higher affinity of nicotinamide mononucleotide adenylyltransferase for NMN than for NaMN (14), NAD may also be synthesized via direct conversion of NMN to NAD in the tissues. For a better understanding of the regulation of NAD biosynthesis in higher organisms, including humans, further analyses on quinolinic acid phosphoribosyltransferase and nicotinic acid phosphoribosyltransferase expression in animal tissues are under investigation.
In the present study, we identified glutamine- and
ammonia-dependent human NAD synthetases, NADsyn1 and
NADsyn2, respectively, with distinct tissue distribution of the
synthetases, and we obtained evidence that the CN_hydrolase domain
confers glutamine dependence on the former enzyme. Our results suggest
that the glutamine-dependent NAD synthetase is classified
as a glutamine amidase into the nitrilase family and that the newly
identified metabolic pathway involving the
ammonia-dependent NAD synthetase plays a role in NAD
biosynthesis. These are important clues to better understand detailed
structures fundamental to catalytic activity of the enzyme and to
elucidate regulatory mechanisms of cellular NAD metabolism.
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ACKNOWLEDGEMENT |
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We thank K. Tsuchie for technical assistance.
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FOOTNOTES |
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* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AB091316 (NADsyn1) and AB091317 (NADsyn2).
To whom correspondence should be addressed. Tel.: 81-853-20-2121;
Fax: 81-853-20-2120; E-mail: mikat@shimane-med.ac.jp.
Published, JBC Papers in Press, January 23, 2003, DOI 10.1074/jbc.M209203200
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ABBREVIATIONS |
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The abbreviations used are: NaMN, nicotinic acid mononucleotide; CN_hydrolase, carbon-nitrogen hydrolase; HPLC, high performance liquid chromatography; NaAD, nicotinic acid adenine dinucleotide; kb, kilobase(s).
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Lee, H. C.
(1997)
Physiol. Rev.
77,
1133-1164 |
2. |
Hara, N.,
Tsuchiya, M.,
and Shimoyama, M.
(1996)
J. Biol. Chem.
271,
29552-29555 |
3. |
Okazaki, I. J.,
and Moss, J.
(1998)
J. Biol. Chem.
273,
23617-23620 |
4. | Haag, F., and Koch-Nolte, F. (1998) J. Biol. Regul. Homeost. Agents 12, 53-62[Medline] [Order article via Infotrieve] |
5. |
Lindahl, T.,
and Wood, R. D.
(1999)
Science
286,
1897-1905 |
6. | Pieper, A. A., Verma, A., Zhang, J., and Snyder, S. H. (1999) Trends Pharmacol. Sci. 20, 171-181[CrossRef][Medline] [Order article via Infotrieve] |
7. |
Anderson, R. M.,
Bitterman, K. J.,
Wood, J. G.,
Medvedik, O.,
Cohen, H.,
Lin, S. S.,
Manchester, J. K.,
Gordon, J. I.,
and Sinclair, D. A.
(2002)
J. Biol. Chem.
277,
18881-18890 |
8. | Tissenbaum, H. A., and Guarente, L. (2001) Nature 410, 227-230[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Smith, J. S.,
Brachmann, C. B.,
Celic, I.,
Kenna, M. A.,
Muhammad, S.,
Starai, V. J.,
Avalos, J. L.,
Escalante-Semerena, J. C.,
Grubmeyer, C.,
Wolberger, C.,
and Boeke, J. D.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
6658-6663 |
10. | Luo, J., Nikolaev, A. Y., Imai, S., Chen, D., Su, F., Shiloh, A., Guarente, L., and Gu, W. (2001) Cell 107, 137-148[Medline] [Order article via Infotrieve] |
11. | Foster, J. W., and Moat, A. G. (1980) Microbiol. Rev. 44, 83-105 |
12. | White, H. B. (1982) in Pyridine Nucleotide Coenzyme: Biosynthesis of Salvage Pathways of Pyridine Nucleotide Coenzymes (Everse, J. , Anderson, B. M. , and You, K. S., eds) , pp. 1-17, Academic Press, Inc., New York |
13. | Tritz, G. J. (1987) in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (Neidhardt, F. C. , Ingraham, J. L. , Low, K. B. , Magasanik, B. , Schaechter, M. , and Umbarger, H. E., eds) , pp. 557-563, American Society for Microbiology, Washington, D. C. |
14. |
Emanuelli, M.,
Carnevali, F.,
Saccucci, F.,
Pierella, F.,
Amici, A.,
Raffaelli, N.,
and Magni, G.
(2001)
J. Biol. Chem.
276,
406-412 |
15. | Fukuoka, S., Nyaruhucha, C. M., and Shibata, K. (1998) Biochim. Biophys. Acta 1395, 192-201[Medline] [Order article via Infotrieve] |
16. |
Nessi, C.,
Albertini, A. M.,
Speranza, M. L.,
and Galizzi, A.
(1995)
J. Biol. Chem.
270,
6181-6185 |
17. | Rizzi, M., Nessi, C., Mattevi, A., Coda, A., Bolognesi, M., and Galizzi, A. (1996) EMBO J. 15, 5125-5134[Abstract] |
18. | Cantoni, R., Branzoni, M., Labò, M., Rizzi, M., and Riccardi, G. (1998) J. Bacteriol. 180, 3218-3221[Abstract] |
19. |
Yu, C. K.,
and Dietrich, L. S.
(1972)
J. Biol. Chem.
247,
4794-4802 |
20. | Zerez, C. R., Wong, M. D., and Tanaka, K. R. (1990) Blood 75, 1576-1582[Abstract] |
21. | Zalkin, H. (1985) Methods Enzymol. 113, 297-302[Medline] [Order article via Infotrieve] |
22. | Kozak, M. (1991) J. Cell Biol. 115, 887-903[Abstract] |
23. | Tesmer, J. J. G., Klem, T. J., Deras, M. L., Davisson, V. J., and Smith, J. L. (1996) Nat. Struct. Biol. 3, 74-86[Medline] [Order article via Infotrieve] |
24. | Kobayashi, M., Yanaka, N., Nagasawa, T., and Yamada, H. (1992) Biochemistry 31, 9000-9007[Medline] [Order article via Infotrieve] |
25. | Kobayashi, M., Izui, H., Nagasawa, T., and Yamada, H. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 247-251[Abstract] |
26. |
Bork, P.,
and Koonin, E. V.
(1994)
Protein Sci.
3,
1344-1346 |
27. | Novo, C., Tata, R., Clemente, A., and Brown, P. R. (1995) FEBS Lett. 367, 275-279[CrossRef][Medline] [Order article via Infotrieve] |
28. |
Spencer, R. L.,
and Preiss, J.
(1967)
J. Biol. Chem.
242,
385-392 |
29. | Willison, J. C., and Tissot, G. (1994) J. Bacteriol. 176, 3400-3402[Abstract] |
30. | Zalkin, H., and Smith, J. L. (1998) Adv. Enzymol. Relat. Areas Mol. Biol. 72, 87-144[Medline] [Order article via Infotrieve] |
31. | Elgadi, K. M., Meguid, R. A., Qian, M., Souba, W. W., and Abcouwer, S. F. (1999) Physiol. Genomics 1, 51-62[Medline] [Order article via Infotrieve] |