(Received for publication, August 1, 1994; and in revised form, November 16, 1994)
From the
We have cloned and sequenced the hxA gene coding for the xanthine dehydrogenase (purine hydroxylase I) of Aspergillus nidulans. The gene codes for a polypeptide of 1363 amino acids. The sequencing of a nonsense mutation, hxA5, proves formally that the clones isolated correspond to the hxA gene. The gene sequence is interrupted by three introns. Similarity searches reveal two iron-sulfur centers and a NAD/FAD-binding domain and have enabled a consensus sequence to be determined for the molybdenum cofactor-binding domain. The A. nidulans sequence is a useful outclass for the other known sequences, which are all from metazoans. In particular, it gives added significance to the missense mutations sequenced in Drosophila melanogaster and leads to the conclusion that while one of the recently sequenced human genes codes for a xanthine dehydrogenase, the other one must code for a different molybdenum-containing hydroxylase, possibly an aldehyde oxidase. The transcription of the hxA gene is induced by the uric acid analogue 2-thiouric acid and repressed by ammonium. Induction necessitates the product of the uaY regulatory gene.
Xanthine dehydrogenases are ubiquitous enzymes that have been
thoroughly studied from the biochemical point of view. Their structure
is conserved throughout evolution. They are dimers, with each monomer
of 150 kDa containing a complex electron transport chain with a
pterin-bound molybdenum, a flavin, and two iron-sulfur centers as
cofactors. NAD is usually the terminal electron acceptor (Coughlan,
1980; Rajagopalan, 1991; Wootton et al., 1991). A number of
enzymes related to the xanthine dehydrogenases have been described.
These show the same overall general structure, but different substrate
specificities. Among these are the aldehyde oxidases from a wide range
of metazoans, including mammals and Drosophila melanogaster,
and the pyridoxal dehydrogenase from D. melanogaster (Krenitsky et al., 1972, 1974; Branzoli and Massey, 1974;
Courtright, 1976).
The M of the Aspergillus
nidulans xanthine dehydrogenase has been estimated to be 304,000.
Its cofactor content is identical to that of other enzymes of this
group (Scazzocchio et al., 1973; Lewis et al., 1978;
Scazzocchio and Sealy-Lewis, 1978; Mehra and Coughlan, 1989). Many
loss-of-function mutations of the A. nidulans enzyme have been
obtained (Darlington et al., 1965; Alderson and Scazzocchio,
1967; Darlington and Scazzocchio, 1967). Two classes of substrate
specificity mutations have also been obtained. The first shows an
altered kinetics of inhibition by allopurinol (Scazzocchio, 1966). The
second shows a number of pleiotropic effects, the most striking of
which is that while the wild-type enzyme hydroxylates 2-hydroxypurine
at position 8, the mutant enzyme hydroxylates this same analogue at
position 6 (Scazzocchio and Sealy-Lewis, 1978). All these mutations map
in one gene, hxA, located in chromosome V. A fine structure
map has been constructed, which positions all substrate specificity
mutations in a discrete domain of the gene. (
)Transformation
techniques (see below) permit the physical location of any mutation and
introduction of new ones at selected places in the gene. The enzyme of D. melanogaster has also been subject to a thorough genetic
analysis (Chovnick et al., 1977, 1990; Gray et al.,
1991; Hughes et al., 1992a, 1992b; Doyle and Bray, 1994). The
selection techniques that are available for A. nidulans and D. melanogaster are not identical; thus, the classes of
mutations extant in the two organisms are only partially overlapping.
In particular, substrate specificity mutations have been obtained only
in A. nidulans.
A second enzyme, purine hydroxylase II, has been described in A. nidulans. This is physiologically a nicotinate hydroxylase, and it is inducible by 6-hydroxynicotinate (Sealy-Lewis et al., 1979) and controlled with other enzymes of the nicotinate degradation pathway (Scazzocchio et al., 1973; Scazzocchio, 1980, 1994). The enzyme accepts hypoxanthine, but not xanthine, as a substrate and presents some interesting kinetic and mechanistic peculiarities (Lewis et al., 1978; Coughlan et al., 1984; Scazzocchio, 1994 (for review)). Other nicotinate hydroxylases have been described in bacteria, but none has a hypoxanthine hydroxylase activity (Hirschberg and Ensign, 1972; Imhoff and Andreesen, 1979). Thus, a comparison of the two purine hydroxylases of A. nidulans would be of evolutionary and mechanistic interest.
The genetic system of A. nidulans has provided a
complete picture of the gene-protein relationships of the
molybdenum-containing enzymes. The structural genes coding for the
three molybdenum-containing enzymes, xanthine dehydrogenase, purine
hydroxylase II, and nitrate reductase, have been identified and mapped,
and many alleles in each have been characterized and ordered in fine
structure maps (Scazzocchio et al., 1973; Scazzocchio and
Sealy-Lewis, 1978; Scazzocchio, 1980; Tomsett and Cove, 1979). The genes involved in the molybdenum cofactor biosynthesis have
been identified. The concept of a common molybdenum-containing cofactor
was first derived from the existence of six genes in which mutations
led to the loss of nitrate reductase and xanthine dehydrogenase
activities (Pateman et al., 1964). Later it was found that
purine hydroxylase II also requires the molybdenum cofactor
(Scazzocchio, 1973; Scazzocchio, 1980). Another gene, hxB,
codes for a protein needed for the substrate-specific hydroxylation
activity of both purine hydroxylases I and II, but not for some
ancillary activities such as the NADH dehydrogenase activity
(Scazzocchio et al., 1973; Sealy-Lewis et al., 1978;
Scazzocchio, 1980, 1994). Mutations in this gene do not affect nitrate
reductase activity. This gene would seen to code for a protein
necessary for a post-translational modification and may well be
isofunctional with the ma-l gene of D. melanogaster,
coding for a protein involved in the generation of the molybdenum-bound
sulfur atom (Wahl et al., 1982).
The three molybdoproteins
of A. nidulans are inducible, each by a specific metabolite.
Each enzyme is coinduced with other enzymes of the same pathway. The
specific regulatory genes involved in each induction process have been
genetically characterized (Cove, 1979; Scazzocchio, 1994 (for reviews))
and, in the cases of the regulatory genes involved in nitrate and
purine assimilation, cloned and sequenced (Burger et al.,
1991a, 1991b; Suárez et al., 1991). ()The protein coded by the uaY gene mediates uric
acid induction of at least eight activities of the purine utilization
pathway, including xanthine dehydrogenase (Scazzocchio et al.,
1982; Scazzocchio, 1994). The transcription of the genes of the purine
utilization pathway necessitates a specific inducer, uric acid, but
occurs efficiently only in the absence of ammonium or glutamine.
However, neither the response to specific induction nor ammonia
repression is identical in all genes of the pathway, with the basal
noninduced level of xanthine dehydrogenase being considerably higher
than that of the uric-acid permease (Gorfinkiel et al., 1993).
A GATA factor, the product of the areA gene, mediates ammonia
and glutamine repression of possibly all genes coding for enzymes and
permeases involved in the utilization of nitrogen sources (Arst and
Cove, 1973; Kudla et al., 1990).
The genomic and cDNA sequences of the genes coding for urate oxidase and the urate-xanthine permease have been reported. The transcription of these genes under different conditions and in wild-type and mutant backgrounds has been studied (Oestreicher and Scazzocchio, 1993; Gorfinkiel et al., 1993).
The cloning and sequencing of the hxA gene are of dual interest. From the point of view of regulation, it will contribute to our understanding of the different responses to uric acid induction and ammonium repression of different genes responding to the same regulatory gene products. From the point of view of enzymology, the comparison of enzymes from organisms as different as mammals, insects, and fungi will allow the identification of putative, discrete functional domains. In addition, the availability of a unique set of mutations will permit the definitive identification of functional domains and residues determining substrate binding and specificity.
Figure 1:
Restriction
map of plasmid pBAN884 and sequencing strategy. The continuoussegment indicates the BglII fragment cloned in
plasmid pBAN884. The shortarrows indicate the extent
and direction of each sequencing reaction. The whole BglII-SacII region was sequenced on both strands; the
direction of transcription is from left to right. This was determined
by hybridization with the single-stranded plasmids pBAN873X and
pBAN876X, which contain the same fragment cloned in opposite
orientations. Plasmid pBAN816C was used as a probe for screening the
ZAPII cDNA library. B, BglII; C, ClaI; S, SacII; Xb, XbaI.
Other plasmids used either for sequencing or transformation experiments
are also shown. kb, kilobase.
Figure 2:
Transcriptional regulation of the hxA gene. Northern blotting was performed as described under
``Experimental Procedures.'' The hxA probe was the
6.1-kbp BglII-BglII fragment (see Fig. 1)
labeled with [-
P]dCTP using random
hexanucleotide primers (Amersham Corp.). The same blots were also
hybridized with the actin gene (Fidel et al., 1988) similarly
labeled as a control of RNA loading. In all panels, urea indicates growth for 20 h on 5 mM urea as nitrogen
source, and amm. indicates growth for 20 h on limiting
ammonium (2.5 mM) as nitrogen source. Lanes 1, no
additions; lanes 2, addition of both inducer and corepressor
(27 µM 2-thiouric acid and 10 mM ammonium ion,
respectively); lanes 3, addition of inducer. In a, we
compare a uaY
strain and a uaY205 strain. In b, two comparisons are made: first, a uaY6 strain with a wild-type (indicated as uaY
) strain grown on urea as nitrogen source
in lanes 1-3; second, we show the repressing effect of
growth on limiting ammonium (2.5 mM) on the wild-type strain.
In c, we compare a wild-type (indicated as areA
) and an areA600 strain grown on
limiting ammonium (2.5 mM) as sole nitrogen source. The wild
type grown on urea under inducing conditions is also included as a
standard (lane 3). Each panel corresponds to a different and
separate experiment. In c (as in the other panels), all lanes
come from one and the same Northern blot, but other lanes irrelevant to
this experiment were cut out. To permit all relevant comparisons, the
experiments shown in the three panels overlap partially. b and c are overexposed (in relation to a) to reveal more
clearly the low amounts of xanthine dehydrogenase mRNA present under
noninduced or repressed conditions. Thus, lanes 3 for the wild
type (indicated as uaY
and areA
, respectively) are saturated, and this
results in an apparently lower induced/noninduced ratio than in a.
Figure 3: hxA nucleotide sequence and deduced amino acid sequence. Noncoding regions and introns are presented in lower-case letters. The upper numbers to the right of the sequence indicate nucleotides, and the lower numbers indicate amino acids. The arrows are above the 5`-WGATAR sequences that are putative AreA-binding sites. The dashed line shows the TATA-like sequence. The pyrimidine-rich sequence is underlined. The verticals half-arrows indicate transcription initiation and termination points. Boldface type indicates the 5`- and 3`-splicing and lariat intron sites. The box corresponds to the AATAA sequence that is probably involved in RNA termination. The double-headed arrow indicates a sequence that has been implicated in mRNA termination (see ``Results and Discussion''). The mutational change in hxA5 is indicated above the nucleotide sequence. Also indicated are the HindIII and EcoRV sites that limit the fragment determined by transformation experiments to contain the hxA5 mutation and the ClaI sites that correspond to the limits of plasmid pBAN882C, which has been shown to contain the wild-type sequences corresponding to four substrate specificity mutations (see ``Results and Discussion'').
The size of the transcribed mRNA was estimated at 4.5 kilobases on Northern blots and is entirely contained in the 5.5-kbp fragment sequenced (data not shown; see below). A putative TATA box, 5`-TATTAA, is 37 nucleotides upstream from the start of transcription. Three 5`-WGATAR sequences that are potential AreA-binding sites (Fu and Marzluf, 1990; Merika and Orkin, 1993) are found upstream from the start of transcription. A pyrimidine-rich region precedes the start of transcription, as seen frequently in Ascomycetes. The different cDNA clones analyzed have poly(A) tails from 33 to 36 nucleotides long. The sequence 5`-AATAA upstream from the termination point of the mRNA could be the polyadenylation signal (Fitzgerald and Shenk, 1981; Hawkins, 1987). However, a 5`-CATGGTGAT sequence is also present; this sequence has also been implicated in mRNA termination (Upshall et al., 1986). The 5`-, 3`-, and lariat intron sequences show some agreement with the sequences previously proposed as consensus sequences (Ballance, 1986; Gurr et al., 1987).
The sequence surrounding the putative ATG codon shows an excellent agreement with other sequences described for lower eukaryotes, in particular for the uaZ and uapA genes. The codon usage does not differ significantly from the codon usage reported for most genes of A. nidulans (Lloyd and Sharp, 1991).
The translated peptidic
sequence has 1363 amino acids. The RNA transcript of the hxA gene contains 4.425 nucleotides. The size of the mature mRNA,
4.284 nucleotides after subtraction of the intron sequences, compares
well with the 4.2 kilobases estimated by preparative
ultracentrifugation followed by identification by in vitro translation (Hanselman, 1984). The calculated M of the polypeptide is 149,410. This compares well with the M
of the dimer estimated at 304,000 on
nondenaturing gels (Lewis et al., 1978), but is larger than
the value of 135,000 estimated for the monomer run on denaturing gels
after immunoprecipitation (Hanselman, 1984). The pI of 5.86 estimated
using the program MacVector
(Kodak Scientific Imaging
Systems, New Haven, CT) is similar to those of other xanthine
dehydrogenases (Keith et al., 1987; Riley, 1989; Houde et
al., 1989).
Figure 4: Comparison of eight xanthine dehydrogenase peptidic sequences. The comparison has been carried out with the Pileup program of the University of Wisconsin Genetics Computer Group software and printed with the Prettybox program. Black, identical amino acids; dark and light gray, similar amino acids; white, nonconserved residues. Note that the program actually minimizes identities and similarities, as it compares sequences to a consensus sequence established when at least five out of the eight proteins have the same residue at one given position. For example, at position 26 of the A. nidulanssequence, we found an arginine in four sequences and glycine in four others. This is recorded as white (nonconserved) by the Prettybox program. In some places, slightly different alignments could be done by eye, which would increase similarities, but will create more gaps. An example of this can be found around positions 420-423 of the sequence of A. nidulans, where by creating a new gap, a universally conserved (except for H2) methionine will fall into place. The sequences are the following: A. nidulans (An), D. melanogaster (Dm) (Lee et al., 1987; Keith et al., 1987), D. pseudoobscura (Dp) (Riley, 1989), C. vicina (Cv) (Houde et al., 1989), rat liver (R) (Amaya et al., 1990), mouse liver (M) (Terao et al., 1992), human liver H1 (Ichida et al., 1993), and human liver H2 (Wright et al., 1993).
The amino acid identity of the A. nidulans xanthine dehydrogenase with the mammalian and insect enzymes is around 40-46% according to the organism or the alignment program used (see Fig. 4). The enzyme of A. nidulans is a few amino acids longer than all others. The additional amino acids are mainly at the amino terminus. It can be noticed that the amino terminus of the protein is highly variable, the similarity starting with a universally conserved phenylalanine at position 39 of A. nidulans and at position 8 of the D. melanogaster sequence.
The xanthine dehydrogenase of D. melanogaster has been shown to be peroxisomal (Beard and Holtzman, 1987). The putative peroxisomal localization signals of the enzyme (Gould et al., 1989), the AKL motif at positions 253-255 and AKI at positions 616-618, are conserved in the A. nidulans enzyme at positions 292-294 and 643-645, respectively.
The xanthine dehydrogenases have two
[2Fe-2S] centers, described as an electron sink (Edmondson et al., 1973; Coughlan and Rajagopalan, 1980). The first
center, revealed by analyzing the amino acid sequence with the PROSITE
program (Bairoch, 1991), belongs to the same type found in the
ferredoxin from a number of photosynthetic organisms, bacterial
fumarate reductase, and eukaryotic succinic dehydrogenase. The general
sequence of this motif is
CXCX
CX
C,
where X is any amino acid and n equals 11 in succinic
dehydrogenase, 29 in the ferredoxins, and 21 in all reported xanthine
dehydrogenases. The second putative iron-sulfur center has been located
between amino acids 78 and 176 in the sequence of the D.
melanogaster xanthine dehydrogenase (Wootton et al.,
1991). This corresponds to residues 108-206 in the A.
nidulans sequence. An alignment between this region and a sequence
of the iron-sulfur center of the ferredoxin of Clostridium
pasteurianum has been proposed (Hughes et al., 1992b).
All xanthine dehydrogenases have the sequence (H/N)G(S/T)QCGFCTP, while
the sequence in the bacterial ferredoxin is NGKQQFCYS, showing a
significant conservation around the second cysteine of the
carboxyl-terminal iron-sulfur center.
The putative FAD-binding site cannot be precisely identified by sequence comparison. There are no obvious similarities to the consensus sequences described by Correll et al.(1993). The enzymatic data underlying the tentative identification of the FAD-binding site in the D. melanogaster enzyme (Hughes et al., 1992a) have now been withdrawn (Doyle and Bray, 1994). There is a cluster of missense mutations mapping in this region (residues 348, 353, and 357 in D. melanogaster) (Hughes et al., 1992a). This sequence is conserved in A. nidulans (residues 387-395). The A. nidulans enzyme shows three conservative changes in this region, one of which is shared with all the mammalian enzymes, with the exception of one of the human enzymes (H2 in Fig. 4). The fact that this conserved region maps just upstream from the NAD-binding site makes it likely that it is in fact the FAD-binding site. However, a direct demonstration has not yet been obtained.
The
NAD-binding site has been chemically identified in the chicken xanthine
dehydrogenase. The analogue
5`-p-fluorosulfonylbenzoyladenosine inactivates the enzyme by
covalently binding to a tyrosine (Nishino and Nishino, 1987, 1989).
This corresponds to tyrosine 429 in the sequence of A.
nidulans. Excluding the aberrant human enzyme (H2 in Fig. 4), the following consensus sequence can be established
starting at position 425 of the enzyme of A. nidulans:
FFXGY*R(T/N)X(I/L)XPXH, where Y*
denotes the tyrosine that is labeled by
5`-p-fluorosulfonylbenzoyladenosine in the chicken enzyme. All
residues marked X differ between the mammalian and insect
sequences. In every case, both the A. nidulans enzyme and the
human H2 enzyme have a unique residue at these positions, which differs
from both the mammalian and dipteran enzymes. This is of some relevance
because Amaya et al.(1990) have argued that the environment of
the Y* residue determines whether the conversion of a NAD-dependent
form to an O-dependent form is possible. No oxidase
activity has been observed for the A. nidulans enzyme, either
in its native form or following partial proteolysis (Mehra and
Coughlan, 1989).
The domain involved in the fixation of the
molybdenum cofactor has been tentatively identified by comparing the
nitrate reductases from Arabidopsis thaliana and A.
nidulans, the sulfite oxidases from rat and chicken liver, and the
xanthine dehydrogenases from rat and D. melanogaster (Hughes et al., 1992b). The sequencing of the A. nidulans hxA gene permits the comparison of a nitrate reductase (Kinghorn and
Campbell, 1989; Johnstone et al., 1990) and a xanthine
dehydrogenase from the same organism. We have compared 11 available
eukaryotic nitrate reductases, two sulfite oxidases, and the eight
available xanthine dehydrogenases. We have derived a consensus sequence
for each group of enzymes. This is shown in Fig. 5, with a
suggested ``consensus'' sequence for the molybdenum
cofactor-binding domain of eukaryotic molybdopterin-containing enzymes.
The BLAST program revealed a clear similarity between the A.
nidulans xanthine dehydrogenase and two prokaryotic enzymes, the
aldehyde dehydrogenase from Acetobacter polyoxogenes (Tamaki et al., 1989) and the nicotine dehydrogenase from Arthrobacter nicotinovorans (GenBank accession
number X75338). The putative molybdenum cofactor-binding domains of
these two enzymes are also shown in Fig. 5. The nicotine
dehydrogenase has been shown to contain molybdopterin (Freudenberg et al., 1988). There is no information about the cofactor
complement of the aldehyde dehydrogenase (Tamaki et al.,
1989), but the similarity data suggest that it may also contain a
molybdenum cofactor. Brandsch (
)has also observed
similarities in the putative molybdenum cofactor-binding domain of the
nicotine dehydrogenase and the eukaryotic xanthine dehydrogenases.
Thoenes et al.(1994) have recently reported similarities
between the aldehyde oxidoreductase from Desulfovibrio gigas and the metazoan xanthine dehydrogenases. They propose a large
``molybdopterin-binding domain'' that extends far beyond
toward the carboxyl-terminal of what is proposed by us. A sequence
similar to that shown in Fig. 5can also be found toward the
amino terminus of their proposed molybdopterin-binding domain.
Figure 5:
Putative molybdenum cofactor-binding
domain. Ten sequences of nitrate reductase were aligned using the
Pileup program of the University of Wisconsin Genetics Computer Group
software including A. nidulans (Johnstone et al.,
1990), Fusarium oxisporum (Diolez et al., 1993), Neurospora crassa (Okamoto et al., 1991), Ustilago maydis (Banks et al., 1993), Volvox
carteri (Gruber et al., 1992), A. thaliana (Cheng et al., 1988), barley (Schnorr et al.,
1991), Nicotiana tabacum (Vaucheret et al., 1989),
rice (Cheng et al., 1989), and spinach (Prosser and Lazarus,
1990); the program Prettybox was utilized, and the consensus sequence
was noted. The same procedure was applied for two sequences of sulfite
oxidase: rat (Garrett and Rajagopalan, 1994) and chicken (Neame and
Barber, 1989) liver. The alignment of the xanthine dehydrogenases is as
described in the legend to Fig. 4. The BLAST program aligned the
sequence of the xanthine dehydrogenase of A. nidulans with the
aldehyde dehydrogenase of A. polyoxogenes (Tamaki et
al., 1989) and the nicotine dehydrogenase of A.
nicotinovorans. The sequence of the aldehyde oxidoreductase of D. gigas (Thoenes et al., 1994) was aligned by hand.
The final alignment was made by hand. The double boxes represent the universally conserved amino acids; the single
boxes represent the amino acids where we found conservative
substitutions; and the numbers represent the residues between
conserved amino acids. S.O. indicates the consensus sequence
for sulfite oxidase (it corresponds to positions 137-214 of the
rat enzyme). N.R. indicates the consensus sequence for the
nitrate reductases (positions 72-157 of A. nidulans). XDH indicates the consensus sequence for the xanthine
dehydrogenases (positions 773-862 of A. nidulans). Nic.D. indicates the sequence of the prokaryotic nicotine
dehydrogenase (positions 203-288 of ndhC). Ald.D. indicates the prokaryotic aldehyde dehydrogenase (positions
368-452). Ald.O. indicates the prokaryotic aldehyde
oxidoreductase (positions 369-446). Note that the later three
enzymes do not have a cysteine that is conserved in all eukaryotic
enzymes. cons. indicates the ``consensus of
consensus'' for all eukaryotic sequences. corresponds to
hydrophobic amino acids, and
to asparagine or glutamine (we find
an exception, serine in the U. maydis sequence);
is
equivalent to histidine or asparagine or glutamine, and
indicates
an aspartic or glutamic acid. For 9 out of the 10 nitrate reductases,
the sixth spacing between conserved residues (between
and G) is
of 16 or 19 amino acids; the exception is the nitrate reductase from N. crassa, which shows a spacing of 30 amino acids. Residues
marked with asterisks are those where the H2 enzyme differs
from other, conventional xanthine
dehydrogenases.
Cofactor (cnx) and hxB mutations result in loss of xanthine dehydrogenase activity while maintaining NADH dehydrogenase activity (Scazzocchio, 1973; Lewis and Scazzocchio, 1977). But cofactor mutations also affect the stability of the A. nidulans xanthine dehydrogenase dimer, and in strains carrying such mutations, it is possible to detect on acrylamide gels the xanthine dehydrogenase monomer by its NADH dehydrogenase activity, while in hxB mutants, the stability of the dimer is not affected (Lewis and Scazzocchio, 1977). This makes a straightforward prediction on the phenotype that should be obtained by mutating the conserved residues in the putative molybdenum cofactor-binding domain.
A number of universally conserved stretches can be noticed
carboxyl-terminal to the putative molybdenum cofactor-binding domain.
Transformation experiments reported above have shown that at least some
substrate specificity mutations map in this region. Thus, we propose
that at least some of the conserved amino acids carboxyl-terminal to
the putative molybdenum cofactor-binding domain participate in
substrate binding. We observe that some residues that are conserved in
rats, mice, human sequence H1, D. melanogaster, Drosophila
pseudoobscura, and A. nidulans are not conserved in the
other reported human sequences (H2 in Fig. 4). The sequence
ERXXXH (A. nidulans positions 910-915) is
universally conserved, except for H2 (Wright et al., 1993),
which has a EMXXXK sequence. Mutagenesis experiments to be
reported elsewhere have shown that conserved amino acids in the stretch
ERXXXH at positions 910-915 of A. nidulans are
actually involved in determining substrate specificity. This sequence is within plasmid pBAN882C, which has been shown to
contain the sequences altered in three substrate specificity mutations, hxA101, hxA102, and hxA143.
The proposed domain organization of the A. nidulans xanthine dehydrogenase is shown in Fig. 6. On the whole, the analysis of the A. nidulans sequence confirms the domain organization proposed previously (Hughes et al. 1992a, 1992b; Wootton et al., 1991), pinpoints a number of residues conserved in organisms as far apart as metazoans and fungi, and proposes a consensus for the molybdenum cofactor-binding domain.
Figure 6: Putative functional domains and intron position in the xanthine dehydrogenases. I and II indicate the iron-sulfur centers. F indicates amino acids involved in the binding of FAD, N indicates those involved in the binding of NAD, and M indicates amino acids involved in complexing the molybdenum cofactor according to the consensus sequence drawn in Fig. 5. S indicates a region of high similarity between all xanthine dehydrogenases corresponding to the region where the substrate specificity mutations map in the enzyme of A. nidulans (see ``Results and Discussion''). F and N are ``minimal'' domains (see ``Results and Discussion''), as the FAD and NAD domains may well overlap. The S region may be extended or limited as more substrate specificity mutations are located and sequenced. Short arrows, positions of introns in A. nidulans; double-headed arrows, positions of introns present in the three sequenced genes from Diptera; long single-headed arrow, intron present in both Drosophila species; long single-headed dashed arrow, intron present in D. pseudoobscura and C. vicina, but not in D. melanogaster.
The domain organization favored
by us and others (Wootton et al., 1979; Hughes et
al., 1992a, 1992b) on the basis of sequence comparisons may seem
to conflict with the results obtained with the chicken xanthine
dehydrogenase by Coughlan et al. (1979). These authors have
shown that one product of subtilisin digestion carries the
molybdenum-binding domain and the iron-sulfur centers, but not the FAD
domain. The contradiction is only apparent, as this
``M 65,000 domain'' obtained by
subtilisin digestion has been shown to be composed of several peptides
that stick together and are copurified. Such a complex could reflect
the tertiary rather than the primary structure of the protein. Magnetic
coupling studies of the native enzyme imply that the molybdenum and
iron-sulfur domains lie near each other (8-14 Å) (Lowe et al., 1972; Lowe and Bray, 1978; Cottman and Buettner,
1979). This proposal is consistent with the more recent results of
Amaya et al.(1990) on the rat liver enzyme.
The three
introns interrupting the A. nidulans hxA gene are not in the
same positions as the introns of the genes sequenced in Diptera. It
would be particularly interesting to compare the A. nidulans intron positions with the mammalian ones, as on the whole, the A. nidulans enzyme is more similar to the mammalian than to
the insect sequences. Unfortunately, no mammalian xanthine
dehydrogenase genomic sequences are known. The introns of the A.
nidulans hxA gene interrupt putative functional domains: the first
and second interrupt the iron-sulfur domains, and the third interrupts
the putative molybdenum cofactor-binding domain. This intron is in the
-G interval (Fig. 5), which is the more variable interval
in the proposed molybdenum cofactor-binding domain. The positions of
the introns in the A. nidulans and insect enzymes are shown in Fig. 6.
While the human H2 enzyme shows marginally higher overall similarity to the D. melanogaster enzyme than to the A. nidulans enzyme, H2 differs from all other enzymes in several crucial sequences besides those in the putative NAD- and reducing substrate-binding sites (see above). The AKL putative peroxisomal domain is absent in this enzyme (residues 292-294 in the A. nidulans sequence). A hydrophobic residue (alanine in the mammalian and A. nidulans sequences and glycine in the dipteran sequences) in the putative FAD-binding domain is substituted with an arginine (this is the residue that is substituted with an aspartic acid in a mutant of D. melanogaster; see below). Three other universally conserved residues in this domain, a glutamine (position 388), a serine (position 393), and an isoleucine (position 395), are substituted with histidine, histidine, and aspartate, respectively. H2 is the only enzyme described that does not have a tyrosine in the putative NAD-binding domain (position 429 in the enzyme of A. nidulans); this residue is substituted with a cysteine. Other universally conserved residues, flanking this tyrosine, are also different in this human enzyme. On the other hand, the putative iron-sulfur centers and molybdenum cofactor-binding domains are well conserved in H2 (but see Fig. 5, which shows differences between H2 and all other xanthine dehydrogenases in the putative molybdenum cofactor-binding domain). We would like to propose that all the sequences, including the human enzyme H1 described by Ichida et al.(1993), but not the human enzyme H2 described by Wright et al.(1993), correspond to the same enzyme with identical or very similar substrate specificity. A number of enzymes called purine hydroxylases or, more imprecisely, ``aldehyde oxidases'' have been described. Krenitsky et al.(1972, 1974) have shown that the aldehyde oxidase from rat liver is very similar to the xanthine dehydrogenases, but differs in substrate specificity. These authors defined operationally aldehyde oxidases as enzymes able to hydroxylate 6-methylpurine, but not xanthine, while the xanthine oxidases (and by implication, the dehydrogenases) accept xanthine, but not 6-methylpurine as a substrate. By this criterion, mammals, including humans, have both a xanthine dehydrogenase (or oxidase) and an aldehyde oxidase. Moreover, most aldehyde oxidases, including the human liver enzyme, do not accept NAD as the oxidizing substrate (Krenitsky et al., 1974). This could well account for some of the sequence differences noted above. We propose that the human enzyme that has been cloned and sequenced by Wright et al. is another molybdenum-containing hydroxylase, related to but different from the classical xanthine dehydrogenases, and we anticipate that this enzyme will show different reducing and perhaps also oxidizing substrate specificities. As this enzyme has been isolated from a human liver cDNA, it may well be the human liver aldehyde oxidase described by Krenitsky et al.(1974).
It is interesting to look again at the missense D. melanogaster mutations now that the sequence of a non-metazoan enzyme is known. If we exclude the aberrant human H2 enzyme, all the missense mutations known (Hughes et al., 1992b) affect universally conserved residues. The only apparent exception is a conservative change, an alanine in the putative FAD-binding domain of the D. melanogaster enzyme (position 353) that is changed to aspartic acid in one of the mutants. At variance with the three diptera, the mammalian and A. nidulans enzymes have a glycine in this position.
The cloning and sequencing of the hxA gene of A. nidulans will allow the use of all the panoply of genetics and reverse genetics techniques to dissect the electron transport chain, the determinants of substrate specificity, the dimerization domains, and the region of interaction with post-translational modification functions, such as that coded by hxB. The strict conservation of the enzyme structure throughout evolution ensures that these studies will have general significance.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBank(TM)/EMBL Data Bank with accession number(s) X82827[GenBank].
Addendum-While this article was being reviewed, a third cDNA human xanthine dehydrogenase sequence was published (Xu et al., 1994). The peptidic sequence is extremely similar (but not identical) to the enzyme sequence reported by Ichida et al. (1993) (H1 in Fig. 4). Xu et al. propose, as we do above, that the cDNA sequence determined by Wright et al.(1993) (H2 in Fig. 4) cannot correspond to a xanthine dehydrogenase. However, at variance with Xu et al., we propose (Fig. 5) that the cDNA sequence of Wright et al. represents an enzyme containing a molybdenum cofactor-binding domain.