Cloning, Sequencing, and Heterologous Expression of the Murine Peroxisomal Flavoprotein, N1-Acetylated Polyamine Oxidase*

Tianyun Wu, Victoria Yankovskaya and William S. McIntire {ddagger}

From the Molecular Biology Division of the Department of Veterans Affairs Medical Center, San Francisco, the Northern California Institute for Research and Education, San Francisco, California 94121, Department of Biochemistry and Biophysics, University of California, San Francisco, California 94143

Received for publication, August 27, 2002 , and in revised form, March 18, 2003.
    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The aminoacyl sequences of three regions of pure bovine N1-acetylated polyamine oxidase (PAO) were obtained and used to search GenBankTM. This led to the cloning and sequencing of a complete coding cDNA for murine PAO (mPAO) and the 5'-truncated coding region of the bovine pao (bpao) gene. A search of GenBankTM indicated that mpao maps to murine chromosome 7 as seven exons. The translated amino acid sequences of mpao and bpao have a –Pro-Arg-Leu peroxisomal targeting signal at the extreme C termini. A {beta}-{alpha}-{beta} FAD-binding motif is present in the N-terminal portion of mPAO. This and several other regions of mPAO and bPAO are highly similar to corresponding sections of other flavoprotein amine oxidases, although the overall identity of aligned sequences indicates that PAO represents a new subfamily of flavoproteins. A fragment of mpao was used as a probe to establish the relative transcription levels of this gene in various mature murine tissues and murine embryonic and breast tissues at different developmental stages. An Escherichia coli expression system has been developed for manufacturing mPAO at a reasonable level. The mPAO so produced was purified to homogeneity and characterized. It was demonstrated definitively that PAO oxidizes N1-acetylspermine to spermidine and 3-acetamidopropanal and that it also oxidizes N1-acetylspermidine to putrescine and 3-acetamidopropanal. Thus, this is the classical polyamine oxidase (EC 1.5.3.11 [EC] ) that is defined as the enzyme that oxidizes these N1-acetylated polyamines on the exo-side of their N4-amino groups. This enzyme is distinguishable from the plant polyamine oxidase that oxidizes spermine on the endo-side of the N4-nitrogen. It differs also from mammalian spermine oxidase that oxidizes spermine (but not N1-acetylspermine or N1-acetylspermidine) at the exo-carbon of its N4-amino group. This report provides details of the biochemical, spectral, oxidation-reduction, and steady-state kinetic properties of pure mPAO.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Spermine (SPM)1 and spermidine (SPD) are important polyamines required for numerous fundamentally important cellular processes including wound healing, tissue growth, tissue differentiation, and tumor growth (1, 2, 3, 4, 5, 6, 7, 8, 9). In mammalian cells, polyamine pool homeostasis is maintained by a balance of enzymatic and transport processes as follows. (a) Putrescine (PUT) is derived from ornithine via ornithine decarboxylase and (b) condenses with decarboxylate S-adenosyl-L-methionine to yield SPD by the action of SPD synthase. (c) SPM is generated from SPD and decarboxylate S-adenosyl-L-methionine via SPM synthase. (Decarboxylated S-adenosyl-L-methionine is the product of S-adenosyl-L-methionine decarboxylase.) (d) Acetyl-CoA:SPD/SPM N1-acetyltransferase (SSAT) transforms SPM and SPD into N1-acetyl-SPM and N1-acetyl-SPD. (e) N1-Acetylated polyamine oxidase (PAO; EC 1.5.3.11 [EC] ) converts N1-acetyl-SPM to SPD and 3-acetamidopropanal, and it converts N1-acetyl-SPD to PUT and 3-acetamidopropanal (Fig. 1). It also oxidizes inefficiently SPM to SPD and 3-aminopropanal. (f) Spermine oxidase (SMO) oxidizes SPM to SPD and 3-aminopropanal (10, 11, 12) but is unable to oxidize N1-acetyl-SPM or N1-acetyl-SPD. (g) The N1-acetylated polyamines are transported from cells to the blood and then to the kidneys for urine excretion. (h) PUT, SPD, and SPM derived from ingested foods are efficiently transported into cells (1).



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FIG. 1.
The oxidation of N1-acetyl-SPM and N1-acetyl-SPD by the mammalian peroxisomal flavoprotein PAO. The figure shows that the oxidation of enzyme-bound N1-acetyl-SPM produces 2-electron reduced FAD and 1 mol/mol each of 3-acetamidopropanal and SPD. The oxidation of reduced FAD by O2 produces 1 mol/mol of H2O2. N1-Acetyl-SPD is oxidized similarly to produce 1 mol/mol each of PUT, 3-acetamidopropanal, and H2O2. The structures of N1-acetyl-SPD/PUT differ from those of N1-acetyl-SPM/SPD by the lack of 3-aminopropyl groups. (The dashed line differentiates the structures and reactions for the two substrates.) The numbering system used in this paper is shown with the upper left structure. The endo- and exo-carbon centers discussed herein are indicated in the middle left structure. R represents the ribityl-ADP portion of FAD.

 

Peroxisomal mammalian PAO contains noncovalently bound FAD as the redox cofactor. When PAO oxidizes N1-acetyl-SPM or N1-acetyl-SPD, FAD is reduced, which is oxidized subsequently by O2 to generate H2O2 (1) (Fig. 1). Thus, PAO has been proposed to play a role in triggering and/or participating in the progression of apoptosis (13, 14, 15, 16, 17). Additionally, another product of the oxidation of the N1-acetylated polyamines, 3-acetamidopropanal, may be enzymatically deacetylated to yield cytotoxic 3-aminopropanal (18) that is thought to contribute, either alone or in concert with H2O2, to tissue damage following traumatic injury (19, 20, 21).

Prior to the present study, the gene sequences of four other forms of polyamine-oxidizing flavoproteins were known. These are the N1-acetyl-SPD oxidase from Candida boidinii peroxisomes (22), the SPM/SPD oxidase from corn (cPAO) (23), and (apparently) cytosolic human (10, 12) and murine SMO (11).2 These flavoproteins have similarities with the murine PAO (mPAO) and bovine PAO (bPAO) described herein. In particular, all have an easily identified FAD-binding motif near their N termini. This and other features of the primary structures indicate that these enzymes are members of a larger family of amine oxidases that includes mitochondrial monoamine oxidase A and B (26), MAO-N from Aspergillus niger (27, 28), PUT oxidase from Micrococcus rubens (29), and tyramine oxidase from Micrococcus luteus (30). (MAO-A and MAO-B both contain covalently bound FAD (24, 25), whereas other members of this family harbor noncovalently bound FAD.) These amine oxidases belong to an even larger enzyme superfamily (the glutathione reductase (GR) superfamily) that includes various disulfide reductases (GR1 family), glucose oxidase, fumarate reductase, cholesterol oxidase, D-amino acid oxidase (GR2 family, for which PAO is a member), protoporphyrinogen oxidase, and phytoene desaturase (31, 32). Of the amine oxidases of this superfamily, only the structures of cPAO (26) and MAO-B are known (33).

Although PAO has significant clinical and pharmacological relevance pertaining to cancer, ischemic tissue damage, apoptosis, etc., there is a paucity of solid data regarding the biochemical properties, mechanism of substrate oxidation, mechanism of inhibition by highly selective compounds such as N1,N4-bis(butadienyl)-1,4-diaminobutane (MDL 72527) and N1-butadienyl-1,4-diaminobutane (MDL 72521) (1), or structural information for any mammalian PAO. This situation prompted us to initiate a program to create a system to produce PAO heterologously. This paper reports the cloning and the complete sequencing of murine pao (mpao), the sequencing of all but a small portion of bovine pao (bpao), and the production of active mPAO by Escherichia coli at a reasonable level. Various aspects of UV-visible biochemical, spectral, redox, and steady-state kinetic properties of the heterologously produced pure mPAO are presented below.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials— The chemicals and vendors are as follows: SPD trihydrochloride (Calbiochem, gold label); benzylamine hydrochloride, Coomassie Blue, SPM tetrahydrochloride, PUT dihydrochloride, 4-aminoantipyrine, vanillic acid, horseradish peroxidase (type II), MOPS, HEPES, flavin adenine dinucleotide (FAD), bovine liver catalase, and dansyl chloride (Sigma); N1-acetyl-SPM trihydrochloride and N1-acetyl-SPD dihydrochloride (Fluka Chemika); A. niger glucose oxidase (Miles Laboratories, Inc.); acrolein, 1-amino-3,3-diethoxypropane, proline (+99%), 2,4-dinitrophenylhydrazine (moist solid, 30–35% water) (Acros Organics); N-(3-aminopropyl)-1,10-diamindecane trihydrochloride, N1,N12-bis(ethyl)-SPM dihydrochloride (BESPM), and N1,N11-bis(ethyl)-nor-SPM dihydrochloride (BENSPM) (Tocris Cookson); [32P]ATP (Amersham Biosciences). Silica gel (HETLC-GHL) and Avicel (cellulose, 250 µm) TLC plates were from Analtech. Silica gel, for flash chromatography (40-µm particle size), was from J. T. Baker, Inc. All other chemicals, purchased from common vendors, were of reagent grade or better. Molecular Biology reagents were from various vendors as follows: ethidium bromide (Amersham Biosciences), restriction enzymes (Invitrogen and New England Biolabs), and isopropyl thio-{beta}-D-galactoside (Amersham Biosciences), for example.

Analytical Procedures—Electrospray ionization mass spectrometry (ESI-MS; positive-ion mode and negative-ion mode) and elemental analyses of organic chemicals were done by HT Laboratories, San Diego, CA. One-dimensional 300 MHz (Nicolet/GE NT 300) 1H NMR spectra were obtained from Acorn NMR, Inc., Livermore, CA. The liquid chromatography-electrospray ionization mass spectral analysis of pure mPAO was done using a Finnigan LCQ Deca XP (ion trap mass spectrometer). The work was performed at the State University of New York Health Science Center Mass Spectrometry Facility (Brooklyn, NY). Uncorrected capillary tube melting points were determined using a Misco aluminum block device.

Synthesis of 1-Acetamido-3,3-diethoxypropane—1-Amino-3,3-diethoxypropane (the diethyl acetal of 3-aminopropanal), 6.5 ml (5.9 g, 0.04 mol), was dissolved in 50 ml of dry pyridine at 0 °C. Over a 25-min period, with stirring, 6.7 ml (7.25 g, 0.071 mol) of acetic anhydride were added dropwise. The mixture was warmed to room temperature and stirred overnight. A small spot of the reaction mixture, dried on an Avicel TLC plate, was sprayed with a solution of ninhydrin (0.1% w/v in n-butyl alcohol). Heating the plate for several minutes at 200 °C indicated that all of the material had been acetylated. The pyridine, acetic acid, and acetic anhydride were removed in a rotary evaporator under high vacuum at 50–65 °C to yield 7.86 g of material (expected yield 7.57 g). The recovered material was distilled under vacuum (0.45 mm Hg); b.p. 111 °C, 6.88 g, 91% yield. 1H NMR, CDCl3 (ppm relative to tetramethylsilane), 7.29 (0.98, s, amide-NH), 4.57 (1.00, t, propyl-CH), 3.68 (2.02, m, ethyl-CH2), 3.52 (2.05, m, ethyl-CH2), 3.35 (2.07, m, propyl-CH2), 1.95 (2.92, s, amide-CH3), 1.82 (2.04, m, propyl-CH2), 1.22 (6.00, t, ethyl-CH3). To our knowledge this compound has not been described previously.

Synthesis of the 2,4-Dinitrophenylhydrazone of 3-Acetamidopropanal—1-Acetamido-3,3-diethoxypropane, 250 mg (1.3 mmol), was mixed with 1.0 ml of 1.5 N HCl. After 2–3 min, this was added to a boiling solution of 5 ml of ethanol/0.5 ml of concentrated HCl containing 295 mg of 2,4-dintrophenylhydrazine (1.5 mmol). Heating was stopped immediately, and 10 ml of room temperature ethanol were added. In a few minutes, the 2,4-dinitrophenylhydrazone began to crystallize. After 1 h at room temperature and 1 h on ice, the solid was filtered and washed with a small volume of ice-cold ethanol. The yield of the 2,4-dinitrophenylhydrazone of 3-acetamidopropanal was 335 mg (1.13 mmol, 86% yield); m.p. 157.5–158 °C. 1H NMR, d6-Me2SO (ppm relative to tetramethylsilane), 11.35 (0.95, s, hydrazone-NH), 9.82 (0.86, d, aromatic-H), 8.33 (1.01, d-d, aromatic-H), 7.97 (1.91, m, propyl-CH and amide-H), 7.86 (1.04, d, aromatic-H), 3.31 (~2, m, propyl-CH2, overlaps with H2O peak), 2.49 (~2, m, propyl-CH2, overlaps with Me2SO peaks), 1.81 (2.97, s, amide-CH3); ESI-MS, m/z, 294 [M — H] (negative ion mode), 296 [M + H]+, 237 [M — acetamido]+ (positive ion mode).

Purification of bPAO and Amino Acid Sequencing of Segments of This Enzyme—Bovine livers were covered with ice and transported to the laboratory within 1 h after their removal from live animals at a local slaughterhouse. Following a published procedure (34), bPAO was purified from fresh liver, or tissue that had been cut into 1-inch cubes from fresh liver and immediately frozen and stored at —80 °C. About 1 mg of nearly pure bPAO was obtained from 1 kg of tissue. The enzyme was purified further on a 10% Tris-HCl SDS "Ready Gel" (Bio-Rad), then electro-transferred onto an Immobilon-PSQ membrane (Millipore), and Coomassie Blue-stained. The membrane was submitted to the Biomolecular Resource Center (University of California, San Francisco, CA) for the N-terminal sequence analysis.

Purified bPAO was electrophoresed as before, and the Coomassie Blue-stained bPAO band was excised and subjected to an in-gel tryptic digest at the Protein Sequencing Center at the State University of New York, Brooklyn, NY. Two major internal peptides (Peptide I and Peptide II; see Fig. 2) were purified and sequenced.



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FIG. 2.
The DNA and translated protein sequences for mPAO. The overlined segment represents the 5'-segment that was missing from the original truncated mpao1 clone. This sequence was obtained by using the 5'-rapid amplification of cDNA ends PCR method. The double underlined nucleotide sequence denotes the region corresponding to the antisense primer (bases 333 to 309) used for the PCR 5'-extension experiment. The underlined portions of the amino acid sequence correspond to the regions of bPAO that were sequenced by the Edman degradation method, and the asterisk denotes the stop codon.

 

Cloning and Sequencing of bpao and mpao—GenBankTM EST data bases were searched using three bPAO peptide sequences (see above). Two murine ESTs (GenBankTM accession numbers AA437705 [GenBank] and AI098814 [GenBank] ) were found to code for aminoacyl sequences that were ~85% identical to that of bPAO Peptide I (see Fig. 2). Both clones (IMAGE numbers 819909 and 1482295, respectively) were purchased from Genome Systems, Inc. (St. Louis, MO), and plasmids were isolated using a Qiagen kit and sequenced. (All DNA sequencing work was done at the Biomolecular Resource Center, University of California, San Francisco, CA). The AA437705 [GenBank] cDNA fragment (mpao1) was released from its plasmid by an XbaI/SalI digestion. This fragment was the template for generating a mixture of [32P]dATP-labeled probes, by using a random-primed DNA labeling kit (Roche Applied Science) (35). The 32P-labeled probes were used to screen a {lambda}gt 10-mouse 17-day embryo cDNA library (Clontech) and a Uni-ZAP XR bovine liver cDNA library (Stratagene). Seventeen positive phage plaques from the bovine cDNA library and 16 positive plaques from the mouse cDNA library were obtained and rescreened. The final positive clones, containing different length mpao and bpao cDNA inserts, were isolated and confirmed by standard Southern blotting (35) using the same 32P-labeled probes. The largest mpao fragments from these clones were sequenced. A fragment of bpao cDNA was excised from the Uni-ZP XR vector of the pBluescript phagemid. This 1.6-kb fragment (bpao1) was sequenced using flanking primers (T3/T7), and it coded for all but a small section of the N-terminal portion of bPAO. The 5'-end of the fragment coded for a sequence that matched exactly the C-terminal portion of Peptide I (see Fig. 2). A region near the 3'-end coded for a protein sequence identical to that of Peptide II, which was determined later to be the C terminus of the enzyme (see Fig. 2). The high similarity between the sequences of bpao1 and mpao1 confirmed that the mpao1 screening probe codes for a portion of mPAO.

DNA isolated from one plaque of the mouse cDNA library was cloned into the SalI sites of pUC19 to give the plasmid, pUC19_MPAO1, that was used for double-stranded sequencing. The sequence of this fragment (mpao2) was missing the 5'-end of the complete mpao sequence (see under "Results").A5'-extension (see Fig. 2) was obtained using the 5'-rapid amplification of cDNA ends PCR method with mouse 17-day embryo Marathon Ready cDNA (Clontech) as the template and using a SMARTTM PCR cDNA Synthesis kit (Clontech) for the PCRs. The mpao gene-specific antisense primer, mpao1R (5'-GTTCTCTTCCGATAATTCTTTCTCC-3'), spans nucleotides 333 to 309 of mpao (see Fig. 2), and the Clontech AP1 universal sense adaptor primer was specific for the Marathon Ready cDNA; 5 cycles for 30 s at 94 °C and 3 min at 72 °C, 5 cycles for 30 s at 94 °C and 3 min at 70 °C, and 30 cycles for 20 s at 94 °C and 3 min at 68 °C. By using a 50-fold dilution of the resulting PCR product as template, and AP2 (an AP1-nested primer) and mpao1R as primers, a second PCR was carried out under the same conditions. The resulting cDNA fragment, about 400 bp long, was isolated from a 1% agarose gel using a Geneclean kit (Bio 101). Melded with mpao2, the sequenced PCR product provided a 1.7-kb section of mpao that coded for full-length mPAO.

Measuring the Relative Levels of mpao mRNA in Different Murine Tissues—Murine Rapid-Scan Gene Expression Panels were purchased from OriGene Technologies, Inc. Two gene-specific primers were designed according to the manufacturer's instruction. The sense primer, mpao2F (5'-TCGGAAGAGAACCAGCTTGTGG-3', 22-mer), and the antisense primer, mpao2R (5'-CAATGACATGATGTGCAGGCA-3', 22-mer), generated a 570-bp-long mpao cDNA fragment by PCR. The 24 mouse cDNA samples, serially diluted over a 4-log range (x1000, 100, 100, and 1) by the manufacturer, were arrayed into a 96-well PCR plate. The first step of the PCRs were carried out at 94 °C for 3 min, which was followed by 35 cycles: 94 °C for 30 s, 55 °C for 1 min, and 72 °C for 2 min. The control-primer pair for detection of {beta}-actin cDNA, provided by the manufacturer, was used for a PCR that was carried out as just described with 25 cycles. The amplified fragments were electrophoresed on a 1% agarose gel and ethidium bromide-stained to provided a measure of mpao mRNA in each tissue.

Expression of mpao in E. coli—The pET 29 c(+) vector (Novagen) was used to construct a mpao prokaryotic expression system, and E. coli DH5{alpha} was used for plasmid subcloning. First, a 5'-end fragment was generated by PCR using mpao1 as the template for the gene-specific antisense primer mpao1R (see above), and a sense primer, mpao1F, which contains SacI and NdeI sites and an ATG start codon (5'-GCGAGCTCATACATATGGCGTTCCCTGGCCCGCGG-3'). The underlined regions indicate SacI and NdeI sites, respectively. A SacI/BamHI fragment of the PCR product was subcloned into pUC19-MPAO1 to give pUC19-MPAO. This construct contained the entire mpao gene. Next, the full-length mpao cDNA was ligated into NdeI and HindIII sites of pET 29c to give a plasmid denoted pET-MPAO. E. coli BL21 GOLD (DE3) (Invitrogen) was transformed with this plasmid for mPAO production.

Growth of Transformed Bacteria—A culture of the E. coli transformant was grown on Luria-Bertani (LB) agar plates containing 30 µg/ml kanamycin. A single positive colony was inoculated into 3 ml of LB broth containing 30 µg/ml kanamycin (LB-kan) for overnight growth at 37 °C. This culture (500 µl) was transferred to 80 ml of fresh LB-kan medium for overnight growth. Five milliliters of this culture were transferred to each of five 2-liter flasks containing 1 liter of fresh LB-kan medium for overnight growth at 37 °C, with shaking. Each flask was added to one of five 14-liter New Brunswick FS-614 fermentors containing 12 liters of LB-kan media. The cell culture was incubated at 30 °C with rapid stirring and vigorous aeration. When the A600 of the culture reached 0.6–0.7, isopropyl thio-{beta}-D-galactoside was added (final concentration, 50 µM). Growth was continued until the A600 reached 1.5–2.0. About 260 g of centrifuged cell paste were obtained from the 60 liters of growth media. The paste was stored at —80 °C.

Purification of Heterologously Produced mPAO—Selected fractions for the various steps in the purification were assayed for N1-acetyl-SPM oxidase activities by a published method (36). This assay measured the time-dependent formation of H2O2 (Fig. 1). The assay stock solutions are as follows: (A) 100 mM vanillic acid, pH 7.0 with KOH; (B) 50 mM 4-aminopyrine; (C) 400 units/ml horseradish peroxidase; (D) 50 mM N1-acetyl-SPM; (E) 100 mM glycine/KOH, pH 9.5, the pH for maximal activity (37). Thirty microliters each of solutions A–D were mixed with 2.88 ml of solution E, and 50 µl of the resulting solution were pipetted into individual wells of a 96-well plate. Anywhere from 1 to 50 µl of a particular fraction was added to a well. The relative activities of different fractions were assessed visually from the time-dependent intensity change of the developing pink color. The purity of various fractions were assessed by SDS-PAGE using pre-cast 10–20% Tris-HCl "Ready Gels" (Bio-Rad), following the manufacturer's instructions.

Unless noted otherwise, all purification steps were carried out at 4 °C. Frozen E. coli cell paste (260 g) was thawed in a beaker with 10 mM MOPS buffer, pH 7.25. (The pH was adjusted at 21 °C; the estimated pH at 4 °C is 7.35). The 800-ml suspension was homogenized with a large glass/Teflon piston (Potter/Elvehjem) tissue grinder, and then passed twice through an Avestin Emulsiflex C5 Homogenizer at 15–20,000 pounds/square inch. Next, 15 mg of solid FAD were dissolved in the suspension, and it was centrifuged (50,000 x g, for 30 min). The supernatant was dialyzed in the dark against 13 liters of 10 mM MOPS buffer, pH 7.25, for 4 h and then overnight against 13 liters of fresh buffer. The resulting solution was diluted to 2 liters with this buffer and applied to a 14 x 25-cm DEAE-cellulose (Whatman, DE53) column with a flow rate of 20 ml/min. The column was then washed with 2 liters of the buffer and eluted with an 8-liter gradient from 0 to 400 mM KCl in the 10 mM MOPS, pH 7.25 buffer. Activity eluted from 4.7–7.8 liters after the gradient was initiated. The 3.1-liter volume was reduced to ~500 ml using 350 ml of Amicon pressure concentrators fitted with Amicon YM-10 membranes. After dissolving 15 mg of FAD, the resulting solution was dialyzed in the dark for 4 h against 13 liters of 10 mM HEPES buffer, pH 7.8 (pH adjusted at 21 °C; estimated pH at 4 °C was 8.05), and then overnight against 13 liters of fresh buffer. The resulting sample was applied to a 5 x 39-cm DEAE-Spherodex LS column (100–300 µm bead size; Ciphergen) already equilibrated with the 10 mM HEPES buffer. The column was washed with 500 ml of this buffer before starting a 2.4-liter gradient from 0 to 500 mM KCl in the same buffer. The column, with a 7-foot pressure head, was run at the maximum flow rate. Once the gradient was started, 26-ml fractions were collected. The majority of the activity eluted in tubes 82–108, which were combined (~700 ml) and concentrated to ~50 ml as described earlier. This solution was dialyzed for 4 h, against 7 liters of 10 mM KH2PO4/KOH buffer, pH 7.2, and then overnight against 7 liters of fresh buffer.

The sample was chromatographed on a Mono P HR 5/20 column (Amersham Biosciences) at room temperature. After injecting 2 ml of the sample at a flow rate of 1 ml/min with solution I (H2O), proteins were eluted with the following gradient: 0–1% II (1 M KH2PO4/KOH, pH 7.2) in 4 min; 1–30% II in 125 min. mPAO, eluting from 38 to 41 min, was collected as a single fraction and immediately put on ice. This procedure was repeated until the entire sample had been processed. The mPAO fractions from all of the Mono P runs were combined, then concentrated, and washed into 1 mM KH2PO4/KOH buffer, pH 7.2, using 2-ml Centricon-10 centrifuge concentrators (Amicon). The final volume was 2 ml in the 1 mM buffer.

This sample was chromatographed on a 1 x 10-cm ceramic hydroxyapatite (Bio-Rad, type II) column (Amersham Biosciences HR 10/10 column), at room temperature. The mPAO sample (100 µl), diluted to 1 ml with H2O, was injected immediately onto the hydroxyapatite column with a flow rate of 2 ml/min. The elution was carried out as follows: 0% solution II for 7 min; 0–1% II in 2 min; hold at 1% II for 10 min. mPAO eluted as a broad peak from 14 to 17 min. This step was repeated until the entire sample had been processed. The combined fractions were concentrated as for the Mono P fraction. The solution was washed into 10 mM KH2PO4/KOH buffer, pH 7.2, to give a solution that was 3.68 mg/ml mPAO (based on an {epsilon}458 = 10,400 M1 cm1 and a molecular mass = 56,101 Da for the enzyme; see under "Results"). The enzyme was judged pure by SDS-PAGE, by ion-exchange chromatography on an analytical TSK DEAE 2SW column (0.4 x 25 cm; a 0.75 ml/min flow rate, with a gradient from 1 to 50% solution II in 30 min; a single sharp peak eluted at 23 min), and gel filtration chromatography on a TSK 3000SW column (0.7 x 30 cm; 0.5 ml/min flow rate; 250 mM KH2PO4/KOH buffer, pH 7.2). The purity and integrity of the protein was confirmed also by the electrospray ionization (ESI) mass spectral analysis, which provided a peak of 55,311 ± 6 mass units (the mass of apo-mPAO based on the sequence is 55,316 mass units). The yield of pure mPAO was 36.8 mg.

By using the conditions for the steady-state kinetic assay described below, it was found that the enzyme, at 2–4 mg/ml, was stable when frozen at —20 or —80 °C and thawed through several cycles. However, at a concentration of 30 µg/ml, activity was lost quickly after several freeze/thaw cycles, with more rapid loss occurring at —80 °C. When 33% (v/v) ethylene glycol was added, mPAO was stable for several of freeze/thaw cycles for solutions containing 20 µg/ml to 4 mg/ml, regardless of the storage temperature. It was decided to store the enzyme at —20 °C in the presence of 33% (v/v) ethylene glycol. The enzyme maintains full activity and measured biochemical, redox, Mr, and kinetic properties after 16 months of storage under these conditions. Ethylene glycol removal and buffer exchange were accomplished easily by several concentration/dilution cycles using Centricon-10 centrifuge concentrators.

Binding of FAD to mPAO—The spectrum of a 0.1 mg/ml (0.8 ml) solution of mPAO in 10 mM KH2PO4/KOH buffer, pH 7.2, indicated that the sample contained 7.4 nmol of FAD. This solution was treated with 80 µl of 55% trichloroacetic acid (38) and centrifuged to give a clear yellow supernatant and a white pellet. Overnight incubation of the isolated solution at room temperature in the dark resulted in the conversion of the liberated FAD to FMN. Fluorescence analysis with a Hatachi F-4010 fluorescence spectrophotometer (450 nm excitation, 525 nm emission; reference-authentic FMN) indicated that FAD was released quantitatively from mPAO. Thus, FAD is noncovalently bound to mPAO.

Spectral Characterization and Redox Properties of mPAO—All UV-visible spectra were recorded with a Hewlett-Packard 8452A diode array spectrophotometer. mPAO, in 50 mM KH2PO4/KOH buffer, pH 7.6, at 25 °C, was titrated anaerobically with a solution of sodium dithionite. This solution was standardized by using it to titrate anaerobically a FAD solution of known concentration. The anaerobic cuvette and other details of this procedure are described elsewhere (39, 40, 41). The anaerobic mPAO solution contained also 50 mM D-glucose, 3 µg of catalase, and 50 µg of glucose oxidase to scavenge trace dissolved O2. The spectral data were subjected to "Factor Analysis" using the SPECFIT program (Spectrum Software Associates, Chapel Hill, NC) as described earlier (40, 41).

A 1.20 µM solution of mPAO was titrated anaerobically with a solution of 0.5 mM N1-acetyl-SPD in 50 mM KH2PO4/KOH buffer, pH 7.6, at 25 °C. The enzyme and substrate solutions contained 50 mM D-glucose, 3 µg of catalase, and 50 µg of glucose oxidase.

Steady-state Kinetic Experiments—Spectrophotometric assays were done at 30 °C in 50 mM KH2PO4/KOH buffer, pH 7.6, following a published procedure (36). This method provided a continuous monitor of the H2O2 produced in the reactions. The assays were done in 1-ml, 1 cm-path length cuvette with 0.8 ml of solution containing varying amounts of substrate, 0.1–0.2 µg of mPAO, 1 mM vanillic acid, 0.5 mM 4-aminopyrine, and 4 units of horseradish peroxidase. By varying each of the last three components, it was found that none were inhibitory. The reactions were monitored at 498 nm with a UVIKON 840 spectrophotometer (Kontron Instruments) for the formation of the quinoneimine dye ({epsilon} = 4650 M1 cm1 at pH 7.6; see Ref. 36), the condensation product of vanillic acid and oxidized 4-aminopyrine. The latter was produced from 4-aminopyrine by its interaction with horseradish peroxidase that had been oxidized by H2O2 (36). Assays were done by varying the concentration of the amine substrate, whereas the dissolved [O2] was constant at the air-saturated level (237 µM) in the buffer at 30 °C. The data were fit by nonlinear regression (42) to the appropriate steady-state kinetic equations.

The value for the apparent dissociation constant, KD (= KI), for each inhibitor was estimated by measuring the rates of N1-acetyl-SPM oxidation as its concentrations and that of the inhibitor were varied. It was assumed that these substances, which are either very poor or nonsubstrates, are competitive inhibitors for the oxidation of the substrate. These data were analyzed also by nonlinear regression using the appropriate equation.

Steady-state kinetic assays were done also by varying the [N1-acetyl-SPM] in buffer saturated with pure O2 (1.12 mM). After several minutes of bubbling a cuvette solution with a stream of pure O2 gas, a small aliquot of substrate was added followed by the enzyme. Once the enzyme was added, the bubbling was terminated, and the absorbance change was recorded.

Some (oxygraph) assays were carried out by monitoring directly the dissolved O2 consumption in air-saturated buffer (dissolved [O2] = 0.237 mM) or in buffer saturated with pure O2 at 30 °C (dissolved [O2] = 1.12 mM). The dissolved [O2] was measured with a YSI Inc. model 53 oxygen monitor equipped with a Clark-type oxygen electrode in a glass 1.4-ml reaction chamber. The true kcat and KO values for the oxidation of N1-acetyl-SPM and N1-acetyl-SPD were determined by progress curve analyses of reactions that were allowed to go to completion (dissolved [O2] = 0 at t = {infty}). By using a published procedure (43), the data were fitted to the integrated Michaelis-Menten equation. The analyses were done using Maple VI (Windows 2000) software (Waterloo Maple, Inc.) running on a PC computer. By using KI values (see Table I) as a gauge, the saturating concentrations of N1-acetyl-SPM and N1-acetyl-SPD were made high enough (3.7 mM) so that product inhibition by SPD or PUT, respectively, was insignificant at all times during the reaction (see Table I). Inhibition by the H2O2, formed as a product of polyamine oxidation by mPAO, was assessed by addition of 2 µl of 30 mg/ml (30,000 units/mg) solutions of catalase before a reaction was started. The catalase converted immediately each mole of H2O2 formed to 0.5 mol of dissolved O2. After correcting the data by a factor of 2, the rate of O2 consumption was the same as in the absence of catalase. This indicated a lack of inhibition by the H2O2 formed in the assays lacking catalase.


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TABLE I
Steady-state kinetic parameter for the reaction of various amines (S) and O2 (O) with pure mPAO

 

Analyses of the Aldehyde Produced When N1-Acetyl-SPM and N1-Acetyl-SPD Are Oxidized by mPAO—mPAO, 29 µg, was dissolved in 2 ml of 50 mM KH2PO4/KOH buffer, pH 7.6, containing 0.8 mM N1-acetyl-SPM and 30 µg of catalase (30 units). An identical solution containing 0.8 mM N1-acetyl-SPD in the place of N1-acetyl-SPM was also prepared. Each solution was stirred at room temperature for 2 h. After dansylating a small aliquot, HPLC analysis (see below) indicated that the substrates had been oxidized completely for both solutions. Each solution (100 µl) was mixed separately with 100 µl of a 2,4-dinitrophenylhydrazine solution (100 mg in 94 ml of ethanol/6 ml of concentrated HCl). Each of the resulting solutions (25 µl) was injected onto a Prodigy octadecylsilyl silica gel HPLC column (5 µm particle size, 0.46 x 5.0 cm; Phenomenex): flow rate, 1 ml/min; gradient elution 0% B for 0.5 min, 0–35% B from 0.5 to 1.5 min, hold at 35% B from 1.5 to 5.0 min, 35 to 100% B from 5.0 to 9.0 min; solutions A and B were H2O and acetonitrile, respectively, both containing 0.5% (v/v) trifluoroacetic acid. The HPLC system used SpetraSYSTEM P2000 gradient pumps, a UV6000LP Diode Array Detector, and the ThermoQuest ChromQuest Chromatography Data System (Thermo Separation Products). The 368-nm chromatograms were used for quantitative analyses.

Authentic 3-acetamidopropanal, the expected product of mPAO oxidation of N1-acetyl-SPM and N1-acetyl-SPD, was generated by treating 10 µl (~10 mg, 53 µmol) of 1-acetamido-3,3-diethoxypropane with 100 µl of 1.5 N HCl for 1 min and then diluting immediately to 0.8 mM with 50 mM KH2PO4/KOH buffer, pH 7.6. A 0.8 mM solution of 3-aminopropanal was produced similarly after treating 10 µl (9.1 mg, 62 µmol) of 1-amino-3,3-diethoxypropane with 100 µlof1.5 N HCl for 1 min. A 0.8 mM solution of commercial acrolein was also prepared. Each of these solutions was mixed immediately, 1:1 (v/v), with the 2,4-dintrophenylhydrazine reagent and analyzed by HPLC as described in the previous paragraph. Another stock solution containing 0.8 mM of all three aldehydes was treated and analyzed in the same manner. These analyses provided the retention times and reference peak areas for unreacted 2,4-dinitrophenylhydrazine and the 2,4-dinitrophenylhydazones of each aldehyde (see Fig. 6).



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FIG. 6.
HPLC analyses of the aldehyde products generated when N1-acetyl-SPM and N1-acetyl-SPD are oxidized completely by mPAO. The details of the 2,4-dinitrophenylhydrazine derivatizations and the analyses are presented in the "Experimental Procedures." The upper chromatogram is for the analysis of a standards solution that contained 0.8 mM each of 3-aminopropanal, 3-acetamidopropanal, and acrolein (each peak represents 10 nmol of the 2,4-dinitrophenylhydrazone of each aldehyde). The middle and lower chromatograms are for the analyses of the N1-acetyl-SPM and N1-acetyl-SPD reactions, respectively; 0.8 mM substrate at t = 0. Unreacted 2,4-dinitrophenylhydrazine eluted at 4.5 min. The peaks at 8.4 min are due to the 2,4-dinitrophenylhydrazone of acetaldehyde, which is a trace contaminant of the ethanol solution used to dissolve 2,4-dinitrophenylhydrazine. The other minor peaks are of unknown origin.

 

The following experiment was carried out in order to obtain an analytical amount of the aldehyde produced by mPAO oxidation of N1-acetyl-SPM. A 10-ml solution of 50 mM KH2PO4/KOH buffer, pH 7.5, containing 5.0 mM N1-acetyl-SPM (50 µmol or 17.7 mg total), 7.25 µg/ml mPAO, and 7.5 µg/ml catalase was stirred at room temperature. The reaction was monitored by treating 10 µl of the solution with dansyl chloride (see below) and analyzing by TLC; silica gel plates using cyclohexane/ethyl acetate, 2:3 (v/v); Rf values were 0.04 for N1-acetyl-SPM and 0.77 for SPD. After 23 h, there remained no N1-acetyl-SPM. The solution was filtered using Centricon-10 centrifuge concentrators (Millipore), and 1.25 ml of concentrated HCl was added with stirring. Next, 23 mg (115 µmol) of 2,4-dinitrophenylhydrazine in 0.625 ml of tetrahydrofuran were added to the filtrate. After stirring for 1–2 min, the solution became slightly cloudy. The mixture was put on ice for 2 h and filtered, and the solid material was washed with 1–2 ml of ice-cold 1.5 M HCl followed by 6–7 ml of ice-cold water. The yield of the dry yellow solid was 13.2 mg (yield, 89% based on Mr of the 2,4-dintrophenylhydrazone of 3-acetamidopropanal). 1H NMR, d6-Me2SO (ppm relative to tetramethylsilane) 11.36 (0.98, s, hydrazone-NH), 9.82 (0.83, d, aromatic-H), 8.34 (1.01, d-d, aromatic-H), 7.97 (1.95, m, propyl-CH and amide-H), 7.87 (1.04, d, aromatic-H), 3.31 (~2, m, propyl-CH2, overlaps with H2O peak), 2.49 (~2, m, propyl-CH2, overlaps with Me2SO peaks), 1.81 (2.90, s, amide-CH3); ESI-MS, m/z, 294 [M — H] (negative ion mode) 296 [M + H]+, 237 [M — acetamido]+ (positive ion mode). The NMR and mass spectral data indicate that this material is identical to the 2,4-dinitrophenylhydrazone of 3-acetamidopropanal made by chemical synthesis (see above).

Analyses of the Polyamine Products of mPAO Oxidation of N1-Acetyl-SPM and N1-Acetyl-SPD—The 2-h N1-acetyl-SPM and N1-acetyl-SPD enzyme reaction solutions described at the beginning of the previous section were used for these analyses. To 0.5 ml of each solution was added 10 µl of a 15 mM solution of N-(3-aminopropyl)-1,10-diaminodecane trihydrochloride (the internal standard; 0.3 mM final concentration), which was determined to be a mPAO non-substrate. Separate solutions of 0.8 mM N1-acetyl-SPM, N1-acetyl-SPD, SPM, SPD, and PUT, and 0.3 mM N-(3-aminopropyl)-1,10-diaminodecane were prepared. A solution containing 0.8 mM N1-acetyl-SPM, N1-acetyl-SPD, SPM, SPD, and PUT, and 0.3 mM N-(3-aminopropyl)-1,10-diaminodecane was also prepared. The reference solutions established retention times and signal intensities for each compound and the internal standard.

A literature procedure for dansylation with dansyl chloride and sample preparation was used with minor modifications (44). To 50 µl of an unknown or reference solution, in a 1.5-ml screw-cap plastic vial, was added 200 µl of a saturated Na2CO3 solution and 200 µl of a 10 mg/ml dansyl chloride solution in acetone. Each sample was vortex mixed for 20 s before incubation at 65 °C for 10 min. After cooling on ice for several minutes, 100 µl of a proline solution (250 mg/ml) was added, and the sample was vortex mixed for 10 s. The phases were separated by centrifugation, and the upper organic phase (~350 µl) was removed. Ten microliters of this phase were injected onto a Prodigy HPLC column (octadecylsilyl silica gel, 5-µm particle size, 0.46 x 5.0 cm), using a flow rate of 1 ml/min, and the following elution gradient: 0–45% B from 0 to 0.1 min, 45–80% B 0.1 to 8 min, hold at 80% B from 8 to 11 min, 80–90% B from 11 to 12 min. Detection was accomplished with a Gilson Spectra/Glo fluorescence detector using a 7-51X excitation filter (330–400 nm) and a 3-72M emission filter (460–600 nm). The retention times (min) are (data not shown) as follows: N1-acetyl-SPD, 10.4; PUT, 11.5; N1-acetyl-SPM, 13.4; SPD, 14.8; N-(3-aminopropyl)-1,10-diaminodecane, 16.4; SPM, 17.9. Base-line separation of all peaks was achieved.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cloning and Sequencing of mpao and Sequence Analysis— With the hope of studying the properties of a mammalian PAO, this enzyme was purified from bovine liver following a published protocol (34, 45). Although it was reported that ~20 mg of PAO could be obtained from 1 kg of bovine liver, we obtained ~1 mg from this mass of tissue in two separate attempts. In order to carry out careful biochemical and kinetic studies, larger amounts of PAO were required. Therefore, we decided to clone and sequence the gene for a mammalian PAO and attempt to produce the enzyme in a heterologous system. Because a mammalian peroxisomal pao gene was unavailable, in order to clone such a gene, amino acid sequence information was needed. The amino acid analysis of purified bPAO provided the N-terminal sequence, EAEAPGRGPRVLVVGGGIAGL. The underlined segment identifies bPAO as a member of family of FAD-containing oxidases (31) that includes human MAO-A, human MAO-B, corn (maize) polyamine oxidase (cPAO), and an N1-acetyl-SPM oxidase from C. boidinii (Cb Ac-SMO) (22, 23, 26). The GXGXXG sequence is a flavoprotein fingerprint. This motif is present in members the GR1 (glutathione reductase) and GR2 family of flavoproteins, which have known structures (32). MAO-A, MAO-B, cPAO, and several non-amine oxidizing enzymes are known to be of the GR2 structural type.

The sequencing of two tryptic bPAO peptides resulted in the internal protein sequences SEHSFGGVVEVGAHWIHGPS (Peptide I) and LMTLWDPQAQWPEPR (Peptide II). The underlined segment of Peptide I aligns with the end of the FAD-containing enzyme superfamily motif near the N termini of these proteins (31).

Translated EST GenBankTM sequences were screened using these bPAO sequences, and two mouse GenBankTM EST sequences (accession numbers AA437705 [GenBank] and AI098814 [GenBank] ) containing a translated sequence very similar to Peptide I (85% identity) were identified. The plasmid DNA for AA437705 [GenBank] and AI098814 [GenBank] were purified and sequenced. Both clones were truncated at the 5'-end. The DNA from EST clone AA437705 [GenBank] , with a longer mpao insert, was subjected to restriction enzyme digestions and purified. A 968-bp segment (mpao1) was excised from this clone and sequenced. The mpao1 cDNA fragment was used as a library screening probe ("Experimental Procedures"). This process allowed us to clone a nearly complete mpao cDNA (mpao2; 1710 bp) from the mouse embryo library. The sequence of the full-length mpao (1770 bp; Fig. 2) was obtained by combining the sequence information obtained for mpao2 and from a 5'-rapid amplification of cDNA ends PCR procedure. However, we failed to obtain the missing 5'-end of bpao. A search of GenBankTM revealed that mpao maps to murine chromosome 7 (cytogenic position 7F4) as 7 exons (GenBankTM accession number NW_000335).

During the review of the current work, a paper by Vujcic et al. (46) appeared that reported the cDNA sequences for mPAO and hPAO. These sequences were found by BLAST searching GenBankTM using the SMO cDNA sequence. Although the details are not provided herein, we have also cloned and sequenced the hpao gene and submitted its sequence in GenBankTM (accession number AF312698 [GenBank] ) on October 11, 2000. Vujcic et al. (46) made no attempt to sequence the mpao and hpao cDNA from the clones that they obtained commercially. Compared with our cDNA sequences, their sequences differ at numerous positions. We are confident that our bPAO, mPAO, and hPAO sequences are correct because we sequenced each at least twice; ambiguous regions were sequenced three or four times.

Vujcic et al. (46) reported also the transient transfection of HEK-293 cultured human kidney cells with mpao and hpao cDNA. Although these cells expressed the enzymes from the transfected genes, no effort was made to purify the proteins for biochemical characterization (46).

A Comparison of mPAO and bPAO with Each Other and with Other Known Amine Oxidases, and the Identification of These as Peroxisomal Proteins—The full-length mpao cDNA (1770 bp) (GenBankTM accession number AF226656 [GenBank] ) and deduced mPAO amino acid sequences are presented in Fig. 2. Upstream from the ATG start codon, there is a single TAA stop codon. No other possible translational start sites were found upstream from this ATG codon. The mpao coding region terminates with a single TGA stop codon. The gene contains a 1512-bp open reading frame that encodes for 504 amino acids. The mature apoprotein (minus the N-terminal Met) has a mass of 55,316 Da. The incompleted bpao nucleotide (1625 bp) and deduced amino acid (452 amino acids) sequences can be found at GenBankTM (accession number AF226658 [GenBank] ). Its 5'-end nucleotide sequence is missing. The coding region of bpao terminates with a single TGA stop codon. Both mpao and bpao have an ATAAA sequence as polyadenylation signals that are near to the poly(A) tails.

An inspection of the bPAO and mPAO sequences (Fig. 3) indicates the presence of the peroxisomal targeting signal sequence, -PRL, at the C termini of these proteins. This consensus sequence -(S/A/C/P)-(K/H/R)-(I/L/M) (47), which is not cleaved after protein import into the peroxisome, is seen also in the Cb Ac-SMO and MAO-N sequences (i.e. -SKL and -ARL, respectively) (Fig. 3), indicating that these enzymes reside also in the peroxisomes of the respective host yeast cells. Like many oxidases, PAO is localized in the peroxisomes where its oxidation product H2O2 can be degraded by catalase.



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FIG. 3.
Alignment of the protein sequences of various flavoprotein amine oxidases. The alignment and (default) shading was accomplished using ClustalW (version 1.8) within the BioEdit© Program, version 5.0.9 (Dept. of Microbiology, North Carolina State University) (48). Aligned are bovine (peroxisomal) PAO, murine (peroxisomal) PAO, human SMO (GenBankTM accession number AY033889 [GenBank] ) (10, 12), murine SMO (GenBankTM accession number BC004831 [GenBank] ) (11), human MAO-A (GenBankTM accession number M69226 [GenBank] ) (24), human MAO-B (GenBankTM accession number M69177 [GenBank] ) (24), S. gairdneri (trout liver) MAO (fMAO; GenBankTM accession number L37878 [GenBank] ), M. tuberculosis amine oxidase (Mt AmOx; GenBankTM accession number AL021646 [GenBank] ), PUT oxidase from M. rubens (Put-Ox; GenBankTM accession number D12511 [GenBank] ) (29), Zea mays (corn) PAO (GenBankTM accession number AJ002204 [GenBank] ) (23, 26), M. luteus tyramine oxidase (Ml TyrOx; GenBankTM accession number 3298360) (30), A. niger MAO (MAO-N; GenBankTM accession number L38858 [GenBank] ), and C. boidinii N1-acetyl-SPD oxidase (GenBankTM accession number AB018223 [GenBank] ) (22). The question marks for the bPAO sequence indicate a region of unknown composition. The composition of the segment preceding this region was obtained by protein sequencing, whereas the sequence following this region was deduced from the translated cDNA sequence. The cPAO sequence is the only one with a recognizable N terminus transport signal sequence, which is underlined. At the C termini of bPAO, mPAO, MAO-N, and Cb Ac-SMO, the tripeptide peroxisomal transport signals are indicated by asterisks. The position of Cys residues that are covalently linked to the FAD in MAO-A, MAO-B (24, 25), and fMAO are indicated by the {ddagger} symbol. The = symbols above the sequences indicate regions that are highly conserved in this alignment. For example, the regions labeled Beta-1, Alpha, and Beta-2 are components of the {beta}1{alpha}{beta}2 motif near the N termini that interacts with the ADP portion of FAD. The positive end of the {alpha}-helix of this motif interacts with the diphosphoryl group of the ADP moiety. The regions labeled Fl or Flx are in the flavin-binding domains of MAO-B and cPAO (26, 33). These regions constitute elements of the Rossmann fold. The helix that has its positive end interacting with the N1/C2/C2-O locus of FAD is labeled Flx (near the C termini) (33). The regions labeled Sub are conserved regions in the substrate-binding domain, which are remote from the FAD and seemingly remote from the substrate/inhibitor-binding site (26, 33). The extended C-terminal regions of MAO-A, MAO-B, and fMAO anchor these proteins to the outer surface of mitochondrion (33).

 

A ClustalW (version 1.8) alignment of many (but not all) known flavoprotein amine oxidase amino acid sequences is provided in Fig. 3. Among these sequences, there are two regions of high similarity. One, near the N termini, is clearly a {beta}{alpha}{beta} consensus domain that interacts with the ADP moiety of FAD (32, 33, 49). The second conserved region, near the C termini, is involved also in FAD binding. The C-terminal region contains a conserved region that harbors the Cys residues that are covalently linked to FAD in the human monoamine oxidases: Cys406 (MAO-A) and Cys397 (MAO-B) (24, 25). For bPAO and mPAO, a Ser (Ser429 of mPAO) aligns with these Cys residues. We have determined that mPAO bind FAD noncovalently (see under "Experimental Procedures").

The ClustalW analysis (Fig. 3) provided the following percent identities (percent similarities) between mPAO and the other flavin-containing amine oxidases: bPAO, 73% (82%); human and murine SMO, 37% (53%); Micrococcus rubens PUT-Ox, 18% (32%); cPAO, 17% (34%); Salmo gairdneri MAO, 16% (30%); Mycobacterium tuberculosis amine oxidase, 17% (30%); human MAO-B, 15% (30%); human MAO-A, 14% (30%); C. boidinii N1-acetyl-SPD oxidase, 13% (32%); M. luteus tyramine oxidase, 13% (28%); A. niger MAO-N, 12% (25%). Overall, the amino acid sequences identity between mPAO and the other protein is low, generally less than 20%, except for the 37% identity with the newly discovered human and murine SMO. This indicates that peroxisomal PAO represents a new subfamily of mammalian flavoprotein amine oxidases.

The Distribution of mpao mRNA in Murine Tissues—The availability of mpao cDNA allows, for the first time, the determination of the transcription level of this gene in mammalian tissues. We probed PCR-amplified mRNA of murine tissue from numerous organs and murine tissues at different developmental stages (see under "Experimental Procedures"). mpao mRNA is detected in all the murine tissues tested (Fig. 4), with the liver and stomach having the highest levels. This is in accord with an earlier finding of large levels of mPAO in the liver of various mammals (34, 37, 50, 51). Lesser but significant levels of mpao were detected in heart, spleen, thymus, small intestine, muscle, pancreas, uterus, and breast at various developmental stages. Relatively lower levels of mpao mRNA are expressed in brain, kidney, lung, testis, skin, adrenal gland, and prostate gland. The 100x panel (Fig. 4) clearly shows that the mRNA level increases during embryonic development; there is a gradual increase in the tissues on going from 8.5- to 19-day embryos. mRNA levels change also with breast development. Fig. 4 shows that level of mpao mRNA is very low in the virgin breasts, is quite high in the pregnant breasts, but is decreased in lactating and involuting breasts. These findings for breast and embryo were confirmed by repeating this analysis with a murine multiple tissue panel from a different lot (OriGene). Apparently, high mpao transcription and presumably translation is important in tissue growth and development (1). These data suggest that mpao expressions are regulated by growth hormones. It has been proposed that PAO, via its participation in the polyamine interconversion pathway, is an important regulator for maintaining cellular polyamine and tissue homeostasis.



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FIG. 4.
Agarose electrophoresis of "Rapid-Scan Gene Expression Panel" PCR-amplified mpao cDNA samples for 24 major mouse tissues and developmental stages. The right frame presents the results for the PCR-amplified cDNA of a 540-bp portion of the {beta}-actin gene for each tissue, which is the control. The middle frame presents the results for the PCR-amplified cDNA of a 570-bp portion of mpao for each tissue using a high level of first-strand cDNA for the PCR (the 100x panel). The right frame displays the results for the panel using 100x lower first-strand murine cDNA (the 1x panel). The total mRNA of each tissue was subjected to oligo(dT) selection, and the first-strand cDNA used for the PCRs for each tissue were generated from the poly(A+) mRNA using oligo(dT) primers and Moloney murine leukemia virus-reverse transcriptase. The amplified fragments were electrophoresed on an agarose gel, and the intensity of the ethidium bromide-stained bands provided a measure of the level of mpao mRNA in each tissue.

 

Currently, we do not know how these mRNA levels relate to the amount of the mPAO protein in different murine tissues. However, a study measuring PAO enzyme activity in various rat tissues has been reported (1). High activity was found in the pancreas and liver. Lower but significant activity was seen in spleen, kidney, small intestines, testes, prostate tissue, thymus, brain, heart, and lung; and very low activity was observed in skeletal muscle. It was found that PAO activity increased from a low level at birth to quite high levels at 70 days post-natal and beyond in rat brain and liver.

It is important to note that Vujcic et al. (46) reported the relative levels of the transcripts for pao and smo in numerous normal and neoplastic human tissues. For the few normal tissues probed, the agreement with our findings for murine tissues is fair, except that no pao mRNA was detected in normal human spleen.

Heterologous Production of mPAO by E. coli and Characterization of the Pure Enzyme—The cloned mpao gene was heterologously expressed in E. coli, and active mPAO was purified to homogeneity and characterized. PAGE, gel filtration, ion exchange chromatography, and mass spectral analysis indicated that the highly purified, homogeneous enzyme is a monomer of the expected molecular mass. Fig. 5 displays the UV-visible spectrum of the pure oxidized enzyme. The {lambda}max values (and relative absorbances) for the protein are 274 (1.0), 377 (0.09), and 456 (0.11). The calculated {epsilon}274 = 66,000 M—1 cm1 for the protein component, based on the amino acid composition (52), and an estimated {epsilon}274 = 26,000 M—1 cm1 for the bound FAD (assuming 1 mol of FAD/mol of protein) (53) provide an {epsilon}274 of 89,000 M—1 cm1. By using this value, and assuming that the {epsilon}458 is the same as free FAD, i.e. 11,300 M—1 cm1, the estimated molar ratio of protein to FAD is 1:0.90. This supports the contention of 1 mol of noncovalently bound FAD/mol of enzyme. The pI and the molecular mass values for apo-mPAO (minus the N-terminal Met), calculated from the amino acid composition, are 4.84 and 55,316 Da, respectively, whereas the calculated molecular mass of the holoenzyme (FAD-containing) is 56,101 Da. (Because of the negative charges of the phosphate groups of FAD, the pI value is expected to be somewhat lower than 4.84.) ESI mass spectral analysis of purified mPAO gave a molecular mass = 55,311 ± 6 mass units, in perfect agreement with that deduced from the cDNA-translated protein sequence.



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FIG. 5.
The anaerobic dithionite titration of pure mPAO. The titration was done in a 1-ml, 1-cm path anaerobic cuvette, in 50 mM KH2PO4/KOH buffer, pH 7.6, at 21 °C. The concentration of the standardized sodium dithionite solution was 0.541 mM. A shows the spectrum of the oxidized enzyme (- -), those obtained at the beginning of the titration (solid lines; 2.16 and 4.33 nmol of dithionite added), and that of fully reduced enzyme (–––; 17.3 nmol dithionite added). The arrows indicate the direction of the absorbance changes that occurred as more dithionite was added. In the 380-nm region, the increase in absorbance indicates the formation of the red radical, whereas the small increase in the 550–700 nm region indicates the formation of a small amount of the blue radical (54). B displays the spectral changes that occurred in the latter phase of the titration. The arrows indicate the direction of the absorbance changes that took place as progressively more dithionite was added: 4.33, 6.49, 8.66, 10.8, 13.0, 15.1, and 17.3 nmol. Although impossible to see in this reproduction, the absorbance in the 550–700-nm region increased slightly and then decreased during this phase of the titration. The inset to B shows a graph of A377, A458, and A590 versus the amount of dithionite added. From this plot, it was determined that 15.2 nmol of dithionite were required to fully reduce the enzyme sample. C displays the spectra of the fully oxidized (–––), the radical (solid line), and the fully reduced (solid line) forms of FAD bound to mPAO that resulted from the factor analysis of the titration data presented in A and B.

 

mPAO was titrated anaerobically with a standardized solution of sodium dithionite (Fig. 5). A significant amount of the one-electron reduced flavin radical formed in the initial phase of the titration and disappeared in the final phase as it converted to the two-electron fully reduced form of bound FAD. The intermediate one electron-reduced species was predominantly the anionic (so-called "red") radical, but a trace of the neutral (so-called "blue") radical was evident also by the low absorbance in the 500–650 nm region of the spectrum (Fig. 5) (54). From this titration, the {epsilon}458 for the bound FAD was found to be 10,600 M—1 cm1, whereas the {epsilon}274 was determined to be 99,200 M—1 cm1.

mPAO was titrated anaerobically with N1-acetyl-SPD (data not shown). We chose N1-acetyl-SPD as the mPAO reductant rather than the better substrate N1-acetyl-SPM because we wanted to avoid the possible slow reduction of the high concentration enzyme by SPD, the N1-acetyl-SPM oxidation product. Because mPAO oxidizes N1-acetyl-SPM, N1-acetyl-SPD, or SPM at the exo-carbon of secondary amino groups (see below), there is no chance that the N1-acetyl-SPD oxidation product PUT (which does not have a secondary amino group) would be oxidized during the titration. Based on the A458, the concentration of enzyme was 1.20 µM, whereas a concentration of 1.16 µM was determined from the N1-acetyl-SPD titration; as expected 1 mol of substrate reduced 1 mol of FAD. Thus, all enzyme molecules in the preparation are capable of oxidizing the substrate. No trace of a flavin radical was detected during this titration. This indicates rapid transfer of 2 electrons from enzyme-bound substrate to enzyme-bound FAD.

The Steady-state Kinetic Properties of mPAO—It was assumed that steady-state mechanism for the oxidation of the various polyamine derivatives listed in Table I is of the ping-pong type. This is supported by the fact that the apparent kcat/KS values for assay done in air-saturated (0.237 mM O2) and pure O2-saturated (1.2 mM O2) buffers were approximately equal. Further support for this contention is provided by the kcat/KO values for N1-acetyl-SPM and N1-acetyl-SPD. These values are equal (Table I), as expected for a ping-pong type mechanism (42).

The steady-state kinetic studies indicated that N1-acetyl-SPM is the best substrate for mPAO, although N1-acetyl-SPD is also a good substrate (Table I). The kcat/KS value (the so-called "specificity constant") for the former substrate is over an order of magnitude higher than for the latter. Whereas SPM can be oxidized by the enzyme, it is much less efficient than for the oxidation of N1-acetyl-SPM or N1-acetyl-SPD; the kcat/KSPM value is 4 orders of magnitude lower than that for N1-acetyl-SPM. It was found that SPD, PUT, N8-acetyl-SPD, and benzylamine were not oxidized by mPAO.

Vujcic et al. (46) expressed cloned hpao and mpao genes in a human kidney cell line and determined that these cells oxidized substrates with the following preference: N1-acetyl-SPM {approx} N1-acetyl-SPD > N1,N12-diacetyl-SPM >> spermine. These findings are basically the same as those reported herein.

Interestingly, both BESPM and BENSPM are fairly good mPAO substrates, both being better than SPM (Table I). This is an important finding because BENSPM has been used for phase II cancer clinical trials (55). These N-ethylated polyamines have been used widely also to study the physiological effects of polyamine-metabolizing enzymes. They down-regulate polyamine biosynthetic enzymes, but dramatically up-regulate SSAT synthesis (13, 14), which results in mammalian cells becoming apoptotic.

It has been reported that terminally alkylated polyamine analogs like BESPM and BENSPM are oxidatively dealkylated by PAO; for BESPM and BENSPM this would result in the formation of N1-ethyl-SPM and N1-ethyl-nor-SPM, respectively, and acetaldehyde (56, 57, 58, 59). In contrast, it was reported recently that the lysates of cultured human cells transiently transfected with mpao or hpao cDNA did not dealkylate BESPM and BENSPM but converted them to N1-ethyl-SPD and N1-ethyl-nor-SPD ({3-[(3-aminopropyl)amino]propyl}-ethylamine), respectively, and N-ethyl-3-aminopropanal. It was not possible for us to resolve this dilemma by analyzing the products formed when BESPM or BENSPM were oxidized by pure mPAO, as was done for N1-acetyl-SPM and N1-acetyl-SPD (see below). The appropriate reference compounds (i.e. N1-ethyl-SPM, N1-ethyl-nor-SPM, N1-ethyl-SPD, N1-ethyl-nor-SPD, or N-ethyl-3-aminopropanal) are not available commercially and are not conveniently synthesized.

Inspection of KI values (Table I) indicates that SPM and PUT (but not benzylamine) are weak inhibitors for the oxidation of N1-acetyl-SPM (Table I), whereas N8-acetyl-SPD and SPD are somewhat better inhibitors. All of these compounds are competitive inhibitors, because in assay with each inhibitor at levels that would produce significant inhibition, the inclusion of N1-acetyl-SPM at a saturating concentration eliminated the inhibition.

Ideally, we would like to compare the biochemical and kinetic properties of the E. coli-produced mPAO with that isolated from a natural source. However, this was not feasible for several reasons. First, it is not prudent to compare the E. coli-produced mouse enzyme with the enzyme purified from bovine liver because of the species difference. Furthermore, the specific activities obtained from two different purifications of the bPAO were not the same and were lower than expected for a fully functional enzyme. Because only 1 mg of PAO could be obtained from 1 kg of bovine liver, it would be difficult to obtain sufficient quantities of PAO from mouse liver for comparative studies. We attempted to obtain a full-length bpao coding cDNA fragment. Unfortunately, we could not identify the appropriate clones from any cDNA library that was screened. Therefore, we were not able to heterologously produce bPAO for comparison with the enzyme obtained from bovine liver. Furthermore, a kcat/KS value of 2.54 x 106 M—1 s1 for oxidation of N1-acetyl-SPM by mPAO is that expected for a native, fully active enzyme (60). Additionally, recombinant mPAO is a highly stable, monomeric protein that maintains its biochemical, redox, and kinetic properties even after prolonged storage. Finally, 1 mol of enzyme FAD is reduced efficiently by 1 mol of substrate in the reductive titration (see above). Thus, we are confident that this E. coli-produced mPAO is in its native, fully active form.

The Nature of the Product Resulting from the Oxidation of N1-Acetyl-SPM and N1-Acetyl-SPD by mPAO—By using the HPLC method described under "Experimental Procedures," it was found that complete oxidation of N1-acetyl-SPM and N1-acetyl-SPD by mPAO yielded 96 and 94% (based on 1 mol of aldehyde/mol of substrate), respectively, of the 2,4-dinitrophenylhydrazone of 3-acetamidopropanal (Fig. 6). There was no trace of any other 2,4-dinitrophenylhydrazones such as would be seen if the enzyme oxidized the substrates on the endo-side of their N4-nitrogens. We expect that the phenylhydrazone of this aldehyde, because of its positive charge, would have very short HPLC retention times; the retention time of the 2,4-dinitritophenylhydrazone of 3-aminopropanal (a reference compound that is positively charged in the HPLC solutions containing trifluoroacetic acid) has a much shorter retention time than the same derivatives of 3-acetamidopropanal and acrolein, which are uncharged.

The acetyl group of 3-acetamidopropanal did not hydrolyze during the enzymatic reaction or during the workup preceding the analyses. This hydrolysis would produce 3-aminopropanal, which can spontaneously convert to acrolein (18). However, the 2,4-dinitrophenylhydrazones of either 3-aminopropanal or acrolein were not detected in the HPLC analyses of the enzyme reaction solutions (Fig. 6).

To prove definitively that 3-acetamidopropanal was the true product of these enzymatic oxidations, a larger scale reaction between N1-acetyl-SPM and mPAO was carried out, and the 2,4-dintorphenylhydrazone of the product aldehyde was isolated in 89% yield. The chemical properties of this compound and the 2,4-ditrophenylhydrazone of 3-acetamidopropanal generated by organic synthesis were compared. The two substances were identical in all respects, and 1H NMR and mass spectral analyses prove incontrovertibly that 3-acetamidopropanal is the enzymatic oxidation product.

By using an HPLC method to analyze dansylate polyamines, it was found that complete mPAO oxidation of N1-acetyl-SPM and N1-acetyl-SPD produced 95% SPD and 91% PUT (based on 1 mol/mol of substrate), respectively (data not shown). Another research group (46) exposed these substrates to lysates from HEK-293 cultured human kidney cells transiently transfected with the genes for hpao or mpao. By using an HPLC method similar to that described herein for analyzing dansylated polyamines, they found also that mPAO and hPAO converted these substrates to SPD and PUT, respectively.

These observations indicate that N1-acetyl-SPM and N1-acetyl-SPD are always oxidized at the carbon on the exo-side of their N4-nitrogens. Thus, there can be no doubt that mPAO is the classical polyamine oxidase that has been described and studied over the past few decades.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Peroxisomal PAO, an integral component of polyamine interconversion pathway, is an important player in regulating cellular polyamine levels. Thus, understanding the precise biochemical and structural properties of PAO are essential for a deeper understanding of its participation in many fundamental cellular processes. With this in mind, we set out to develop a system that would provide, in good yield, a highly purified preparation of a mammalian peroxisomal PAO. In the course of accomplishing this goal, we cloned and sequenced the entire mpao (Fig. 2) gene and most of the bpao gene (GenBankTM accession number AF226658 [GenBank] ). Based on a comparison of primary structures (Fig. 3), the sequence identity with other flavin-containing amine oxidases is less than 40%. This indicates that PAO represents a new subfamily of flavoproteins.

Inspection of the translated mPAO and bPAO sequences indicated the presence of a {beta}1{alpha}{beta}2 FAD-binding fingerprint motif (Fig. 3) that interacts with the ADP moiety of the enzyme-bound FAD (26, 33). This motif along with numerous other conserved regions are found in the "FAD-binding domain" and are elements of the Rossmann fold (Fig. 3). Two other conserved regions are located in the "substrate-binding domain" (26, 33).

Cys406, Cys397, and Cys399, the residues that are covalently attached to the 8{alpha}-carbon of the isoalloxazine ring of FAD in of MAO-B, MAO-A (24, 25, 33), and fMAO, respectively, are pointed out in the aligned sequences of Fig. 3. These Cys residues align with Ser residues in bPAO, mPAO (Ser429), and the murine and human SMO (Ser481 in both). However, except for the mitochondrial monoamine oxidases, FAD is noncovalently bound to all known amine oxidases of this family.

As with MAO-A, MAO-B (61, 62), and MAO-N (28), mPAO forms an intermediate anionic (red) radical when titrated with dithionite. This indicates that there is either a positively charged aminoacyl group (i.e. Arg or Lys) or the positive end of an {alpha}-helix dipole is near the N-1 position of the flavin's isoalloxazine ring. This positively charged environment stabilizes the negative charge of the red radical, which is localized at the N1/C2/C2O locus of the isoalloxazine ring of the flavin. In the MAO-B and cPAO structures, the positive end of an {alpha}-helix interacts with the N1/C2/C2-O locus of FAD (26, 33). The helices span residues from Met438 to Met454 of MAO-B and residues from His469 to Gln487 of cPAO (Fig. 3). These segments of cPAO and MAO-B align with the highly conserved region near the C termini (25) of the other flavoprotein amine oxidases (Fig. 3; for mPAO, Thr475–Gln496), and the secondary structure prediction program "Psi-Pred" (version 2, at the web site, insulin.brunel.ac.uk/psipred/) indicated that this region of mPAO forms an {alpha}-helix.

Whereas SPM can be oxidized by peroxisomal mPAO, it is a poor substrate when compared with N1-acetyl-SPM and N1-acetyl-SPD. SPM and SPD are acetylated for transport from the cells and eventual excretion from the body (1). High PAO levels could prevent transport of N1-acetyl-SPM and N1-acetyl-SPD from the cell and increase the SPD and PUT levels of the polyamine pool of the cell (Fig. 1). In contrast, hSMO and mSMO oxidize SPM (KS = 18 µM) (10) but not N1-acetyl-SPM or N1-acetyl-SPD (10, 11). In addition to the absence of a peroxisomal transport signal at the C termini of hSMO and MSMO, there are other significant sequence differences between these mPAO (Fig. 3); hSMO and mSMO also do not have N-terminal transport signals, suggesting that they are cytosolic enzymes.

Polyamine interconversion involving peroxisomal PAO helps maintains the intracellular balance of these substances. It has been proposed that in some cells under stress, polyamine oxidation generates the toxic byproducts H2O2 and 3-aminopropanal (generated by enzymatic deacetylation of 3-acetamidopropanal, the product of N1-acetyl-SPM oxidation by PAO; Fig. 1), which can initiate cell death (15, 16, 17, 64, 65, 66, 67, 68, 69). In addition, 3-aminopropanal can spontaneously convert to the extreme cytotoxin acrolein (18). H2O2 can be inactivated by catalase in peroxisomes, unless the levels of N1-acetyl-SPM and PAO are extremely high (or the catalase level low), which seems to be the case in some pre-apoptotic cells. It is believed also that the level of SSAT, which produces N1-acetyl-SPD and N1-acetyl-SPM from the polyamine pool, is elevated in these cells (15, 16, 17, 64, 65, 66, 67, 68). However, in some cultured cancer cells, the levels of SSAT and thus the N1-acetylated polyamines are high, but PAO is low. In fact, the level of PAO activity decreases as the histological grade of breast cancer tumors increases (63). From these observations, it can be proposed that a PAO-dependent apoptosis initiation mechanism is intact in some precancerous cells. However, events take place whereby PAO activity is interrupted, shutting down cell death, and cellular proliferation ensues.

In contrast, for tissue damaged by ischemia/reperfusion, the level of SSAT, N1-acetylate polyamines, and PAO increase (19, 20, 21). This results in the production of high levels of H2O2 and 3-aminopropanal (and perhaps acrolein), which contributes to tissue damage.

With the work described herein, a program has been initiated to study the detailed chemical, biochemical, structural, kinetic, inhibition, and mechanistic properties of a mammalian peroxisomal PAO. Hopefully, this will lead to a richer appreciation of its involvement in apoptosis, cellular proliferation, cell signaling, tissue damage, wound healing, tissue development, and differentiation, etc. and aid in the development of clinically relevant approaches to treat cancer and ameliorate ischemic tissue damage.


    FOOTNOTES
 
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AF226656 [GenBank] and AF226658 [GenBank] .

* This work was supported by the Department of Veterans Affairs and the NHLBI, National Institutes of Health. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed: Dept. of Veterans Affairs Medical Center, Molecular Biology Division (151-S), 4150 Clement St., San Francisco, CA 94121. Tel.: 415-387-1431; Fax: 415-750-6959; E-mail: wsm{at}itsa.ucsf.edu.

1 The abbreviations used are: SPM, spermine; N1-acetyl-SPD, N1-acetylspermidine; N1-acetyl-SPM, N1-acetylspermine; BENSPM, N1,N11-bis(ethyl)-nor-SPM; BESPM, N1,N12-bis(ethyl)-SPM; bPAO and bpao, bovine N1-acetylated polyamine oxidase and its gene; Cb Ac-SMO, N1-acetylspermine oxidase from C. boidinii; ESI-MS, electrospray ionization mass spectrometry; hPAO and hpao, human murine N1-acetylated PAO and its gene; hSMO, human spermine oxidase; MAO-A and MAO-B, monoamine oxidase form A and form B from humans and other mammals; MAO-N monoamine oxidase from A. niger; MDL 72527, N1,N4-bis(butadienyl)-1,4-diaminobutane; mPAO and mpao, murine N1-acetylated PAO and its gene; mSMO, murine spermine oxidase; PAO and pao, N1-acetylated polyamine oxidase and its gene; PUT, putrescine; SPD, spermidine; SMO, spermine oxidase; HPLC, high pressure liquid chromatography; GR, glutathione reductase; SSAT, SPD/SPM N1-acetyltransferase; cPAO, corn PAO; MOPS, 4-morpholinepropanesulfonic acid. Back

2 SMO, referred to as PAO in Refs. 10 and 12, oxidizes SPM but not the N1-acetylated polyamines. By definition, PAO, designated EC 1.5.3.11 [EC] , is a flavoprotein that oxidizes specifically N1-acetyl-SPM and N1-acetyl-SPD. To date, an EC number for SMO has not been assigned. To avoid further confusion concerning the identity of these two polyamine-oxidizing enzymes, the acronyms SMO and PAO taken from Ref. 11 will be used. Back



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Seiler, N. (1995) Prog. Brain Res. 106, 333–344[Medline] [Order article via Infotrieve]
  2. Wallace, H. M. (1998) Biochem. Soc. Trans. 26, 569–571[Medline] [Order article via Infotrieve]
  3. McIntire, W. S., and Hartmann, C. (1993) in Principals and Applications of Quinoproteins (Davidson, V. L., ed) pp. 97–171, Marcel Dekker, Inc., New York
  4. Morgan, D. M. (1998) Biochem. Soc. Trans. 26, 586–591[Medline] [Order article via Infotrieve]
  5. Heby, O., and Persson, L. (1990) Trends Biochem. Sci. 15, 153–158[CrossRef][Medline] [Order article via Infotrieve]
  6. Tabor, C. W., and Tabor, H. (1984) Annu. Rev. Biochem. 53, 749–790[CrossRef][Medline] [Order article via Infotrieve]
  7. Pegg, A. E. (1986) Biochem. J. 234, 249–262[Medline] [Order article via Infotrieve]
  8. Pegg, A, E., and McCann, P. P. (1988) ISI Atlas of Sci.: Biochem. 1, 11–18
  9. Nishioka, K. (1993) Cancer Res. 53, 2689–2692[Medline] [Order article via Infotrieve]
  10. Wang, Y., Devereux, W., Woster, P. M., Stewart, T. M., Hacker, A., and Casero, R. A., Jr. (2001) Cancer Res. 61, 5370–5373[Abstract/Free Full Text]
  11. Vujcic, S., Diegelman, P. Bacchi, C. J. Kramer, D. L., and Porter, C. W. (2002) Biochem. J. 367, 665–675[CrossRef][Medline] [Order article via Infotrieve]
  12. Murray-Stewart, T., Wang, Y., Devereux, W., and Casero, R. A. (2002) Biochem. J. 368, 673–677[CrossRef][Medline] [Order article via Infotrieve]
  13. Hu, R. H., and Pegg, A. E. (1997) Biochem. J. 328, 307–316[Medline] [Order article via Infotrieve]
  14. Kramer, D. L., Vujcic, S., Diegelman, P., Alderfer, J., Miller, J. T., Black, J. D., Bergeron, R. J., and Porter, C. W. (1999) Cancer Res. 59, 1278–1286[Abstract/Free Full Text]
  15. Mank-Seymour, A. R., Murray, T. R., Berkey, K. A., Xiao, L., Kern, S., and Casero, R. A. (1998) Clin. Cancer Res. 4, 2003–2008[Abstract]
  16. Lindsay, G. S., and Wallace, H. M. (1999) Biochem. J. 337, 83–87[CrossRef][Medline] [Order article via Infotrieve]
  17. Chopra, S., and Wallace, H. M. (1998) Biochem. Pharmacol. 55, 1119–1123[CrossRef][Medline] [Order article via Infotrieve]
  18. Houen, G., Bock, K., and Jensen, A. L. (1994) Acta Chem. Scand. 48, 52–60[Medline] [Order article via Infotrieve]
  19. Ivanova, S., Botchkina, G. I., Al-Abed, Y., Meistrell, M., Batliwalla, F., Dubinsky, J. M., Iadecola, C., Wang, H., Gregersen, P. K., Eaton, J. W., and Tracey, K. J. (1998) J. Exp. Med. 188, 327–340[Abstract/Free Full Text]
  20. Dogan, A., Rao, A. M., Baskaya, M. K., Hatcher, J., Temiz, C., Rao, V. L., and Dempsey, R. J. (1999) J. Neurosurg. 90, 1078–1082[Medline] [Order article via Infotrieve]
  21. Dogan, A., Rao, A. M., Hatcher, J., Rao, V. L., Baskaya, M. K., and Dempsey, R. J. (1999) J. Neurochem. 72, 765–770[CrossRef][Medline] [Order article via Infotrieve]
  22. Nishikawa, M., Hagishita, T., Yurimoto, H., Kato, N., Sakai, Y., and Hatanaka, T. (2000) FEBS Lett. 476, 150–154[CrossRef][Medline] [Order article via Infotrieve]
  23. Tavladoraki, P., Schininà, M. E., Cecconi, F., Di Agostino, S., Manera, F., Rea, G., Mariottini, P., Federico, R., and Angelini, R. (1998) FEBS Lett. 426, 62–66[CrossRef][Medline] [Order article via Infotrieve]
  24. Bach, A. W. J., Lan, N. C., Johnson, D. H., Abell, C. W., Bembenek, M. E., Kwan, S.-W., Seeburg, P. H., and Shih, J. C. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 4934–4938[Abstract]
  25. Weyler, W., Hsu, Y.-P. P., and Breakfield, X. O. (1990) Pharmacol. Ther. 47, 319–417
  26. Binda, C., Coda, A., Angelini, R., Federico, R., Ascenzi, P., and Mattevi, A. (1999) Structure 7, 265–276[CrossRef][Medline] [Order article via Infotrieve]
  27. Schilling, B., and Lerch, C. (1995) Biochim. Biophys. Acta 1243, 529–537[Medline] [Order article via Infotrieve]
  28. Sablin, S. O., Yankovskaya, V., Bernard, S., Cronin, C. N., and Singer, T. P. (1998) Eur. J. Biochem. 253, 270–279[Abstract]
  29. Ishizuka, H., Horinouchi, S., and Beppu, T. (1993) J. Gen. Microbiol. 139, 425–432[Medline] [Order article via Infotrieve]
  30. Roh, J. H., Wouters, J., Depiereux, E., Yukawa, H., Inui, M., Minami, H., Suzuki, H., and Kumagai, H. (2000) Biochem. Biophys. Res. Commun. 268, 293–297[CrossRef][Medline] [Order article via Infotrieve]
  31. Dailey, T. A., and Dailey, H. A. (1998) J. Biol. Chem. 273, 13658–13662[Abstract/Free Full Text]
  32. Dym, O., and Eisenberg, D. (2001) Protein Sci. 10, 1712–1728[Abstract/Free Full Text]
  33. Binda, C., Newton-Vinson, P., Hubalek, F., Edmondson, D. E., and Mattevi, A. (2002) Nat. Struct. Biol. 9, 22–26[CrossRef][Medline] [Order article via Infotrieve]
  34. Gasparyan, V. K. (1995) Biokhimiya 60, 1632–1636
  35. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  36. Holt, A., Sharman, D. F., Baker, G. B., and Palcic, M. M. (1997) Anal. Biochem. 244, 384–392[CrossRef][Medline] [Order article via Infotrieve]
  37. Hölttä, E. (1983) Methods Enzymol. 94, 306–311[Medline] [Order article via Infotrieve]
  38. Singer, T. P., and McIntire, W. S. (1984) Methods Enzymol. 106, 369–378[Medline] [Order article via Infotrieve]
  39. Edmondson, D. E., and Singer, T. P. (1973) J. Biol. Chem. 248, 8144–8149[Abstract/Free Full Text]
  40. Efimov, I., Cronin, C. N., and McIntire, W. S. (2001) Biochemistry 40, 2155–2166[CrossRef][Medline] [Order article via Infotrieve]
  41. Engst, S., Kuusk, V., Efimov, I., Cronin, C. N., and McIntire, W. S. (1999) Biochemistry 38, 16620–16628[CrossRef][Medline] [Order article via Infotrieve]
  42. McIntire, W., Hopper, D. J., and Singer, T. P. (1985) Biochem. J. 228, 325–335[Medline] [Order article via Infotrieve]
  43. Goudar, C. T., Sonnad, J. R., and Duggleby, R. G. (1999) Biochim. Biophys. Acta 1429, 377–383[Medline] [Order article via Infotrieve]
  44. Hunter, K. J. (1998) Methods Mol. Biol. 79, 119–123[Medline] [Order article via Infotrieve]
  45. Gasparyan, V. K., and Nalbandian, R. M. (1991) Biokhimiya 55, 1223–1227
  46. Vujcic, S., Liang, P., Diegelman, P., Kramer, D. L., and Porter, C. W. (2003) Biochem. J. 370, 19–28[CrossRef][Medline] [Order article via Infotrieve]
  47. Gould, S. J., Keller, G. A., Hosken, N., Wilkinson, J., and Subramani, S. (1989) J. Cell Biol. 108, 1657–1664[Abstract]
  48. Hall T. A. (1999) Nucleic Acids Symp. Ser. 41, 94–98
  49. Wierenga, R. K., Terpstra, P., and Hol, W. G. (1986) J. Mol. Biol. 187, 101–107[Medline] [Order article via Infotrieve]
  50. Hölttä, E. (1977) Biochemistry 16, 91–100[Medline] [Order article via Infotrieve]
  51. Tsukada, T., Furusako, S., Maekawa, S., Hibasami, H., and Nakashima, K. (1998) Int. J. Biochem. 20, 695–702[CrossRef]
  52. Pace, C. N., Vajdos, F, Fee, L., Grimsley, G., and Gray, T. (1995) Protein Sci. 4, 2411–2423[Abstract/Free Full Text]
  53. Müller, F. (1991) in Chemistry and Biochemistry of Flavoenzymes (Müller, F., ed) Vol. 1, pp. 1–71, CRC Press, Inc., Boca Raton, FL
  54. Massey, V., and Hemmerich, P. (1980) Biochem. Soc. Trans. 8, 246–257[Medline] [Order article via Infotrieve]
  55. Bergeron, R. J., Mueller, R., Bussenius, J., McManis, J. S., Merriman, R. L., Smith, R. E., Yao, H., and Weimar, W. R. (2000) J. Med. Chem. 43, 224–235[CrossRef][Medline] [Order article via Infotrieve]
  56. Bolkenius, F. N., and Seiler, N. (1989) Biol. Chem. Hoppe Seyler 370, 525–531[Medline] [Order article via Infotrieve]
  57. Bitonti, A. J., Dumont, J. A., Bush, T. L., Stemerick, D. M., Edwards, M.L., and McCann, P. P. (1990) J. Biol. Chem. 265, 382–388[Abstract/Free Full Text]
  58. Bergeron, R. J., Feng, Y., Weimar, W. R., McManis, J. S., Dimov, H., Porter, C., Raisler, B., and Phanstiel, O. (1997) J. Med. Chem. 40, 1475–1494[CrossRef][Medline] [Order article via Infotrieve]
  59. Bergeron, R. J., Merriman, R. L., Olson, S. G., Wiegand, J., Bender, J., Streiff, R. R., and Weimar, W. R. (2000) Cancer Res. 60, 4433–4439[Abstract/Free Full Text]
  60. Fersht, A. (1985) Enzymes Structure and Function, 2nd Ed., pp. 121–154, W. H. Freeman & Co., New York
  61. Sablin, S. O., and Ramsay, R. R. (1998) J. Biol. Chem. 273, 14074–14076[Abstract/Free Full Text]
  62. Edmondson, D. E. (1995) Xenobiotica 25, 735–753[Medline] [Order article via Infotrieve]
  63. Wallace, H. M., Duthie, J., Evans, D. M., Lamond, S., Nicoll, K. M., and Heys, S. D. (2000) Clin. Cancer Res. 6, 3657–3661[Abstract/Free Full Text]
  64. Ha, H. C., Woster, P. M., Yager, J. D., and Casero, R. A. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 11557–11562[Abstract/Free Full Text]
  65. Ferioli, M. E., Pinotti, O., and Pirona, L. (1999) Biochem. Pharmacol. 58, 1907–1914[CrossRef][Medline] [Order article via Infotrieve]
  66. Rao, A. M., Hatcher, J. F., Dogan, A., and Dempsey, R. J. (2000) J. Neurochem. 74, 1106–1111[Medline] [Order article via Infotrieve]
  67. Hatcher, J., Rao, A. M., Dogan, A., and Dempsey, R. J. (2000) Soc. Neurosci. Abstr. 26, 769.9
  68. Zoli, M., Pedrazzi, P., Zini, I., and Agnati, L. F. (1996) Brain Res. Mol. Brain Res. 38, 122–134[Medline] [Order article via Infotrieve]
  69. Ivanova, S., Batliwalla, F., Mocco, J., Kiss, S., Huang, J., Mack, W., Coon, A., Eaton, J. W., Al-Abed, Y., Gregersen, P. K., Shohami, E., Connolly, E. S., Jr., and Tracey, K. J. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 5579–5584[Abstract/Free Full Text]