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(The FASEB Journal. 2006;20:1097-1108.)
© 2006 FASEB

Divergent cyclooxygenase responses to fatty acid structure and peroxide level in fish and mammalian prostaglandin H synthases

Wen Liu*, Dazhe Cao*,1, Sungwhan F. Oh{dagger}, Charles N. Serhan{dagger} and Richard J. Kulmacz*,2

* Department of Internal Medicine, University of Texas Health Science Center at Houston, Houston, Texas, USA; and

{dagger} Center for Experimental Therapeutics and Reperfusion Injury, Department of Anesthesiology, Perioperative and Pain Medicine, Brigham and Women’s Hospital and Harvard Medical School, Boston, Massachusetts, USA

1Correspondence: Department of Internal Medicine, University of Texas Health Science Center at Houston, 6431 Fannin St., Houston, TX 77030, USA. E-mail: richard.j.kulmacz{at}uth.tmc.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Prostanoid synthesis in mammalian tissues is regulated at the level of prostaglandin H synthase (PGHS) cyclooxygenase catalysis by the availability and structure of substrate fatty acid and the availability of peroxide activator. Two major PGHS isoforms, with distinct pathophysiological functions and catalytic regulation, have been characterized in mammals; a functionally homologous PGHS isoform pair has been cloned from an evolutionarily distant vertebrate, brook trout. The cyclooxygenase activities of recombinant brook trout PGHS-1 and -2 were characterized to test the generality of mammalian regulatory paradigms for substrate specificity, peroxide activation, and product shifting by aspirin. Both trout cyclooxygenases had much more restrictive substrate specificities than their mammalian counterparts, with pronounced discrimination toward arachidonate (20:4n-6) and against eicosapentaenoate (20:5n-3) and docosahexaenoate (22:6n-3), the latter two prominent in trout tissue lipids. Aspirin treatment did not increase lipoxygenase-type catalysis by either trout enzyme. Both trout enzymes had higher requirements for peroxide activator than their mammalian counterparts, though the preferential peroxide activation of PGHS-2 over PGHS-1 seen in mammals was conserved in the fish enzymes. The divergence in cyclooxygenase characteristics between the trout and mammalian PGHS proteins may reflect accomodations to differences among vertebrates in tissue lipid composition and general redox state.—Liu, W., Cao, D., Oh, S. F., Serhan, C. N., Kulmacz, R. J. Divergent cyclooxygenase responses to fatty acid structure and peroxide level in fish and mammalian prostaglandin H synthases.


Key Words: cyclooxygenase substrate specificity • cyclooxygenase activation by peroxide • aspirin-induced lipoxygenase catalysis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
CYCLOOXYGENASE CATALYSIS BY prostaglandin H synthase (PGHS) is a major regulatory point in the biosynthesis of prostanoid signaling molecules (1) . A pair of major PGHS isoforms, arising from genes on separate chromosomes, is found in many vertebrates, ranging from humans to fish; invertebrates appear to have evolved a pair of PGHS isoforms distinct from that in vertebrates (2) . Studies in mammals have established that while PGHS-1 and -2 share extensive structural and enzymatic similarities, they have very different pathophysiological functions and distinct regulatory controls at both the level of gene expression and of catalysis (3) . This differential catalytic control has been dramatically demonstrated in several mammalian cells by observing vigorous catalysis by PGHS-2 cyclooxygenase, whereas PGHS-1 cyclooxygenase remains latent (4) .

One aspect of differential catalytic regulation in the two mammalian isoforms is the more efficient cyclooxygenase activation in PGHS-2 than in PGHS-1, with half-maximal activation at 2 and 20 nM PGG2, respectively (5) . This difference would allow low cellular peroxide levels to support cyclooxygenase catalysis by PGHS-2 but not by PGHS-1 (6) . However, the evidence indicating that cellular peroxide levels are a key factor in differential control of cyclooxygenase catalysis currently is mostly limited to mammalian enzymes (5 , 7 8 9 10) , and tests of the generality of this regulatory scheme are needed. A second aspect to differential cyclooxygenase catalytic regulation in the mammalian isoforms is their selectivity toward fatty acid substrates. Mammalian PGHS-2 cyclooxygenase utilizes a wider variety of fatty acids than does the PGHS-1 cyclooxygenase (11 12 13) . Mammalian PGHS-2 can also oxygenate fatty acid esters and amides that are not substrates for mammalian PGHS-1 (14 , 15) . The two mammalian PGHS isoforms also differ dramatically in the consequences of pretreatment with aspirin, with acetylation of Ser-530 blocking all oxygenation activity in PGHS-1 but shifting PGHS-2 from cyclooxygenase to lipoxygenase (LOX) catalysis (16 17 18) . This shift in PGHS-2 product may contribute to aspirin’s various pharmacological actions (19) .

Polyunsaturated fatty acids (PUFA) of the n-3 family are increasingly recognized as a beneficial dietary component in humans (20 , 21) . The n-3 PUFA have long been known to be essential dietary components in many fish species (22) , and n-3 fatty acids predominate over n-6 fatty acids in tissue lipids of both freshwater and marine fish (23) . Despite this abundance of n-3 PUFA, prostanoid synthesis by fish tissues, whether from endogenous or from exogenous precursors, generally prefers 20:4n-6 over 20:5n-3 and discriminates strongly against 22:6n-3 (24) . It has remained a puzzle whether this discrimination against the abundant n-3 PUFA is a property of the fish PGHS proteins themselves or is dictated by other cellular factors.

PGHS isoform pairs have been cloned from several species of bony fish (25 26 27 28) . There are several indications that the fish PGHS isoform pair is functionally homologous to the mammalian pair. PGHS-1 is the predominant platelet isoform in both zebrafish and humans (27 , 29) , and the enzyme appears to have comparable hemostatic roles in fish and mammals (30) . Just as in mammals, tissue expression of PGHS-2 is more restricted than that of PGHS-1 in both trout and zebrafish (4 , 26 , 27) . As in mammals, PGHS-2 is inducible by mitogens and cytokines in fish (4 , 25 , 26 , 28 , 31 , 32) . Despite the functional homology, examination of the amino acid sequences indicates there is considerable evolutionary divergence between mammalian and fish PGHS proteins. The amino acid sequence identity is only ~70% between human and trout PGHS-1 and between human and trout PGHS-2, whereas it is nearly 90% among mammalian PGHS-1s and among mammalian PGHS-2s. We exploited the availability of the PGHS isoform pair from this evolutionarily distant vertebrate species to test the generality of the peroxide activation, substrate specificity, and aspirin product shift patterns established in mammalian PGHS isoforms.

There has been considerable attention recently to the role of cellular lipid composition and redox state in modulating the pathophysiology of prostanoid signaling and in the therapeutic/preventive potential of dietary intervention (33 , 34) . The present results provide some fresh and unexpected biochemical insights into how the structure of available fatty acid substrates and the redox state can affect the formation of potent lipid mediators that are relevant to human systems.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Fatty acids were purchased from NuChek Preps (Elysian, MN, USA). Glutathione (GSH) peroxidase-1 (GPx-1), GSH reductase, GSH, hydrogen peroxide, diethyldithiocarbamate, 1-methyl-3-nitro-1-nitrosoguanidine, and heme were from Sigma Aldrich (St. Louis, MO, USA). Tween 20 was from Anatrace (Maumee, OH, USA). [14C]-Labeled fatty acids were from American Radiolabeled Chemicals, Inc. (St. Louis, MO, USA). LK6 silica TLC plates were from Whatman (Clifton, NJ, USA). Eicosanoid standards were from Cayman Chemical Co. (Ann Arbor, MI, USA). Reagents for DNA extraction and purification were from Qiagen (Valencia, CA, USA). Restriction enzymes and DNA-modifying enzymes were from New England Biolabs (Beverly, MA, USA). Oligonucleotides were from Sigma-Genosys (Houston, TX, USA). pVL1392 vector and BaculoGold linearized baculovirus DNA were from Pharmingen (San Diego, CA, USA). Sf9 (Spodoptera frugiperda) insect cells, medium, and culture reagents were from Invitrogen (Carlsbad, CA, USA). DNA sequencing was performed at the Microbiology and Molecular Genetics Core Facility, University of Texas Health Science Center at Houston.

Ovine PGHS-1 (oPGHS-1) was purified from ram seminal vesicles (35) . The well-studied oPGHS-1 has major practical advantages as a representative mammalian PGHS-1, and its reported interactions with fatty acid and inhibitors are similar to those of hPGHS-1. Recombinant human PGHS-2 (hPGHS-2) was expressed in Sf9 cells and purified as described previously (5) . Plasmids containing cDNAs coding for brook trout (Salvelinus fontinalis) PGHS-1 (tPGHS-1) and -2 were generous gifts from Dr. Frederick Goetz at the Woods Hole Marine Biological Laboratory. The 1.8-kb fragment containing brook trout PGHS-1 cDNA was generated by polymerase chain reaction (PCR) using the following primer pair: sense: 5'-GAAGATCTGCCACCATGAGAGGTTTGGTCGTGTGCGTC-3'; antisense: 5'-GCTCTAGATTACAGTTCAGTGGACTGTTTCCTTGG-3'.

The 1.8-kb fragment containing brook trout PGHS-2 cDNA was generated by PCR using the following primer pair: sense: 5'-GAAGATCTGCCACGATGAATAAAGTAGTCTGTATAATT-3'; Antisense: 5'-GCTCTAGATTAAAGCTCTAAAGTCCTTTCTTTGAAC-3'. These primer pairs incorporated an upstream Kozak sequence (GCCACC/G) (36) and removed the 3'-untranslated region. The resulting DNA fragments were digested with BglII and XbaI, ligated separately into pVL1392 vector digested with the same enzymes and used to transform E. coli DH5{alpha} competent cells. The corresponding transfer vector plasmids were isolated from ampicillin-resistant colonies; plasmid integrity was verified by restriction digests and DNA sequencing. Generation and amplification of recombinant baculovirus containing the tPGHS1 or tPGHS2 cDNA and expression of recombinant trout PGHS-1 and -2 proteins followed earlier procedures (5) . The recombinant trout proteins were purified by a modification of the method of Gierse et al. (37) . Briefly, Sf9 cells expressing the recombinant protein were suspended in 25 mM Tris, pH 8.1 containing 0.25 M sucrose, 1 mM EDTA, and 1 mM diethyldithiocarbamate and homogenized by sonication. The membrane fraction was collected by centrifugation for 1 h at 100,000 g and the recombinant protein solubilized by extraction with 1.5% Tween 20. After removal of residual membrane material by pelleting at 135,000 g for 1 h, the extract was loaded on a column of High Q ion exchange resin (Bio-Rad, Hercules, CA, USA) equilibrated with 25 mM Tris, pH 8.1, containing 0.1 mM EDTA, 0.2 mM phenol, and 0.1% Tween 20. The column was eluted with a 0–400 mM NaCl gradient. Active fractions were pooled, concentrated by ultrafiltration on a YM30 membrane (Millipore, Bedford, MA, USA), mixed with 0.25 vol of glycerol, and stored at –70°C. This procedure removes the bulk of contaminating insect cell lipids and gives an electrophoretic purity of ~50% for PGHS-2 and ~10% for PGHS-1. Typical cyclooxygenase specific activities were 2.3 and 15 K units/mg protein for partially purified tPGHS-1 and -2, respectively. No oxygenase activity was observed from the tPGHS-1 and -2 preparations in the absence of fatty acid, confirming removal of all substrate fatty acids derived from the host cells. The Sf9 host cell membranes lack endogenous arachidonate oxygenase activity (5) . There was also a large dilution factor (at least 150-fold) when enzyme was assayed, assuring that the concentration of any extraneous fatty acid binding proteins remained small compared to the levels of fatty acid added.

Cyclooxygenase activity was measured in 3 ml of 0.1 M potassium phosphate, pH 7.2, containing 1 mM phenol, 0.025% Tween 20, and 1 µM heme (35) . One unit of cyclooxygenase activity is 1 nmol O2 consumed/min. Initial assays at 30°C used a "standard" electrode membrane and 100 µM arachidonate. Assays to evaluate substrate dependence used 9–13 distinct levels of fatty acid between 2 and 150 µM at 25°C with a "high sensitivity" membrane to minimize electrode dampening effects (38) . This temperature is roughly midway between typical body temperatures for freshwater fish and mammals. The reaction mixtures contained Tween 20 to mask variations in detergency due to changes in fatty acid concentration, and heme to counter any heme dissociation driven by fatty acid (39) . To ensure that the Km and Vmax values reflected the characteristics of fully active cyclooxygenase, concentration dependence measurements were done in the presence of 100 µM H2O2. Estimates of the stimulatory effect of added hydroperoxide were obtained by comparing the reaction rates for each fatty acid at 100 µM in the presence and absence of 100 µM H2O2. All velocities were normalized to the average velocity observed in multiple calibrating reactions with 100 µM arachidonate, run with the same batch of enzyme on the same day; this permitted reliable comparison of velocities obtained with different fatty acids. Assays to evaluate the temperature dependence of cyclooxygenase activity used the "high sensitivity" membrane and included 100 µM H2O2. Protein levels were kept well below those known to alter fatty acid Km values (40) . Estimates of Km and Vmax and the corresponding SE’s were obtained by fitting reaction rates to the Michaelis-Menten equation using Kaleidagraph (Synergy Software, Reading, PA, USA). The efficiency of cyclooxygenase activation by peroxide was evaluated in reactions containing varying amounts of GPx-1 in 3 ml of potassium phosphate, pH 7.2, containing 1 mM phenol, 0.5 mM GSH, 1 µM heme, 100 µM arachidonate (5) . Reactions were started with 50–70 U of cyclooxygenase. The inhibitory effects of GPx-1 are similar in partially purified and homogeneous PGHS preparations (5 ,41) .

LOX activity in aspirin-treated hPGHS-2 and tPGHS-2 was measured from the increase in absorbance at 235 nM due to the conjugated diene chromophore in the product hydroperoxide (42) . Stock solutions of enzyme (0.55 mg protein/ml) were preincubated with a small vol of 1.0 M aspirin in dimethyl sulfoxide (final concentration 5–10 mM aspirin and 0.5–1.0% dimethylsulfoxide) and 5–20 µl aliquots transferred for assay at room temperature (22–23°C) in stirred spectrophotometer cuvettes containing 2.0 ml of 0.1 M Tris, pH 8, 10% glycerol, 0.10% Tween 20, 0.05% cholate, 0.50 mM L-tryptophan, and 60 µM arachidonate or 20:2n-6. Product formation was calculated from the {Delta}A235 after 60 s and the values were normalized to that for the control without aspirin.

Product profiles were examined using reactions containing 0.30 ml of 25 mM potassium phosphate, pH 7.2/1.0 mM phenol/0.1 mM H2O2/1 µM heme at room temperature. The enzymes were preincubated with or without aspirin before addition of 50–75 µM [14C]-labeled 20:4n-6, 20:5n-3, 22:4n-6 or 22:6n-3. Enzyme amounts were adjusted, based on oxygen measurements, to consume comparable amounts of each fatty acid (Table 1 ). Conditions used for preincubation with aspirin (Table 2 ) were based on measurements of the oxygenase inhibition kinetics for each enzyme with arachidonate as substrate (data not shown) and adjusted to give about a 50% decrease in activity. This is the theoretical decrease in oxygenase activity when aspirin converts hPGHS-2 from cyclooxygenase to 15-lipoxygenase catalysis and provides a consistent benchmark for aspirin treatment of the other enzymes and avoids potential problems from nonspecific acetylation. Reactions were stopped after 3 min by addition of 0.60 ml ice-cold ethyl acetate-toluene (1:1), 7 µl of 1.2 M HCl, and 200 mg of NaCl. After vortexing and centrifugation, the upper organic layer was dried with anhydrous Na2SO4. The aqueous layer was subjected to a second extraction and the combined extracts were stored at –20°C. For TLC analysis, unlabeled standards (arachidonate, ricinoleic acid and PGF2{alpha}) were added to a 0.3 ml aliquot of each extract and the mixture was treated with 3 mM triphenylphosphine for 30–60 min to reduce peroxides and convert PGG and PGH to PGF. The volume was then decreased to 10–20 µl by vacuum evaporation, 10 µl of acetone was added, and the solution was spotted on a TLC plate. The plate was developed in diethyl ether-heptane-acetic acid (85:15:0.1). Standards were visualized with iodine vapor before the plates were exposed to PhosphorImager screens for 20 h; the distribution of radioactivity was analyzed with a Bio-Rad Personal FX imager. Thin layer chromatography (TLC) was used for this general reaction product survey because this technique is relatively insensitive to differences in double bond number and chain length, simplifying identification of analogous products from different fatty acids.


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  		Table 1. Relative amounts of enzyme added to reactions with individual radiolabeled fatty acidsa


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  		Table 2. Preincubation conditions used for screening of product profilesa

For gas chromatography/mass spectrometry (GC-MS) analyses of arachidonate metabolites, trout PGHS-1 and -2 were incubated with unlabeled fatty acid as described above, but without triphenylphosphine treatment. Extracts were dried under a stream of nitrogen gas, dissolved in 10 µl of methanol, and esterified with diazomethane in diethyl ether for 45 min at room temperature. The methyl esters were silylated by overnight treatment with BSTFA reagent (Pierce, Rockford, IL, USA) at room temperature; this procedure is adapted from that described by Melnikova and Pivnitsky (43) . GC-MS analysis of the derivatives used a Hewlett-Packard 5971A mass selective detector quadrupole equipped with a HP Model G1030A workstation and a Model 5890 GC. The column was 30 m x 0.25 mm ID x 0.25 µm film SE-30 with 100% dimethylpolysiloxane (Supelco, Bellefonte, PA, USA). The temperature was held at 150°C for 2 min, increased at 10°C/min to 230°C, then increased at 5°C/min to 280°C. The injector was at 150°C and the transfer line at 300°C. Ionization was by electron impact at 70 eV. Saturated fatty acid methyl esters (C14-C24) were used as chromatographic standards, giving the following retention times in minutes (mean of three injections): C14, 6.50; C16, 8.51; C18, 10.37; C20, 12.35; C22, 14.58; C24, 16.95.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Effects of reducing cosubstrate
Peroxidase reducing cosubstrates, such as phenol, increase the number of cyclooxygenase catalytic turnovers before self-inactivation for mammalian PGHS-1, thereby increasing the apparent cyclooxygenase reaction rate measured with an oxygen electrode (38) . Similar isoform-specific patterns were observed for the fish and mammalian enzymes, with the PGHS-1 isoforms showing a more pronounced initial stimulation at low phenol levels and a steeper later inhibition at higher phenol levels than the PGHS-2 isoforms; the inhibitory phase was most dramatic for fish PGHS-1, which was completely inhibited at 10 mM phenol (data not shown). Stimulatory action was maximal for all enzymes near 1 mM phenol, so that concentration was used in subsequent assays.

Effect of temperature
Brook trout thrive in water near 15°C (44) , an environment much cooler than mammalian body temperatures, raising the possibility that conventional assay temperatures might be detrimental to trout PGHS activity. Accordingly, the trout and human PGHS-2 cyclooxygenase activities were assayed at 5–30°C (Fig. 1 ). Activity increased throughout the temperature range for both enzymes, indicating that their optimal temperatures are at least 30°C, the highest temperature used in the present study. The slope of the curve was steeper for hPGHS-2 than tPGHS-2, suggesting that the trout cyclooxygenase has a lower activation energy than the mammalian enzyme.


Figure 1
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  		Figure 1. Temperature dependence of hPGHS-2 (circles) and tPGHS-2 (triangles) cyclooxygenase activities. Three different amounts of hPGHS-2 and tPGHS-2 were assayed at each of the temperatures indicated; activities are normalized to the average value for each enzyme at 30°C.

Effects of fatty acid structure
Cyclooxygenase kinetics were evaluated for a panel of C18-C22 fatty acids. The Km values were narrowly clustered below 10 µM for the mammalian isoforms, with a range of 1–5 µM for hPGHS-2 and 3–9 µM for oPGHS-1 (Table 3 and Table 4 ). In contrast, the Km values tended to be higher and varied more widely for the two fish isoforms, with a range of 2–36 µM for tPGHS-2 and 13–160 µM for tPGHS-1. The lowest Km value was found with arachidonate except for hPGHS-2, where 20:5n-3 and 22:6n-3 had lower Km values. For a given fatty acid, the PGHS-2 isoform tended to have a lower Km value than the PGHS-1 isoform in both fish and mammalian enzymes.


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  		Table 3. PGHS-1 cyclooxygenase kinetic parameters


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  		Table 4. PGHS-2 cyclooxygenase kinetic parameters

The fatty acid selectivity (VRel/KRel) values (Fig. 2 ) indicate that arachidonate was the preferred substrate for all enzymes except hPGHS-2, for which four fatty acids (20:5n-3, 20:3n-6, 22:4n-6, and 20:4n-6) behaved similarly. The preference for arachidonate was very pronounced in tPGHS-1 and -2, for which only one fatty acid, 20:3n-6, had a selectivity value within an order of magnitude of that for arachidonate. This is in marked contrast to the situation with the mammalian enzymes, for which almost all the fatty acids had selectivity values within an order of magnitude of that observed with arachidonate. The trout cyclooxygenases clearly have dramatically more restrictive substrate specificities than their mammalian counterparts.


Figure 2
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  		Figure 2. Comparison of cyclooxygenase fatty acid specificity factors (Vrel/Krel) for ovine and trout PGHS-1 (oPGHS-1 and tPGHS-1) (A) and human and trout PGHS-2 (hPGHS-2 and tPGHS-2) (B). For each fatty acid, the Vrel value (Table 3 or 4) was divided by the corresponding Krel value (the Km value from Table 3 or 4 divided by that for arachidonate for the same enzyme); the SE of the Vrel/Krel ratio is indicated. Note the logarithmic scale for the Vrel/Krel axis.

The observed cyclooxygenase velocity of all four enzymes was little stimulated by added hydrogen peroxide (HOOH) when arachidonate was the substrate (Tables 3 , 4) . With hPGHS-2, a large stimulation by added HOOH was observed only with 22:6n-3. With oPGHS-1 and tPGHS-1, greater than 2-fold stimulation by HOOH was seen with five of the fatty acids tested. The greatest stimulation of oPGHS-1 by added HOOH was seen with 22:6n-3. Thus, 22:6n-3 is essentially a nonsubstrate, tight binding inhibitor of both mammalian isoforms unless exogenous peroxide is available. With tPGHS-2, 2-fold or greater stimulation was seen with only four fatty acids, and the degree of stimulation was considerably less than for the same fatty acids with tPGHS-1.

Product profiles of fish and mammalian PGHS-1 and -2
Qualitatively similar product profiles were observed for all four enzymes with 20:4n-6, 20:5n-3, and 22:4n-6, with a mixture of prostaglandins and hydroxy fatty acids evident (Fig. 3 ). The other fatty acid examined, 22:6n-3, was metabolized by the mammalian enzymes mainly to hydroxy fatty acids, but was not appreciably metabolized by either of the two fish enzymes. Pretreatment with aspirin increased 15-hydroxyeicosatetraenoic acid synthesis from arachidonate by hPGHS-2, but not from oPGHS-1 (Fig. 3) , as expected from earlier observations with human and ovine PGHS-2 (16 17 18) . A similar increase in hydroxy fatty acid synthesis was seen in aspirin-treated hPGHS-2 reacted with 22:4n-6, but not when 20:5n-3 or 22:6n-3 was the substrate for hPGHS-2. The TLC results indicated that aspirin pretreatment of tPGHS-2, tPGHS-1, and oPGHS-1 did not increase hydroxy fatty acid production with any of the fatty acids tested. Thus, the product shift from prostanoid to hydroxy fatty acid after acetylation of Ser-530 by aspirin appears to be limited to hPGHS-2 reacting with particular fatty acids.


Figure 3
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  		Figure 3. TLC analyses of products from reactions of native and aspirin-treated oPGHS-1, hPGHS-2, tPGHS-1, tPGHS-2 with 20:4n-6, 20:5n-3, 22:4n-6, and 22:6n-3. Enzymes were preincubated with or without aspirin (see Tables 1 , 2 ) before addition of radiolabeled fatty acid. The origin is at the bottom of the portion of the plate shown; the solvent front is at the top. The zones corresponding to unreacted fatty acid (FA), hydroxy fatty acids (FA-OH), and prostaglandins (PGs), indicated by brackets, were identified with standards (arachidonate, 15-hydroxyeicosatetraenoic acid, ricinoleic acid, PGD2, PGE2, and PGF2{alpha}). Radiolabeled 20:5n-3 contained a few percent of a nonmetabolized impurity with higher Rf.

The effects of aspirin treatment on LOX catalysis by hPGHS-2 and tPGHS-2 were also examined in a spectrophotometric assay (Fig. 4 ). Preincubation of hPGHS-2 with 10 mM aspirin led to a progressive loss of LOX activity with 20:2n-6 as substrate (k=0.14±0.01 min–1) and a simultaneous gain in LOX activity with arachidonate as substrate (k=0.16±0.03 min–1) (Fig. 4A ). The latter increase reflects the formation of 11- and 15-HPETE in acetylated mammalian PGHS-2s (16 17 18 ,42) . The decreased activity against 20:2n-6 indicates that this fatty acid cannot bind in a catalytically productive conformation when Ser-530 of hPGHS-2 is acetylated. Preincubation of tPGHS-2 with 5 mM aspirin led to a simultaneous loss of LOX activity with both arachidonate (k=0.064±0.005 min–1) and 20:2n-6 (k=0.059±0.004 min–1) as substrate (Fig. 4B ). Aspirin treatment thus increases LOX activity in hPGHS-2 but not in tPGHS-2, showing substrate selectivity in hPGHS-2, corroborating the results in Fig. 3 .


Figure 4
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  		Figure 4. Effects of aspirin treatment on LOX activity of hPGHS-2 (A) and tPGHS-2 (B). Formation of LOX products was measured from the increase in A235 in reactions with arachidonate (filled triangles) or 20:2n-6 (open circles) with the indicated lengths of preincubation with aspirin (10 mM for hPGHS-2, 5 mM for tPGHS-2) in 0.1 M potassium phosphate, pH 7.2, at 22–23°C. Amounts of product formation after 60 s reaction were normalized to the control value without aspirin. Data shown are pooled from two separate experiments. Details are described in Materials and Methods.

GC-MS analysis of arachidonate metabolism by trout PGHS-1 and -2
The arachidonate metabolites produced by the two trout enzymes were derivatized and subjected to GC-MS analysis. In both cases, the chromatograms monitored at m/z 173, an ion characteristic of arachidonate derivatives (45) , had multiple peaks (data not shown). SIM searches at m/z 582 (molecular ion of the PGD2 and PGE2 derivatives) and at m/z 584 (molecular ion of the PGF22a derivative) revealed four peaks for m/z 582 but no significant peaks for m/z 584; derivatized arachidonate products from trout PGHS-1 and -2 showed similar SIM chromatograms and fragment patterns. Ions of diagnostic value for the molecular ion at m/z 582 were identified in the trout PGHS-1 and -2 products as isomeric derivatives of PGD2 and PGE2 by fragment pattern and retention time matching with the authentic PGD2 and PGE2 derivatives (43) . The strongest signal for both m/z 173 and 582 was identified as a single peak with a retention time of 17.2 min, which was identified as a PGD2 derivative (Fig. 5 ).


Figure 5
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  		Figure 5. Mass spectrometric analysis of arachidonate (AA) metabolites produced by tPGHS-1. A) Endoperoxide-derived region of m/z 173 SIM chromatogram for derivatized arachidonate metabolites produced by tPGHS-1. B) Mass spectrum of PGD2 derivative eluting at 17.2 min in panel A.

Cyclooxygenase regulation by peroxide in trout PGHS-1 and -2
Titrations with GPx-1 were used to evaluate the cyclooxygenase activation efficiencies of the two PGHS isoforms from trout. (Fig. 6 ). In this experiment, the cyclooxygenase (Cox) activation efficiency is proportional to the GPx/Cox ratio needed to suppress cyclooxygenase activity (5) . Two separate batches of each trout isoform were tested, with very similar results. Increasing the amount of added GPx-1 led to a progressive suppression of the tPGHS-1 cyclooxygenase velocity, with half of the cyclooxygenase remaining latent when the GPx/Cox ratio was ~10. With tPGHS-2, little decrease in cyclooxygenase rate was seen until the GPx/Cox ratio was near 40; additional GPx produced marked suppression of the cyclooxygenase, with half of the activity remaining at a GPx/Cox ratio of ~80. Extrapolation of the data to the x axis indicates the GPx/Cox ratio needed for complete suppression of the cyclooxygenase activity (the titration "end point"); tPGHS-1 had a 6-fold lower end point than did tPGHS-2. GPx titration end points can be used to estimate the concentration of lipid peroxide (PGG2) required for half-maximal cyclooxygenase activation, termed the Kp value (5 , 46) . Using this process, the titration end points in Fig. 6 correspond to Kp values of 63 and 11 nM PGG2 for tPGHS-1 and -2, respectively. This difference between the activation efficiency of the two trout isoforms is comparable to that between the two mammalian isoforms (5) , indicating that the trout isoforms are amenable to the same sort of differential regulation by cellular peroxide scavengers proposed for the mammalian pair (5 , 6) . Data from titration of oPGHS-1 and hPGHS-2 cyclooxygenases (Fig. 5) indicated Kp values of 20 nM and 2 nM PGG2, respectively, as found previously (5) . Thus, the peroxide activation efficiency of each trout cyclooxygenase was considerably less than that of its mammalian counterpart.


Figure 6
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  		Figure 6. Suppression of trout PGHS-1 and -2 cyclooxygenase activity by GPx-1. The cyclooxygenase velocity produced by a fixed amount of each enzyme was measured in the presence of increasing amounts of GPx-1; values were normalized to the corresponding control reactions. Two separate batches of partially purified, recombinant enzyme were examined for each trout isoform; oPGHS-1 and hPGHS-2 (inset) also were examined. The control cyclooxygenase velocities were 64 and 71 U for the two batches of tPGHS-1, 50 and 51 U for the two batches of tPGHS-2.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Cyclooxygenase product profiles
The two human PGHS isoforms are known to produce comparable product profiles when reacted with arachidonate (18) , and roughly comparable product profiles were generated when arachidonate was reacted with oPGHS-1, hPGHS-2, and the two trout isoforms (Fig. 3) . This general similarity in product profile for mammalian and trout enzymes extended to three other fatty acids, 20:5n-3, 22:4n-6, and 22:6n-3 (Fig. 3) . Mass spectrometric analyses confirmed that PGD2 and PGE2 were the predominant arachidonate metabolites in both recombinant trout enzymes (shown for tPGHS-1 in Fig. 5 ). Earlier mass spectrometric analyses found PGE2 to be the predominant arachidonate metabolite in cos-7 cells expressing recombinant zebrafish PGHS-1 or -2 (27) . In addition, prostaglandins of the 2- and 3-series have been identified as major eicosanoids produced by several tissues from a variety of fish species in vivo and in vitro (47 48 49 50 51) . All of this indicates that the fish and mammalian enzymes have similar cyclooxygenase catalytic chemistry.

Substrate selectivity
Mammalian PGHS-1 has a more restrictive fatty acid substrate selectivity, favoring 20:3n-6 and 20:4n-6, than does PGHS-2 (11 12 13) . The results in Fig. 2 confirm this pattern for the mammalian pair of isoforms and show that it also holds true in the trout PGHS pair. The functional advantage of the wider substrate range in PGHS-2 is not known. Mammalian PGHS-2 has been found to generate a variety of carboxyl-substituted signaling molecules that are not produced by PGHS-1 (15) , and fish may have similar signaling pathways.

The reaction mixtures used to assess cyclooxygenase kinetic parameters were supplemented with peroxide to avoid the confounding effects of variations in activation efficiency among the fatty acids, such as that observed for oPGHS-1 with 20:4n-6 and 20:5n-3 (52) . Peroxide supplementation was especially important for the two PGHS-1 isoforms tested, as the cyclooxygenase velocity was increased many-fold by added peroxide for most of the substrates tested with these enzymes (Table 3) . Peroxide-dependent increases in cyclooxygenase activity were smaller for the two PGHS-2 isoforms, except with 22:6n-3 (Table 4) . The other 22-carbon substrate tested with hPGHS-2, 22:4n-6, was essentially equivalent to arachidonate in V/K value and stimulation by peroxide (Fig. 2 and Table 4 ). This similarity between arachidonate and 22:4n-6 suggests that mammalian tissues with elevated PGHS-2 levels can produce significant amounts of 22-carbon prostanoids, an expectation borne out for renal medulla (53 , 54) . The observation of considerable stimulation of PGHS-1 cyclooxygenase activity by added peroxide (Table 3) suggests that in cellular contexts where peroxide levels are suppressed, fatty acids other than 20:4n-6 and 20:3n-6 are likely to act more as competitive inhibitors than as substrates toward the PGHS-1 isoforms. Addition of peroxide to the cyclooxygenase assay with 20:5n-3 as substrate was reported to give only a 1.6-fold stimulation of hPGHS-1 and to inhibit hPGHS-2 (12) . However, those studies used microsomal enzyme preparations that are likely to already be activated by endogenous peroxidized lipid.

There were marked substrate selectivity differences between trout and mammalian PGHS-1 and between trout and mammalian PGHS-2 (Fig. 2) . Each fish isozyme has a considerably narrower substrate selectivity than its mammalian counterpart, with 5- to 26-fold (PGHS-1) and 3- to 77-fold (PGHS-2) higher V/K ratios for the mammalian enzymes with 20- and 22-carbon substrates other than arachidonate. This discrimination against substrates other than arachidonate is particularly significant in the case of 20:5n-3 and 22:6n-3, which are generally much more prominent components in fish lipids than in mammalian lipids (23) . Put another way, arachidonate is favored more heavily as a cyclooxygenase substrate in the vertebrate where its relative availability is lower, suggesting that prostanoids derived from arachidonate have some functional advantages over prostanoids derived from 20:5n-3 or 22:6n-3. In this connection, PGE2 has been found to be a more potent agonist than PGE3 in several fish systems (55 56 57) ; among mammalian prostanoid receptors, those for PGF and TXA have higher affinities for the cognate 2-series prostanoid than the corresponding 3-series prostanoid (33) . The weaker bias toward arachidonate in mammalian cyclooxygenases may be a consideration in human dietary intervention regimens if arachidonate and its n-6 precursors become minor components in tissue lipids.

Several studies with tissue homogenates, isolated cell preparations, and primary cell cultures derived from a variety of fish species have examined the relative production of PGE2 and PGE3, reflecting the relative cyclooxygenase utilization of endogenous arachidonate and eicosapentaenoate (48 49 50 51 , 58 59 60 61) . Indications of selective use of arachidonate (i.e., a PGE2/PGE3 ratio greater than the 20:4n-6/20:5n-3 ratio in total lipids or major phospholipids) are evident in some conditions, but not in others. Of particular interest are the results obtained with cultured turbot brain astrocytes, where preferential prostaglandin synthesis from arachidonate appeared to occur on stimulation with calcium ionophore, whereas other agonists led to preferential use of eicosapentaenoate (49) . There are also indications that prostaglandin precursor selectivity might vary during fish development (60) . The balance between arachidonate and eicosapentaenoate conversion to prostanoid in fish tissues is clearly under multifactorial regulation. The present characterization of the inherent substrate preferences of the two trout cyclooxygenases is an important step toward deciphering this regulatory scheme.

Determinants of substrate selectivity
The trout enzymes used in the present studies were expressed in recombinant form, solubilized with detergent, and purified by ion exchange chromatography to remove the bulk of host cell lipids. Negligible oxygen consumption was observed in control reactions lacking exogenous fatty acid (data not shown), confirming the absence of endogenous fatty acid substrates in the partially purified tPGHS-1 and -2. The results presented in Fig. 2 and Tables 3 and 4 thus show that the discrimination against 20:5n-3 and 22:6n-3 reported from studies of intact fish tissues and homogenates (24) originates in the structures of the PGHS proteins themselves, not from the influence of host cell lipids or of other proteins or factors present in fish tissues.

For each PGHS isoform, there is much more divergence between the trout and mammalian amino acid sequences than among mammalian sequences. For example, oPGHS-1 has only 68% identity with tPGHS-1, but 91% identity with hPGHS-1. Amino acid residues lining the cyclooxygenase pocket have been assumed to govern substrate specificity, and mutation of individual residues in the pocket can alter cyclooxygenase kinetics and product distribution (62 63 64) . However, examination of residues within 7.5 Å of bound arachidonate in the oPGHS-1 crystal structure (1DIY; (65) ) indicates very strong conservation: there is no position where the tPGHS-1 side chain differs from that in oPGHS-1 and where tPGHS-2 also differs from hPGHS-2. Thus, no single structural difference in the cyclooxygenase pocket can account for stricter substrate specificity in both trout enzymes compared with the mammalian enzymes; important protein structural determinants of fatty acid specificity remain to be identified.

Effects of aspirin
Aspirin acetylates Ser-530 in mammalian PGHS-1 and -2, leading to complete loss of oxygenase activity in PGHS-1 and to a shift from cyclooxygenase to LOX catalysis in PGHS-2 (16 17 18) . Generation of LOX products by aspirin-treated PGHS-2 may account for some of the pharmacological effects of the drug (19) . Ser-530 is conserved in both tPGHS-1 and -2 (26) , and aspirin treatment produced the expected time-dependent decline in arachidonate-dependent oxygen consumption in both recombinant enzymes (data not shown). However, LOX products did not increase for aspirin-treated tPGHS-2 with any of the four fatty acids tested. In addition, increased LOX catalysis by aspirin-treated hPGHS-2 was apparent only with 20:4n-6 and 22:4n-6, and not with 20:5n-3 or 22:6n-3. Product shifting by aspirin is clearly not a general phenomenon, but one limited by the structure of both the fatty acid substrate and of the PGHS-2 protein. The ability of mammalian PGHS-2, but not PGHS-1, to continue oxygenase catalysis after Ser-530 acetylation has been attributed to side chain differences at residues 434, 513, and 523 (66) . These three residues are conserved in mammalian and trout PGHS-2, so the lack of LOX activity in aspirin-treated tPGHS-2 must have another structural basis.

Regulation of cyclooxygenase catalysis
Distinct pathophysiological roles for PGHS-1 and -2 have been well established in mammals (3) . There is evidence of an analogous functional specialization of the two isoforms in fish, most prominently observations that PGHS-2 is inducible by mitogens and cytokines in a limited subset of cells in fish as in mammals (4 , 25 26 27) . In mammals, prostanoid synthesis by PGHS-1 and -2 is differentially controlled both at the level of gene expression and at the level of catalysis (4) . The differential catalytic control was dramatically demonstrated by the observation that prostanoid produced by activated macrophages and fibroblasts originates from catalysis by the induced PGHS-2 and not from constitutive PGHS-1 in the same cells (67 , 68) .

Several mechanisms have been proposed to explain the ability of PGHS-2 cyclooxygenase activity to proceed in cells where the PGHS-1 protein remains latent, including differences in compartmentation, in coupling to phospholipases or fatty acid carriers, in activation by peroxides, and in cooperativity toward substrate (6) . The appearance of considerable arachidonate in the medium in stimulated macrophages and fibroblasts (67) indicates that intracellular movement of unesterified arachidonate is not significantly restricted and argues against differential intracellular compartmentation of PGHS-1 and -2 or differential physical coupling with phospholipases or fatty acid carriers. In contrast, the more efficient cyclooxygenase activation by peroxide in PGHS-2 than in PGHS-1 readily accounts for PGHS-2 catalysis and PGHS-1 latency when both have access to arachidonate (5) . The lower activation efficiency in PGHS-1 can also account for its cooperative response to low arachidonate levels, emphasizing that peroxide and fatty acid levels are interdependent factors controlling cyclooxygenase catalysis (69) . Overall, the associations observed between individual phospholipases and PGHS isoforms appear to be more usefully analyzed in terms of kinetic coupling, based on the different responses of the two isoforms to cellular levels of peroxide and fatty acid rather than physical coupling between phospholipase and PGHS (70 , 71) .

A previous limitation to the hypothesis of differential regulation of PGHS-1 and -2 by cellular peroxide levels was that it was based on observations in a limited number of mammalian cells. The discovery that fish have two PGHS isoforms with functional specialization similar to those in mammals provided an opportunity for an independent test of the generality of the differential regulation hypothesis. The present observation that cyclooxygenase activation by peroxide is more efficient in tPGHS-2 than in tPGHS-1 (Fig. 6) demonstrates that the two trout isoforms also are differentially regulated by peroxide levels. Retention of the same regulatory mechanism in the widely divergent mammalian and fish species suggests that differential cyclooxygenase regulation by peroxides evolved as a functionally important strategy to control prostanoid synthesis in vertebrates.

Titration of PGHS cyclooxygenase activity with GPx-1 has proved to be a useful way of quantitatively assessing the efficiency of feedback activation by the peroxide product PGG2 (5 , 41 , 72) ; arachidonate is conventionally used as the substrate in these titrations. The results appear likely to reflect in vivo behavior both because GPx-1 is the major peroxidase in most cells (73) and because arachidonate is a favored cyclooxygenase substrate for both isoforms in both mammals and fish (Fig. 2) .

Differences in the side chain structure at position 383 can account for part of the higher cyclooxygenase activation efficiency in mammalian PGHS-2 compared to PGHS-1 (41) . However, this residue is conserved as a threonine in all known fish and mammalian PGHS-2 proteins, so the residue 383 structure cannot explain the lower activation efficiency in tPGHS-2. Measurement of the rates of the individual steps in cyclooxygenase activation process (74) for the trout enzymes will identify any kinetically impaired steps and may help target structural determinants linked to lower activation efficiency.

Cyclooxygenase catalysis and peroxide tone in fish
Each trout isoform had a lower activation efficiency than its mammalian counterpart (Fig. 6) . This indicates that higher peroxide levels are needed to maintain cyclooxygenase catalysis in trout and suggests that the trout enzymes have evolved to operate in higher ambient peroxide levels than their mammalian counterparts. Support for this concept comes from comparison of plasma phospholipid hydroperoxide levels in fish and humans. Phospholipid hydroperoxides are not themselves PGHS-1 activators (75) , but serve as an indicator of general lipid peroxide levels. High plasma phospholipid hydroperoxide levels (10–30 µM) were observed in some fish; other fish had lower levels (1.5–5 µM) (76) . In contrast, plasma phospholipid hydroperoxides are below 0.5 µM in healthy humans (77) , and estimates of total lipid peroxide levels in plasma from healthy humans range from 0.4 to 3 µM (78 79 80 81) .

In summary, the cyclooxygenase activities of individual trout and mammalian PGHS isoforms diverge in peroxide activator requirement and in preference for arachidonic acid, though the two isoform pairs share a differential response of PGHS-1 and -2 to peroxide activator. Thus, the two vertebrate families appear to have retained a functional biochemical mechanism for differential catalytic control by peroxide at this key point in prostanoid synthesis even as the PGHS proteins adapted to different substrate mixtures and ambient oxidant levels.


   ACKNOWLEDGMENTS
 
We thank Dr. Frederick Goetz for generously providing the cDNAs coding for brook trout PGHS-1 and -2, Mr. Stephen Gunther and Dr. Dominique Bureau for information on brook trout growth rates, Dr. William Dowhan for use of the PhosphorImager, Dr. Corina Rogge for providing the purified hPGHS-2, and Dr. William Lands for a critical reading of the manuscript. These studies were supported by National Institutes of Health grants GM 52170 (R.J.K.) and GM 38765 and DK 074448 (C.N.S.).


   FOOTNOTES
 
2 Current address: Department of Bioengineering, Rice University, P.O. Box 1892, Houston, TX 77251, USA.

Received for publication October 30, 2005. Accepted for publication January 25, 2006.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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