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An assay was set up for glyceryl ether monooxygenase activity in tissue samples using the novel substrate 1-O- pyrenedecyl-sn glycerol and high performance liquid chromatographic analysis of reaction mixtures with fluorescence detection, allowing robust detection of enzymatic activity in microgram amounts of tissue homogenates. The activity partially purified from rat liver strictly depended on presence of a tetrahydropteridine. Tetrahydrobiopterin-dependent glyceryl ether monooxygenase activity was observed in all rat tissues tested except female heart, with highest activities in liver, intestine and cerebellum. Activity was not uniformly distributed in brain, but was higher in cerebellum than in striatum or cortex. These data demonstrate that tetrahydrobiopterin-dependent glyceryl ether monooxygenase is not only found in liver and the gastrointestinal tract, but also in brain and other organs of the rat and provides an additional goal for tetrahydrobiopterin biosynthesis in these organs.
The reaction catalyzed by glyceryl ether monooxygenase (E.C. 18.104.22.168) is shown in Figure 1 (1). Glyceryl ether monooxygenase cleaves 1-O alkyl glycerol [I] by tetrahydropteridine - dependent hydroxylation at the carbon adjacent to the alkyl - glycerol ether bond. The primary hydroxylation product [II], a semi-acetal, rapidly rearranges to the corresponding aldehyde[III] and glycerol [IV]. In aerobic systems, the aldehyde is then oxidized to the corresponding carboxylic acid [V]. In cells, this reaction is catalysed by long-chain fatty aldehyde dehydrogenase (HUGO gene symbol ALDH3A2, E.C. 22.214.171.124.) by the aid of NAD+. Long-chain fatty aldehyde dehydrogenase, an enzyme ubiquitously expressed in the body, is defective in Sjörgen-Larsson syndrome (OMIM #270200), which is associated with ichthyosis, mental retardation and spasticity (2). Alternatively, the aldehyde may be enzymatically reduced to the corresponding alcohol [VI] by e.g. alcohol dehydrogenase class 4 (HUGO gene symbol ADH4, E.C. 126.96.36.199.). Despite this potentially multiple fate of the alkyl aldehyde product [III], the corresponding carboxylic acid [V] was always found to be the main product even when no NAD+ was added to the assay mixtures in all tissues investigated (3).
Similar to tetrahydropteridine - dependent amino acid hydroxylases, but in contrast to nitric oxide synthases (4), the tetrahydropteridine cofactor of glyceryl ether monooxygenase is thought to leave the reaction as 4a hydroxy derivative, which after dehydratation results in quinonoid dihydrobiopterin (6,7,[8H] dihydrobiopterin; qH2B). Quinonoid dihydrobiopterin is then recycled to tetrahydrobiopterin by dihydropteridine reductase (EC 188.8.131.52), which accepts both NADH and NADPH as reductants. Glyceryl ether monooxygenase reacts with substrates carrying 1-O alkyl side chains ranging from 11 to 20 carbon atoms, and requires one free hydroxyl group adjacent to the 1-O alkyl ether bond. The third hydroxyl group of glycerol may be coupled to various ligands, including common phospholipid side chains. For example, lyso PAF (lyso platelet activating factor; 1-O hexadecyl-sn-glycerophosphocholine), a degradation product of platelet activating factor, is a good substrate of glyceryl ether monooxygenase (1). Little is known about the physiological role of glyceryl monoalkyl ethers. They accumulate in proliferating cells and inhibit protein kinase C (5). Physiological significance and sequence of glyceryl ether monooxygenase are unknown. Its tissue distribution was thus far thought to be limited to liver and intestine (3). Several investigations characterized its activity in rat liver microsomes (6-11).
In this paper, we present a sensitive and robust assay of the enzyme based on the use of the novel compound 1-O- pyrenedecyl-sn glycerol as substrate, which is converted to 1-pyrenedecanoic acid by the glyceryl ether monooxygenase reaction (see Figure 1; n = 9, R = pyrenyl). We investigated cofactor requirements of glyceryl ether monooxygenase in purified fractions from rat liver microsomes and tissue distribution in rats.
Pyrenedecanol was prepared by Vitride reduction of pyrenedecanoic acid (from FLUKA AG, Buchs, Switzerland) according to standard protocols. Pyrenedecyl mesylate was obtained from pyrenedecanol and methanesulfonylchloride as described for the preparation of long-chain fatty alcohols in (12). A mixture of 13 mg (98,4 μmol) 3,2-isopropylideneglycerol (from Bachem, Bubendorf, Switzerland) and 14 mg KOH in 0.5 ml dimethylsulfoxide was stirred at 50 ° C for one hour. After addition of 50 mg (115,1 μmol) pyrenedecyl mesylate, stirring was continued for additional three hours. The reaction was stopped by addition of 1 ml water. The reaction mixture was extracted three times with diethylether (1 × 5 ml, 2 × 3 ml). The combined organic phases were washed three times with 2.5 ml water and dried over sodium sulphate, followed by removal of the solvent under reduced pressure. The crude 1-O- pyrenedecyl-2,3-isopropylidene-sn-glycerol product showed one main spot on thin layer chromatography (Rf 0.4; silica gel; solvent: methylene chloride) and was used without further purification for the next step: After addition of 800 μl methanol, 120 μl 3 N HCl was added under stirring, yielding a precipitate. The reaction mixture was stirred for three hours and the solvent was removed under reduced pressure. The crude product was purified by preparative thin layer chromatography on silica gel (Rf 0.13) using petroleum ether - diethylether (2/8, v/v) as solvent. The yield of pure product was 18 mg (41,8 μmol), which was stored in aliquots at − 20 °C. While our work was in progress, a report appeared that used a related compound (1-O-[9′-(1″-pyrenyl)]nonyl sn-glycerol) to study the metabolic fate of etherlipids in cells (13).
DL-alpha hexadecyl glycerol was from FLUKA, lyso PAF (1-Hexadecyl-sn-glycero-3-phosphocholine) was from Sigma. R,S-1- Octadecylglycerol (+,− batyl alcohol) was a kind gift of Wilf Armarego, Australian National University, Canberra, Australia. Dihydropteridine reductase from Physarum polycephalum was expressed in E. coli, purified by DEAE chromatography as described (14) and stored at − 20 °C in presence of 50% (v/v) glyercol. Pteridine derivatives were obtained from B. Schircks Laboratories, Jona, Switzerland. Methanol for chromatography and buffer salts were from Merck, Darmstadt, Germany. Catalase (from bovine liver) and nucleotides were from Sigma, Vienna, Austria.
A typical glyceryl ether monooxygenase assay contained the following components in a total volume of 10 μl: 100 mM Tris.HCl, pH 8.5, 1 to 12 μg protein, 0.2 mM NADPH (to promote recycling by dihydropteridine reductase), 0.2 mM NAD+ (to promote oxidation of the aldehyde by ALDH3A2), 0.1 mg/ml catalase (purified by NAP-5 Sephadex G-25 columns, Amersham-Pharmacia, Uppsala, Sweden), 0.5 μmol min−1 ml−1 (0.2 μg/ml) dihydropteridine reductase, 0.2 mM 6R 5,6,7,8-tetrahydro-L-biopterin and 0.1 mM 1-O- pyrenedecyl-sn-glycerol. The reaction was started by addition of protein, and incubation was performed at 37°C in the dark. After 5 to 80 min (40 min for rat tissue samples, above 60 min the reaction was no longer linear in crude homogenates), the reaction was terminated by addition of 30 μl methanol. In assays using rat liver microsomes, addition of NAD+ did not alter the yield of pyrenedecaonic acid and was omitted from the reaction mixtures.
Tissue samples were homogenized in 500 μl 0.1 M Tris.HCl, pH 7.6, containing 0.25 M sucrose and 1 mM PMSF by means of an Ultra Turrax (IKA, Staufen, Germany). After centrifugation at 16 000 g for 2 min at 4 ° C, supernatants were used for determination of protein (Bradford assay, bovine serum albumin standard) and glyceryl ether monooxygenase activity assays. Tissue extracts were diluted to yield less than 0.4 mg protein/ml final concentration in the assay. Samples were spiked with solubilized rat liver microsomes (10% v/v, 0.02 mg protein/ml final concentration) to control for interfering substances potentially present in homogenates. Increase in activity in tissues by spiking was compared to the same amount of solubilized rat liver microsomes diluted in assay buffer. A control sample was generated by stopping the reaction immediately after its start, which yielded always less than 2% pyrene decanoic acid product compared to pyrene decanoic acid levels reached after 40 min of incubation. Reagent blanks (homogenization buffer instead of protein added) were run in parallel and were always negative for pyrene decanoic acid (< 1 nmol/l). Tissues for female and male animals were analysed in a mixed sequential order to avoid bias by drifting of assay and/or analysis conditions with time.
For testing interference of lipids in the assay, 5 mM stock solutions were prepared in methanol and further diluted in assay buffer before added to solubilized microsomes. Control samples contained the same amount of methanol but no lipid added.
A Hewlett Packard 1050 HPLC system with Chemstation software (Agilent, Vienna, Austria) was used. 4 μl sample were injected onto a Zorbax XDB-C8 USP-L7 column (Agilent, Vienna, Austria) eluted with 10 mM potassium phosphate buffer, pH 5.4, containing 79 % (v/v) methanol at a flow rate of 1.0 ml/min for 4.5 min, followed by a gradient to 100 % (v/v) methanol at 5.0 min. After elution with 100 % methanol until 7.5 min, the initial composition (79 % methanol) was restored at 8.0 min, and the column equilibrated until 12.5 min total time to prepare for the next injection. Pyrenedecaonic acid was detected by fluorescence (detector FP-920, Jasco, Vienna, Austria) at excitation 340 nm, emission 400 nm, with a detection limit of 1 nmol/l.
All procedures were carried out at 4 °C. 250 g rat liver (male Sprague Dawley) were homogenized in 375 ml 0.1 M Tris.HCl, pH 7.6, containing 0.25 M sucrose and 1 mM PMSF by means of an Omni Mixer (Sorvall) and centrifuged at 3000 g for 10 min. Supernatants were collected and centrifuged at 13 000 g for 20 min. Microsomal pellets were collected after centrifugation at 40 000 g for 1 hour. After washing with Tris.HCl/sucrose buffer, microsomal pellets were resuspended in 10 ml assay buffer (100 mM Tris.HCl, pH 8.5) containing 10 % (v/v) glycerol, yielding a total protein concentration (Bradford assay) of 11 mg/ml. Microsomes were then solubilized in an equal volume of glycerol and 3 volumes (referring to the initial volume of microsomes) of water containing 1 mM dithiothreitol (DTT) were added. After centrifugation at 40 000 g for 45 min, ethanol was added to the supernatant under stirring to a final concentration of 28 % (v/v), incubated for 30 min, and the pellet was removed after centrifugation at 5 000 g for 15 min. Under stirring, ethanol was added to yield a final concentration of 37.5 % (v/v), and the pellet collected by centrifugation at 5 000 g for 15 min. The pellet was then resuspended in 10 mM Tris. HCl, pH 7.0, containing 10 % (v/v) glycerol and 1 mM DTT. The enrichment in specific activity was about 100 fold compared to crude liver homogenate with a typical overall yield of 0.5 %.
All animal work was conducted in conformity with the Public Health Service (PHS) Policy on Humane Care and Use of Laboratory Animals, and approved by the respective committe of the Austrian Ministry of Science and Education. Sprague Dawley rats (6 female, 231 - 252 g, 6 male 244 - 346 g) were anaesthetized with urethan (i.p.) and decapitated. Tissues were immediateley frozen in liquid nitrogen and stored at −75 °C until analysed.
Figure 2A shows HPLC chromatograms of typical incubation mixtures of glyceryl ether monooxygenase assays in partially purified fractions of rat liver microsomes. The 1-O- pyrenedecyl-sn glycerol substrate is well separated from the product 1 pyrenedecanoic acid. Furthermore, the substrate is virtually free of 1 pyrenedecanoic acid, thus allowing for a sensitive detection of product in incubation mixtures (detection limit 1 nmol/l). Both, substrate and product, were stable to laboratory light for up to 4 hours. Nevertheless, enzyme incubations are performed routinely in the dark to protect the light sensitive pterin cofactors. Previous work has shown that a small amount of alcoholic product is made in addition to the dominant acid product (3). We cannot detect the alcohol in our HPLC chromatograms, however, since it comigrates with the substrate.
With crude rat liver homogenates as source of glyceryl ether monooxygenase, we found the enzymatic product formation was linear with protein amount up to 4 μg, which corresponds to 0.4 mg/ml final homogenate protein concentration in our 10 μl total volume assay (Figure 2B). Similar observations were made with each of the rat tissues tested (data not shown, for a list of tissues tested see Table 2). With purified fractions, the assay is linear up to 12 μg (1.2 mg/ml final protein concentration, not shown). Product increased linearly up to 60 minutes in tissue homogenates (Figure 2C) and up to 80 minutes using purified fractions (not shown). Since we found that ammonium sulfate (5 mM) inhibited the reaction already by 80 %, we omitted this substance that had been included in previous assay protocols from the assay incubation mixture (see Materials and Methods section for details).
1-O-[9′-(1″-pyrenyl)]nonyl-sn-glycerol , a compound similar to our substrate with a nine carbon spacer instead of a ten carbon spacer between glycerol and the pyrenyl fluorophore, is equally suited for the assay. It exhibits a Km (6.2 ± 1.9 μM) comparable to our 10 carbons spacer ether lipid Km (8.9 ± 2.0 μM, mean ± SD, n = 3).
Pyrenedecanoic acid formation was competitively inhibited by hexadecylglycerol (Ki = 12.4 μM). Lyso PAF and octadecyl glycerol also inhibited the pyrene decanoic acid formation. However, more than 85% of pyrenedecanoic acid was recovered when 20 μM of either of the two compounds were added. To control for interference of lipids present in tissue extracts, we spiked all tissue extracts with microsomal preparations and found an average of 75 % recovery (see below).
With a microsomal fraction further purified by ethanol precipitation, we tested for substrate and pteridine cofactor dependence. The enzyme had an apparent Km of 8.9 ± 2.0 μM for the substrate, and 2.6 ± 1.6 μM for tetrahydrobiopterin (mean ± SD, n = 3). Tetrahydrobiopterin was absolutely required for the reaction. In absence of tetrahydrobiopterin, we found less than 1 % of the activity observed in presence of tetrahydrobiopterin (see Figure 2 for an example of respective chromatograms). Tetrahydrobiopterin could not be replaced by either of the following reductants: glutathione (reduced form), FADH2 (formed from FAD by reduced glutathione in presence of glutathione reductase) or L-ascorbic acid (1 mM each), which all displayed activities below 1 % of full activity observed in presence of tetrahydrobiopterin (data not shown). The enzyme activity showed clear preferences for stereochemistry of tetrahydropteridine side chains (Table 1). For example, 6S tetrahydrobiopterin was inactive as cofactor, whereas 6 RS tetrahydroneopterin was a slightly better stimulator of activity as compared to 6R tetrahydrobiopterin.
Table 2 lists glyceryl ether monooxygenase activities in tissues of female and male rats. Highest activities were observed in liver, followed by intestine, cerebellum and testes. Activity was not uniformly distributed in the brain, but was higher in cerebellum as compared to cortex or striatum. Male rats had a higher activity in the liver (p > 0.001), stomach (p < 0.05) and heart (P < 0.02, student’s t-test) as compared to female rats. Tetrahydrobiopterin dependence was observed to a variable degree in all tissues except the female heart. Tetrahydrobiopterin dependence was high in tissues with high activity and declined to about 50 % in tissues with low activity (Table 2).
To control for a potential attenuation of the readout of our assay by endogenously present, non fluorescent substrates, we spiked all tissues with solubilized liver microsomes and observed recovery of activity when spiking tissue extracts in comparison to spiking of the same amount to assay buffer. Mean recovery was 75% in both female and male tissues. Recovery did not correlate with the activity, i.e. was not higher in liver as compared to tissues with much lower activity (Table 2).
To assess glyceryl ether monooxygenase activity in cells and tissues, we have developed a novel assay based on HPLC separation of the fluorescent pyrenedecanoic acid product from the novel substrate 1-O- pyrenedecyl-sn glycerol, which was chemically synthesized to enable this study. Due to the intense fluorescence of the product, our newly developed assay allows a sensitive, robust detection of glycerylether monooxygenase activity. 1-O-[9′-(1″-pyrenyl)]nonyl-sn-glycerol, which had been synthesized for other purposes (13), appears to be equally suited as 1-O- pyrenedecyl-sn glycerol as substrate. However, in contrast to pyrene decaonic acid, the resulting pyrene nonaic acid is not commercially available for calibration of the HPLC method.
In a previous assay based on kinetics of UV absorption, a detection limit of activity of 0.6 nmol/min was found (9). Our assay, in contrast, had a 5 orders of magnitude lower detection limit of 4 fmol/min. This difference arises from our miniaturized assay volume (10 μl instead of 1 ml), and the nanomolar detection limit for the fluorescent substrate as compared to the micromolar concentration changes observed by UV absorption. Other assays used 3H or 14C labelled substrate, thin layer chromatography, scraping and counting of spots (6) or gas-liquid chromatography and detection of labelled CO2 following combustion (3). No minimal detectable activity was mentioned in these reports, but we assume that these assays are sensitive if high radioactive concentrations of substrates are used. However, they appear to be much more laborious and time consuming than our HPLC based assay.
Our competition experiments show that our fluorescent substrate has comparable affinity to the enzyme as hexadecylglycerol, the best substrate of the enzyme (1). Thus, etherlipids present in samples would potentially interfere with the reaction when present in comparable amount to the substrate used. Due to the high sensitivity of the assay, however, dilute tissue extracts were used. We spiked all tissue extract samples with solubilized rat liver microsomes, and found a mean recovery of 75 %. This recovery was not higher in the liver, which had the highest activity, as compared with tissues with much lower activity. These data clearly demonstrate that the differences in glyceryl monooxygenase are not biased by different amounts of interfering compounds potentially present in tissues.
Using a partially purified preparation of glyceryl ether monooxygenase, we could demonstrate that among a panel of other reductants or coenzymes tested, the presence of a tetrahydropterin was essentially required for catalysis. The stimulatory effect of tetrahydropteridines depended on the stereochemistry of the side chain. For example, 6R tetrahydrobiopterin was a very good stimulator, whereas 6S tetrahydrobiopterin was not able to stimulate the enzyme. These results extend previous observations (reviewed in (1)). Since the ability of tetrahydropteridines to stimulate glyceryl ether monooxygenase activity do not correlate with the ability of our dihydropteridine reductase preparation to recycle the various pteridine derivatives (14), which was used in large excess of activity, they clearly suggest a direct interaction of the tetrahydropteridine cofactor with glyceryl ether monooxygenase. We found micromolar Km levels for substrate and cofactor, which are up to ten-fold lower than values found in previous investigations (reviewed in (1)). These differences may originate from differences in the scale, setup and precision of the assay.
In rats, the highest glyceryl ether monooxygenase activities were found in liver, and the intestines. In accordance a previous report, liver of male rats had about three fold higher activity than liver of female rats (15). Interestingly, this difference was confined to a few organs, i.e. liver, heart and stomach, but not reflected in other organs. Activity varied among brain regions, and was higher in cerebellum than in striatum or cortex. With the exception of female heart, tetrahydrobiopterin dependent glyceryl ether monooxygenase activity was detected in all organs investigated. This contrasts previous widely accepted results, that glyceryl ether monooxygenase was only observed in significant activity in liver and intestine (3). We attribute this discrepancy to the better performance of our assay.
Although low residual activity was detected in the tissue homogenates without addition of tetrahydrobiopterin, glyceryl ether monooxygenase activity in these organs might depend totally on tetrahydrobiopterin, if the enzyme activity was partially purified from these tissues, as we have shown here for the liver. The low residual activities may be enabled by the tetrahydrobiopterin cofactor present in the tissues, which were not removed from the homogenates before incubation with the tetrahydrobiopterin-free assay mixture. Alternatively, tissues may contain a low non-tetrahydrobiopterin dependent glyceryl ether monooxygenase activity similar to the one found in Leishmania (16), explaining the decreasing tetrahydrobiopterin-dependence we observed with decreasing activity.
The glyceryl ether monooxygenase activity profile we found in the rat indicates that the enzyme may serve a metabolic function due to its presence in gastrointestinal tract and liver. Localization also in other areas like in testes and brain indicates that glyceryl ether cleavage is also required in peripheral organs, for functions remaining to be investigated. The broad tissue distribution of glyceryl ether monooxygenase should also be considered when interpreting effects of tetrahydrobiopterin treatment and correlation of tetrahydrobiopterin biosynthesis with tetrahydrobiopterin-dependent activities in mammalian tissues. It remains to be determined to what extent tetrahydrobiopterin deficiency related symptoms in mice (17, 18) and humans (19) are caused by impairment of glyceryl ether monooxgenase activity.
This work was supported by the Austrian Research Funds “zur Förderung der wissenschaftlichen Forschung”, Project 19764. The authors are indebted to Petra Loitzl, Nina Madl and Renate Kaus for expert technical assistance. The authors thank Beat Thöny, University Childrens Hospital Zürich, Switzerland, for helpful discussions and Raphael A Zoeller, Department of Physiology and Biophysics, Boston University School of Medicine, for his kind gift of 1-O-[9′-(1″-pyrenyl)]nonyl-sn-glycerol.