Prostaglandins are derived from arachidonic acid (AA) in a pathway dependent on the PGHS (EC 1.14.99.1) family of enzymes, which are commonly known as cyclooxygenase (COX), referring to the first step of enzymatic activity. PGHS converts AA to prostaglandin H₂ (PGH₂), the precursor of all prostanoids. The enzyme contains two active sites: a COX site, where AA is converted into the hydroperoxy endoperoxide, prostaglandin G₂ (PGG₂), and a haem with peroxidase activity that reduces PGG₂ to PGH₂ (For review see 1). The reduction of PGG₂ by the peroxidase element generates the corresponding alcohol. This reaction has previously been demonstrated to concurrently oxidise aminopyrine molecules to aminopyrine free radicals 2. Here, a spin-trapping agent, 1-hydroxy-3-carboxy-pyrrolidine (CPH) is oxidised to 3-carboxy-proxy (CP), probably under the action of the peroxidase, in a similar fashion to that previously seen with aminopyrine (Fig. 1).

Two structurally similar PGHS isoforms exist (PGHS-1 and PGHS-2) which are encoded by different genes and the expression of which varies between tissues 3. PGHS-1 is often referred to as the 'house-keeping' isoform due to its regulatory functions in many tissues. PGHS-2 is virtually undetectable under normal conditions in most tissues and is often referred to as the 'inducible' isoform due to its tendency to be expressed in response to inflammatory stimuli. The exception to this is in the brain and spinal cord, where PGHS-2 is constitutively expressed and plays a role in nociception signaling 4.

The importance of PGHS as a therapeutic target has long been highlighted by the actions of aspirin, 56 the first drug of the family of nonsteroidal anti-inflammatory drugs (NSAIDs) for use as analgesics, anti-inflammatory agents and antithrombotic agents. In contrast to other NSAIDs, such as indomethacin, which reversibly bind at the COX active site 7, aspirin causes an irreversible inhibition of PGHS by rapidly and selectively acetylating the hydroxyl group of a serine residue (Ser 530) near the C-terminus of the enzyme, forming an impediment to the binding of AA 8910. The ensuing irreversible PGHS inhibition requires de novo synthesis of the enzyme for subsequent production of prostaglandins.

Interest in PGHS has been re-ignited recently on account of two advances in the development of novel NSAIDs. Firstly, nitroaspirins 11121314 are being developed in an effort to overcome the gastrotoxic side-effects of aspirin that represent the major limitation to its therapeutic use 151617. Nitroaspirins make use of the protective effects of nitric oxide (NO) to compensate for the potentially damaging impact of aspirin-mediated depletion of protective prostaglandins in the gastric mucosa. Secondly, the suggestion that the gastrotoxic side effects of aspirin are due to the inhibition of housekeeping PGHS-1, whereas its anti-inflammatory effects are due to inhibition of PGHS-2, led to the development of selective inhibitors of the COX-2 activity of PGHS-2, in the hope that the beneficial effects could be retained without injury to the gastric mucosa 18. However, recently, several of these new selective PGHS-2 inhibitors have been withdrawn due to mounting evidence of an increased risk of stroke. The increased risk of thrombus is thought to be due to inhibition of PGHS-2 in endothelial cells leading to down regulation of anti-thrombotic prostaglandins (such as prostacyclin) in relation to the unaffected PGHS-1 derived thromboxanes 192021.

Given the intense interest in PGHS activity and inhibition, it is perhaps surprising that relatively few assays for the activity of this enzyme have been developed. Amongst the most popular techniques is the measurement of thromboxane B₂ (TXB₂), the stable metabolite of PGH₂-derived TXA₂, as a marker of PGHS activity 22. Alternatively, measurement of oxygen consumption using an oxygen electrode 2324, assays using radio-labelled substrate 25 and immunoassays for the prostaglandin products 23 can also be applied. More recently, a commercial chemiluminescent assay (Axxora, Nottingham, UK) has been developed for use on purified enzyme. The assay involves the use of labelled substrate that generates a luminescent product under the action of the hydroperoxidase element of PGHS, most likely via the generation of oxidising free radicals.

Electron paramagnetic resonance (EPR) has been previously utilized to measure free radicals generated by PGHS as a measure of its activity 2627. However these techniques can involve complex mechanisms to trap the short-lived radical species. For instance in an in vitro assay using purified PGHS, liquid nitrogen was utilised to stop the reaction and was required to stabilise the tyrosyl radical species generated, in order that it could be recorded by EPR 26. Another technique which recorded PGHS activity in mouse keratinocytes relied on the use of the antioxidant glutathione to stabilize the generated radical before trapping it with DMPO 27.

Here, we present a novel method for assaying the activity of PGHS-1 in which EPR is used to detect the stable adduct CP, formed from oxidation of commonly available spin-trap, CPH 28, under the action of the peroxidase element of PGHS-1. The aim of the studies described was to validate this technique for assaying PGHS-1 activity in vitro, to determine its effectiveness at establishing the inhibitory effects of conventional NSAIDs and to confirm that a COX-2-selective inhibitor was ineffective in this assay.

Unless otherwise stated, all drugs and chemicals were purchased from Sigma, Dorset, UK. The assay was performed at 37°C in 1 ml of Tyrode's buffer (137 mM NaCl, 2.7 mM KCl, 1.05 mM MgSO₄, 0.4 mM NaH₂PO₄, 12.5 mM NaHCO₃, 5.6 mM Glucose, 10 mM HEPES and 0.8 mM CaCl₂ in deionised water at pH 7.4. 100 units/ml ovine seminal PGHS-1 was incubated with 1.5 μM haematin (5 min, 37°C) prior to the assay. Data from the manufacturers of the PGHS-1 reveals that 1 unit of enzyme consumes 1 nanomole of oxygen at 37°C in the presence of 1 μM haematin and 100 μM AA. Aspirin, salicylic acid, indomethacin, NS398 (Merck Biosciences, Nottingham, UK) or vehicle control (DMSO, 1%) was added and left to incubate for a further 10 min prior to addition of the spin-trap, CPH (1 mM; Axxora). At this point (t = -2 min), a baseline EPR measurement was taken (MS200, Magnettech, Germany. Instrument settings: B0-field, 3356 gauss; sweep width; 50 Gauss, sweep time, 30 sec; modulation amplitude, 1500 mGauss; microwave power, 20 mW). 2 min later, 100 μM AA (as sodium salt) was added (t = 0). Further EPR readings were taken at t = 1.5 (the earliest timepoint at which readings could consistently be taken after addition of AA), 4 and 6 min. The suicidal nature of PGHS-1 activation means that the period of activation is anticipated to be complete within ~1 min of AA addition 3.

The results are corrected for any auto-oxidation of spin-trap by subtraction of values recorded from a duplicate sample run in the absence of AA. The intensity scale on the y-axis of all graphs is an arbitrary scale based upon the area under the curve of the first derivative traces generated.

Here we have validated a new, quick, simple and reproducible method for detecting PGHS-1 activity and inhibition in vitro. The principle of the assay was based on the concomitant oxidation which occurs with the reduction of PGG₂ by the peroxidase element of PGHS 2. This oxidation can be exploited in the absence of antioxidant glutathione to oxidise spin-trapping agent CPH to the stable adduct, CP which generates a characteristic 3-line EPR spectrum. The amplitude of the EPR signal is proportional to the amount of adduct generated.

Our results indicated that oxidation of CPH by the isolated PGHS-1 enzyme upon addition of the enzyme substrate, AA, was sufficient to be easily detectable by EPR. The initial peak in the detected signal recorded at the first reading, had by the subsequent time-points slowed to a rate that was equivalent to the signal generation from substrate-free enzyme, most likely due to autoxidation of the spin trap. The loss of specific enzyme-mediated radical generation is unsurprising, given the well-recognised suicidal nature of activated PGHS-1 3. From these time-course experiments, we selected a 1.5 min time-point for subsequent comparative studies because by this time the AA-dependent free radical generation was complete but the signal was not significantly enhanced by autoxidation of the spin trap.

Aspirin pretreatment (10 min) was shown to have a concentration-dependent inhibitory effect in the published range 30, but the acetyl-free counterpart, SA, failed to significantly inhibit enzyme activity. The lack of effect of SA indicates that aspirin-mediated inhibition of the enzyme is dependent on the acetyl group, the moiety involved in PGHS inhibition, and not due to a non-specific antioxidant effect. SA was not expected to have any effect in this assay because it is known not to affect PGHS-1 or 2 activity 31 except in intact cells 30, possibly by suppressing PGHS-2 transcription in response to exogenous stimuli 32. Furthermore, the reversible, non-specific PGHS inhibitor, indomethacin 33 was demonstrated to have a powerful inhibitory effect, whilst the PGHS-2 specific inhibitor, NS398 at a concentration known to inhibit PGHS-2 29 failed to do so. Signal intensities shown in the figures are after subtraction of an AA-free control from a parallel experiment to account for autooxidation of the spin trap. A negative signal (as observed with indomethacin for example) therefore indicates that the control value was greater than the AA-treated sample, which might suggest that AA has a slight antioxidant effect on its own.

These results confirm that the assay is relevant for NSAIDs with different modes of inhibitory action. As purified PGHS-2 is now available commercially (Sigma), it may be possible to adapt this assay system to help determine the specificity of novel PGHS-2 inhibitors.

This assay provides a convenient screening method for inhibitors of the PGHS enzyme. Whilst various techniques exist to assay the activity of PGHS isoforms, each has its own disadvantages. For instance, recording the oxygen uptake by PGHS is an option to measure its activity but this requires high enzyme concentrations and also accurate control of the initial oxygen concentrations 24. Optical techniques are prone to interference from coloured assay constituents (such as haematin) and require high AA concentrations. Techniques recording uptake of radiolabelled substrate 25 can be complex and expensive. Our technique provides quick inexpensive results that give real-time determination of COX-inhibition. Data obtained, as demonstrated by percentage inhibition of control response achieved with aspirin, is comparable to other in vitro techniques 343536.

The technique described in the present study also offers a simpler alternative than previous EPR techniques where generated radicals are detected following complex radical stabilization steps 2627 and thereby providing potential for less loss of signal. Tsai et al. 26 used dry ice and liquid nitrogen to stop the reaction and trap the radical. The recording of spectra was then carried out under liquid nitrogen. By comparison, our method uses direct oxidation of a spin-trap to generate an adduct that is sufficiently stable to allow successive time-point readings to be taken without the need to freeze the sample at the required time-point. The method by Schreiber et al. 27 did use a spin-trap, but required the use of glutathione to reduce an amine radical to the parent amine, liberating a thiyl radical which was trapped by DMPO. In our method, the spin-trap is oxidised directly, thus reducing the potential for loss of signal. Furthermore, there is some evidence of rapid signal decay in the Schreiber method; the spectra show a decrease of the DMPO-thiyl signal during the course of the measurement, whereby the spectral lines no longer conform to the expected 1:2:2:1 ratio. By comparison, the EPR signal generated in the present technique is much more resilient and does not decay during the time-course of experiments.

It is important to recognize the potential limitations of this assay. Its reliance on the oxidation of CPH might preclude its use in cells, given that the cellular environment is usually very rich in antioxidants such as GSH, and its presence at intracellular concentrations (~5 mM) could effectively compete out the spin-trap and nullify the assay. The susceptibility of polyunsaturated fatty acids such as AA to peroxidative attack by reactive oxygen species may impact on the recorded level of radical if radicals are consumed by free AA in the samples 3738. Furthermore, it is important to recognize that the in vitro nature of the assay does not account for any absorption, metabolic or availability issues that might relate to applied NSAIDs.

In summary, we have validated a new, simple, EPR-based assay for detecting PGHS-1 activity and inhibition. We have demonstrated it to be sensitive to the inhibitory effects of conventional NSAIDs (aspirin and indomethacin) and not to SA or a PGHS-2 specific inhibitor. In principle, this assay should be equally applicable to measuring PGHS-2 activity in isolated enzyme. As such, this assay might prove to be a useful research tool in the ongoing search for novel PGHS inhibitors of both isoforms of the enzyme.

AA, arachidonic acid; COX, cyclooxygenase; CPH, 1-hydroxy-3-carboxy-pyrrolidine; EPR, electron paramagnetic resonance; NO, nitric oxide; NSAID, nonsteroidal anti-inflammatory drugs; PGG₂, prostaglandin G₂; PGH₂, prostaglandin H₂; SA, salicylic acid; TXB₂, thromboxane B₂.

The author(s) declare that they have no competing interests.

Experimental design and procedures were performed by CMT and assisted by DM. AGR and ILM supervised experimental design and procedures. AGR and ILM oversaw manuscript construction, revising it critically for important intellectual content. All authors have given final approval of the version to be published.