methods for lipase detection and assay: a criticalcl.riviere.free.fr/lastlipub.pdf · 2001. 8....

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Eur. J. Lipid Sci. Technol. 2000, 133–153 133

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iew

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Frédéric Beisson*, AliTiss*, Claude Rivière,Robert Verger

Laboratoire de LipolyseEnzymatique (UPR 9025 duCNRS), Institut de BiologieStructurale et Microbiologie,Marseille, France

Methods for lipase detection and assay: a criticalreview1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1342 Physico-chemical methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1352.1 Disappearance of substrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1352.1.1 Nephelometry and turbidimetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1352.1.2 The Wilhelmy plate method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1352.1.2.1 Pure monomolecular films as substrates . . . . . . . . . . . . . . . . . . . . . . . . 1352.1.2.2 Mixed monomolecular films as substrates . . . . . . . . . . . . . . . . . . . . . . . 1362.1.2.3 Interfacial binding and film recovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1362.1.3 Atomic force microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1372.1.4 Infrared spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.2 Appearance of hydrolytic reaction products . . . . . . . . . . . . . . . . . . . . . . 1382.2.1 Proton release as an indirect assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.2.1.1 Coloured indicators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.2.1.2 Titrimetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382.2.2 Analysis of the free fatty acids released. . . . . . . . . . . . . . . . . . . . . . . . . . 1392.2.2.1 Glycerol derived carboxylic esters as substrates . . . . . . . . . . . . . . . . . . 1392.2.2.1.1 Colourimetric assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1392.2.2.1.2 Fluorimetric assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1392.2.2.1.3 Chromatographic assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1402.2.2.1.4 Enzymatic assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1402.2.2.1.5 In situ detection by electron microscopy . . . . . . . . . . . . . . . . . . . . . . . . . 1412.2.2.2 Synthetic carboxylic esters as substrates . . . . . . . . . . . . . . . . . . . . . . . . 1412.2.2.2.1 Radioactive assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1412.2.2.2.2 Colourimetric assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1422.2.2.2.3 Fluorimetric assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1432.2.3 Determination of the released alcohol or thiol . . . . . . . . . . . . . . . . . . . . . 1432.2.3.1 Determination of the released glycerol from TAG . . . . . . . . . . . . . . . . . . 1432.2.3.2 Determination of the released coloured or fluorescent alcohol from

synthetic carboxylic esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1432.2.3.3 Determination of the released thiol from synthetic thioesters . . . . . . . . . 1442.2.4 Electric conductivity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1442.2.5 Acoustic wave conductance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1442.2.6 The oil-drop method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1453 Immunological methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1453.1 ELISA for pure lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1463.2 Lipase immunodetection on physiological media . . . . . . . . . . . . . . . . . . 1463.2.1 Lipase immunodetection in plasma or serum . . . . . . . . . . . . . . . . . . . . . 1463.2.2 Lipase immunodetection in other physiological media . . . . . . . . . . . . . . 1463.3 Immunocytolocalisation of lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1463.4 Immunoblot analysis of lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1464 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1474.1 Pure lipases or crude biological media . . . . . . . . . . . . . . . . . . . . . . . . . . 1474.2 Level of lipase acitity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147

Correspondence: Robert Verger, Laboratoire de Lipolyse Enzy-matique (UPR 9025 du CNRS), Institut de Biologie Structurale etMicrobiologie (IFR1 du CNRS et de l’Université de la Médi-terranée), 31 Chemin Joseph-Aiguier, 13402 Marseille Cedex20, France. Phone: +33 4 91 16 40 93; Fax: +33 4 91 71 58 57; e-mail: [email protected].* These authors have made equal contributions to the present

review.

Abbreviations: AFM, atomic force microscopy; BSA, bovineserum albumin; ELISA, enzyme-linked immuno sorbent assay;FFA, free fatty acids; FTIR, Fourier transform infrared spec-troscopy, HGL, human gastric lipase; HLL, Humicola lanuginosalipase; HPL, human pancreatic lipase; IU, international units(µmoles of FFA released per min.); LPL, lipoprotein lipase; PPL,porcine pancreatic lipase; TAG, triacylglycerol.

1 Introduction

Lipids constitute a large part of the earth’s biomass, andlipolytic enzymes play an important role in the turnover ofthese water-insoluble compounds. Lipolytic enzymes areinvolved in the breakdown and thus in the mobilisation oflipids within the cells of individual organisms as well as inthe transfer of lipids from one organism to another. Oneimportant aspect of lipolytic enzymes is the unique physi-co-chemical character of the reactions they catalyse atlipid-water interfaces, involving interfacial adsorption andsubsequent catalysis sensu stricto. Most of the lipasesare water-soluble enzymes acting on water-insoluble sub-strates (supersubstrates). The heterogeneous characterof this catalysis makes it difficult to accurately quantitateboth the amount of interface (specific surface) [1] as wellas the interfacial parameters (such as the interfacial ten-sion, surface viscosity, surface potential, etc.) responsiblefor the “interfacial quality” of the substrate [2–5], on whichthe lipolytic process greatly depends. The emulsificationof the water-insoluble substrates, which requires thepresence at the interface of surface active amphiphilessuch as detergents, other lipids, proteins, etc., can there-fore drastically influence lipase activity measurements:non-specific inhibition of lipases by proteins present at theoil/water interface is one well-known phenomenon of thiskind [6–10].

Lipases were previously defined in kinetic terms, basedon the “interfacial activation” phenomenon, i.e., on the in-crease in the activity which occurs when a partially water-soluble substrate becomes water-insoluble [11]. Thisprocess was not observed among esterases. The recent-ly determined 3-D structures of lipases show an α/βhydrolase fold as well as a nucleophilic elbow where thecatalytic serine is located [12, 13]. Some, but not all lipas-es show a “lid” controlling access to the active site. How-ever, the above structural features, including the pres-ence of a lid as well as the “interfacial activation” phe-nomenon, are not suitable criteria for classifying specificesterases as lipases. Since enzymes are usually namedafter the type of reaction they catalyse, lipases can bepragmatically redefined as carboxylesterases acting onlong-chain acylglycerols: they are simply fat-splitting ’fer-ments’ [14].

Lipases are ubiquitous enzymes [6, 15–17] which arefound in animals, plants [18, 19], fungi [20] and bacteria[21–23]. In the industrially developed countries, the ediblelipids present in the human diet, which consist mainly oftriacylglycerols (TAGs), from 100 to about 150 grams perday, i.e. 30% of each individual’s daily caloric intake. TAGmolecules cannot cross the intestinal barrier. A series ofhydrolytic and absorption stages are therefore necessaryto produce the chemical energy resources present in the

hydrocarbon chains of biologically usable TAGs. The li-pases in the digestive tract therefore play a particularlyimportant role in nutrition processes, in both humans andhigher animals.

Lipases are now being widely used as enantioselectivecatalysts in aqueous as well as in low-water media, andvarious synthetic molecules can serve as their substrates.In this review, we will deal only with lipase assays involv-ing carboxylic ester hydrolysis and not with the methodsdesigned for studying alcoholysis, acidolysis, ester syn-thesis, or inter- and transesterification reactions. The lat-ter reactions are usually performed under low water activ-ity conditions [24–30]. Furthermore, the reader is referredto published articles concerning the stereoselective as-says involving lipases [31–40].

As can be seen from the literature [6, 41, 42], numerousmethods are available for measuring the hydrolytic activi-ty as well as for the detection of lipases. These methodscan be classified as follows:

1. Titrimetry, 2. spectroscopy (photometry, fluorimetry, in-fra red), 3. chromatography, 4. radioactivity, 5. interfacialtensiometry, 6. turbidimetry, 7. conductimetry, 8. immuno-chemistry, 9. microscopy.

It should be kept in mind, however, that the generaltriacylglycerol hydrolysis reaction catalysed by lipasescan be written as follows:

TAG = triacylglycerols, DAG = diacylglycerols, MAG =monoacylglycerols, FFA = free fatty acids.

As with all reactions catalysed by enzymes, activity mea-surements can be carried out using various physico-chemical methods (by monitoring the disappearance ofthe substrate or the release of the product). Immunologi-cal methods are widely used to quantitate the presence oflipases in biological media, independently from theirlipolytic activity. These two groups of methods will be de-scribed and discussed here. It is worth noting that lipasescan also be quantitated by means of an active site titrationmethod. For instance, Rotticci et al. [43] and Scholze etal. [44] developed a method which is based on the irre-versible inhibition exerted by para-nitrophenyl phospho-nate derivatives. However, since alkylphosphonates arenot specific lipase inhibitors, this method should be re-stricted to the case of purified lipases.

TAG DAG MAG Glycerol

FFA FFA FFA

134 Beisson et al. Eur. J. Lipid Sci. Technol. 2000, 133–153

2 Physico-chemical methods

2.1 Disappearance of substrate

2.1.1 Nephelometry and turbidimetry

In solid media: This method, which is comparable to thoseinvolving the use of coloured indicators in agar plates to-gether with carboxylic esters as the lipase substrate (seechapter 2.2.1.1), consists basically of determining the di-ameter of the product diffusion area. The occurrence oflipolysis will give rise to a clarified zone on the agar plate[45]. This optical phenomenon is observable only if the re-leased fatty acids are partly water-soluble. This clarifica-tion process results from the decrease in the size of thehydrolysed emulsified particles, which in turn causes adecrease in the diffused light. This type of technique canalso be applied to monitor the hydrolysis of Tweens bysome lipases [46]. The chemical structure of Tween es-ters is such, however, that they can be hydrolysed not on-ly by lipases, but also by non-specific esterases.

In liquid media: The method consists of monitoring the de-crease with time in the absorbance of a TAG emulsion[47–51]. This technique is sensitive to artefacts, however,since numerous factors present in blood serum for in-stance can interfere with the process [52, 53]. In addition,the results are not absolute lipase activity values, and toobtain reliable quantitative data, it is therefore necessaryto perform a calibration curve using a pure lipase solution.

This calibration step had hampered a wide diffusion ofthis sensitive method.

A turbidimetric esterase assay was developed using aTween 20 solution in the presence of CaCl2 and Lysobac-ter enzymogenes esterase as enzyme source [54]. Thereaction was monitored by measuring the increase in theoptical density occurring at 500 nm due to the hydrolyticrelease of the fatty acids from Tween 20 and their precip-itation in the form of calcium salts. This turbidimetric as-say was used to determine the specific activity of lipasesfrom Chromobacterium viscosum (87 IU · mg–1) and Can-dida cylindracea (0.5 IU · mg–1). It should be pointed outthat Tweens are not specific substrates for lipases, how-ever.

2.1.2 The Wilhelmy plate method2.1.2.1 Pure monomolecular films assubstrates

Among the interfacial tensiometry methods, themonomolecular film technique at the air-water interfacehas been extensively developed and used by our group[55–64] (Fig. 1) as well as in the group of Brockman[65–67].

This technique consists basically of taking a Teflon troughfilled with an aqueous solution. A thin platinum platedipped in the surface of the aqueous phase is attached to

Eur. J. Lipid Sci. Technol. 2000, 133–153 Methods for lipase detection and assay: a critical review 135

Fig. 1. Set up used with the Wilhelmy plate method to measure the catalytic activity of lipases on monomolecular films.A: Zero order trough for measuring lipase activity on medium chain substrates. B: Zero order trough for measuring lipaseactivity on long chain substrates in the presence of β-CD in the subphase. (1): Teflon™ trough, (2): mobile Teflon™ barrier,(3): electromicrobalance, (4): platinum plate, (5): spread monolayer, (6): reaction compartment, (7): reservoir compartment,(8): surface channel, (9): β-cyclodextrin, E: lipase in solution, E*: lipase absorbed to the lipid film.

an electromicrobalance in order to measure the surfacepressure (π), which is directly related to the interfacial ten-sion (γ) by the relationship π= γo – γ, where γo is the refer-ence value of the interfacial tension with a clean air-waterinterface. This equipment is commercially available atseveral companies: KSV (Helsinki, Finland), Krüss GmbH(Hamburg, Germany) and Kibron Inc. (Helsinki, Finland).A monomolecular film of lipids can be spread at the sur-face of the aqueous phase from a lipid solution in avolatile solvent poorly water-soluble, e.g. chloroform. Thelipid solution is applied in the form of small droplets whichare allowed to evaporate. The area occupied by the lipidfilm can be limited and adjusted with a Teflon barrier thatsweeps the surface of the aqueous phase. The surfacepressure can therefore be maintained automatically con-stant, using the displacement of the Teflon™ barrier that iscontrolled and regulated depending on the output fromthe electromicrobalance. If a lipase solution is injected in-to the aqueous phase below the lipid film, the surfacepressure will decrease, due to the solubilisation of thelipolytic reaction products. Since the barrier moves in or-der to maintain the surface pressure constant, the kineticsof the reaction can be monitored by recording the move-ment of the barrier versus time.

With this technique, it is possible to measure and controlsome important interfacial parameters such as the sur-face pressure (the interfacial free energy) and the molec-ular area of the substrate, as well as the surface excess ofthe water-soluble lipases. One prerequisite of the mono-layer technique, however, is that the water insolublemonomolecular film of substrate should generate water-soluble products during the reaction process. This is whysynthetic medium-acyl chain lipids were originally mainlyused as substrates with lipolytic enzymes [66, 68, 69]. Werecently developed an alternative method using a nonsurface-active agent, β-cyclodextrin, dissolved in theaqueous subphase in order to trap the long-chain lipolyticproducts generated by the lipolysis of monomolecularfilms of long chain neutral acylglycerols [61] or phospho-lipids [63].

The monomolecular film method is suitable for studyingenzymatic reactions on lipid films spread at the air/waterinterface. This method is highly sensitive, and very lowlipid amounts are required to perform reliable kinetic mea-surements. However, in terms of the amounts of lipaseused, the monomolecular film technique requires asmuch lipase as the pH-stat method (from 0.1 to 1 µg perassay).

Nevertheless, the question remains to be answeredwhether the behaviour of a lipid film at the air-water inter-face is actually representative of what is occurring eitherat an oil-water interface or in a complex biomembrane.

2.1.2.2 Mixed monomolecular films assubstrates

Most studies on the kinetics of lipolytic enzyme activitieshave been carried out in vitro with pure lipids as sub-strates. Virtually all biological interfaces are composed ofcomplex mixtures of lipids and proteins. The monolayertechnique is ideally suited for studying the mode of actionof lipolytic enzymes at interfaces using controlled mix-tures of lipids. Two methods of forming mixed lipid mono-layers exist at the air-water interface: either by spreadinga mixture of water-insoluble lipids from a volatile organicsolvent, or by injecting a micellar detergent solution intothe aqueous subphase covered with preformed insolublelipid monolayers.

A new application of the “zero-order” trough was pro-posed by Piéroni and Verger [70] for studying the hydroly-sis of mixed monomolecular films at a constant surfacedensity and a constant lipid composition as shownschematically in Fig. 1A. A Teflon™ barrier was placedtransversally over the small channel of the “zero-order”trough in order to block the surface communication be-tween the reservoir and the reaction compartment. Thesurface pressure was first determined by placing the plat-inum plate in the reaction compartment, where the mixedfilm was spread at the required pressure. The surfacepressure was then measured after switching the platinumplate to the reservoir compartment, where the pure sub-strate film was subsequently spread. The surface pres-sure of the reservoir was equalised with that of the reac-tion compartment by moving the mobile barrier. The barri-er between the two compartments was then removed inorder to allow the surfaces to communicate, and the en-zyme was injected into the reaction compartment and thekinetics were recorded as described [55].

2.1.2.3 Interfacial binding and film recovery

Using the monomolecular film technique, several investi-gators have reported that an optimum occurs in thevelocity-surface pressure profile. The exact value of theoptimum varies considerably with the particular en-zyme/substrate combination used. Qualitative interpreta-tions have been given to explain this phenomenon. Thefirst hypothesis, which was proposed by Hughes [71] andsupported by later workers [72, 73], was that the packing-dependent orientation of the substrate may be one of thefactors on which the regulation of lipolysis depends. Us-ing radiolabelled enzymes, Verger et al. [56] and Pattus etal. [74] subsequently established that the maxima ob-served in the velocity-surface pressure profile disappearwhen they are correlated with the interfacial excess of en-zyme. Indeed, the main difference between the monolay-er and the bulk system lies in the interfacial area to vol-ume ratios, which differ from each other by several orders


of magnitude. In the monolayer system, this ratio is usu-ally about 1 cm–1, depending upon the depth of thetrough, whereas in the bulk system, it can be as high as105 cm–1, depending upon the amount of lipid used andthe state of lipid dispersion. Consequently, under bulkconditions, the adsorption of nearly all the enzyme occursat the interface, whereas with a monolayer, only one en-zyme molecule out of hundred may be at the interface[69]. Owing to this situation, a small but unknown amountof enzyme, which is responsible for the observed hydroly-sis rate, is adsorbed on the monolayer. In order to cir-cumvent this limitation, different methods were proposedfor recovering and measuring the quantity of enzymes ad-sorbed at the interface [62, 67, 75–77].

After performing velocity measurements, Momsen andBrockman [77] transferred the monolayer to a piece of hy-drophobic paper and the adsorbed enzyme was then as-sayed titrimetrically. After correcting for blank rate andsubphase carry-over, the amount of adsorbed enzymewere calculated from the net velocity and the specific en-zyme activity.

In assays performed with radioactive enzymes [75], thefilm was aspirated by inserting the end of a bent glasscapillary into the liquid meniscus emerging above theridge of the Teflon™ compartment walls. As radiolabelledenzyme molecules dissolved in the subphase were un-avoidably aspirated with the film constituents, the resultshad to be corrected by counting the radioactivity in thesame volume of aspirated subphase. The difference be-tween the two values, which actually reflected the excessradioactivity existing at the interface, was attributed to theenzyme molecules having bound to the film. Since it ispossible with the monolayer technique to measure the re-action rate expressed in µmol · cm–2 · min–1 and the inter-facial excess of enzyme in mg · cm–2, it is easy to obtainan enzymatic specific activity value, which can be ex-pressed as usual in µmol · min–1 · mg–1 (IU · mg–1).

By combining a sandwich ELISA technique with themonomolecular film technique, it was possible to mea-sure the enzymatic activity of human gastric lipase (HGL)on 1,2-didecanoyl-sn-glycerol (dicaprin) monolayers aswell as to determine the corresponding interfacial excessof the enzyme [62]. The HGL turnover number increasedsteadily with the lipid packing. The specific activities de-termined on dicaprin films spread at 35 mN · m–1 werefound to be in the same range of the values measured un-der optimal bulk assay conditions, using tributyrin emul-sion as the substrate (i.e. 1000 IU per mg of enzyme). Ata given lipase concentration in the water subphase, theinterfacial binding of HGL to the non hydrolysable eggyolk phosphatidylcholine monolayers was found to be tentimes lower than in the case of dicaprin monolayers [62].

However, we have to keep in mind that the surface-boundenzyme molecules include not only those enzyme mole-cules which are directly involved in the catalysis, but alsoan unknown amount of protein which is present close (ad-sorbed) to the monolayer. These enzyme molecules werenot necessarily involved in the enzymatic hydrolysis of thefilm.

2.1.3 Atomic force microscopy

In order to monitor the kinetics of the hydrolysis of phos-pholipid bilayers by phospholipase A2, Nielsen et al. [78]have performed experiments using atomic force mi-croscopy (AFM) in a liquid medium. Following these stud-ies, the enzymatic hydrolysis of mixed bilayers of acyl-glycerols/phospholipids by Humicola lanuginosa lipase(HLL), was also investigated using AFM. Mica supportedlipid bilayers are hydrolysed by HLL, and as the productsdissolve in the buffer, regions of the bilayer with deep de-fects were detected by the AFM tip. Real time images ofthe hydrolysis occurring in the bilayer were thus obtainedand analysed using purpose built software. These imagesshowed the occurrence of increasingly large nanoscaleindentations at the interface. In addition, the increase inthe area of the holes in the lipid bilayer was recorded as afunction of time and the specific activity of the enzymewas thus estimated, assuming one molecule of lipase tobe acting in each hole. The results were used to model


Fig. 2. Principle of the pH-stat method. The enzyme is in-jected into the thermostated reaction vessel containingthe emulsified substrate. The lipase activity is measuredby recording the amount of titrant (NaOH) added to main-tain the pH at a constant endpoint value during the reac-tion.

the kinetics of the enzyme reaction at the lipid-water in-terface. These data provide the first nanoscale picture ofthe kinetics of lipid degradation by lipases [79].

2.1.4 Infrared spectroscopyA continuous assay for measuring lipase-catalysed hy-drolysis of TAGs in reverse micelles using Fourier trans-form infrared spectroscopy (FTIR), was developed byWalde and Luisi [80]. Lipolysis can be monitored byrecording the FTIR spectrum of the entire reaction mix-ture. Fatty acid esters and FFAs (peak maximum at1751 cm–1 and at 1715 cm–1, respectively) can be quanti-tated on the basis of their molar extinction coefficientsand Beer’s law. This method was applied to measuringthe lipolysis of various substrates (trioctanoylglycerol,vegetable oils).

2.2 Appearance of hydrolytic reactionproducts

2.2.1 Proton release as an indirect assay

2.2.1.1 Coloured indicators

In solid media: When a lipase solution is placed on anagar plate containing a carboxylic ester as substrate, it ispossible to monitor the drop in pH due to the fatty acidsreleased by observing the change in colour of indicatorspreviously incorporated together with the substrate intothe agar gel [22, 45]. There exists a linear relationship be-tween the diameter of the fatty acid diffusion spot and thelogarithm of the enzyme concentration. This technique isvery convenient for rapidly screening lipolytic microorgan-isms growing on agar plates. However, some positivefalse can result from the acidification of the medium, dueto the generation of acidic metabolites other than FFAs,which are released by microbial lipases.

In liquid media: Although it is a qualitative method, thistechnique provides a simple means of detecting lipaseactivity in chromatography column fractions at various li-pase purification stages by observing the changes incolour of indicators mixed with the ester substrates [81].

An even simpler qualitative method is that based on de-tecting the characteristic strong butyric and caproic acidsmell of lipolysed milk droplets used as a lipase substrate.

2.2.1.2 TitrimetryThe well-known pH-stat method (Fig. 2) is generally usedas a reference lipase assay [82, 83]. This is a convenienttechnique for characterising lipase activity and specificity,as well the interfacial activation phenomenon [11, 84, 85].

As shown in Fig. 2 with the pH-stat method, lipase activityis measured on a mechanically stirred emulsion of natur-al or synthetic TAGs by neutralising the FFAs released

with time by adding titrated NaOH in order to maintain thepH at a constant end point value. The pH-stat equipmentis commercially available at Radiometer (Copenhagen,Denmark) or Metrohm Ltd. (Herisau, Switzerland).

In the particular case of the pancreatic lipase/colipasecomplex, the most common routine assay is that per-formed in a thermostated (37 °C) reaction vessel contain-ing 0.5 ml of tributyrin present in 15 ml of a buffered (pH 8)aqueous solution, in the absence of any surfactant oremulsifier other than bile salts [86, 87].

Usually, tributyrin is simply emulsified in situ by the effi-cient mechanical stirring in the pH-stat vessel. The effectof the periodic sonication in the pH-stat method has beenreported, however. Sonication of olive oil emulsified withgum arabic has been shown to improve the sensitivity ofthe assay [88]. Periodic sonication has also been used topromote efficient emulsification of olive oil in the absenceof any surfactant in order to improve the reproducibility ofthe assay [89]. However, sonication procedures are diffi-cult to reproduce experimentally and chemical degrada-tion of the substrate cannot be excluded.

The pH-stat method was used to detect a plant lipase ac-tivity in seedling rape homogenates, in the presence ofdeoxycholate, using triolein emulsified with gum arabic asthe substrate [90]. The pH-stat technique has also beenused to determine lipase activites in serum, plasma andduodenal juice [91, 92]. The pH-stat method is a quantita-tive method which is sensitive to within one µmole of re-leased fatty acid per min. When 0.1 M NaOH is used astitrant, it is not a reliable means of detecting activity levelslower than 0.1 µmole per min. Apart from its low sensitivi-ty, the main disadvantage encountered with the pH-statmethod is the restricted range of pH values which can beinvestigated. More specifically, the detection of the pro-tons released during the hydrolytic reaction catalysed bylipases requires a partial ionisation of the released fattyacids. The end point values of the pH of the reactionmedium must therefore be roughly equal to, or preferablyhigher than the apparent pKa value of the released fattyacid. In fact, it has been reported by Benzonana andDesnuelle that the apparent pKa values of oleic acid couldbe as high as 7.5 under lipase assay conditions [93]. Fur-thermore, it is worth noticing that the ionic strength as wellas the presence of calcium ions will tremendously de-crease the measured values of the apparent pKa. It isgenerally believed that the positive effect of the presenceof calcium ions on lipase assays is due to the water insol-ubility of the calcium soaps of long-chain fatty acids. Thismicro-precipitation phenomenon drives the chemicalequilibrium by virtue of the mass action law.

If at the selected pH value, the FFAs are not fully ionised,their continuous titration is either very inaccurate or im-


possible to perform, even after introducing a correctionfactor. A back-titration has to be carried out, as in the caseof HGL, which is active under acidic pH conditions. TheHGL activity is therefore measured by incubating the en-zyme solution with a TAG emulsion for several min atpH 3. The released non ionised FFAs are back-titrated atthe end of the reaction by inducing a rapid shift of the pHfrom 3 to 9. In a subsequent control experiment (withoutany HGL), the amount of added titrant (NaOH) is thensubstracted from the assay [94, 95].

2.2.2 Analysis of the free fatty acids released

2.2.2.1 Glycerol derived carboxylic esters assubstrates

2.2.2.1.1 Colourimetric assays

A continuous spectrophotometric assay based on themetachromatic properties of the cationic dye safraninehas been described [96]. In this assay, the change in thenet negative charge at the lipid /water interface during thelipolytic reaction is monitored by the absorbance changeof safranine. Lipase activities as low as 50 mU could bedetected using an olive oil emulsion as substrate. Thesensitivity of this method is therefore about two fold high-er than the pH-stat method.

A colourimetric method, based on the formation in an or-ganic phase of a copper soap of the fatty acid in the pres-ence of a dye indicator (copper reagent), was first devel-oped by Duncombe [97]. The copper complex was subse-quently estimated spectrophotometrically at 440 nm. Thesensitivity of the copper reagent and the efficacy of thefatty acid solvent extraction step have been improved bymany workers for specific purposes [98, 99]. Nixon andChan [100] described a procedure which resulted in areference curve showing linearity between 10 and130 nmole of FFAs. In addition, the presence of bovineserum albumin (BSA) and phospholipids was found not tointerfere with the assay.

In the above-described methods using copper soaps,TAGs have to be freed from endogeneous FFAs in orderto obtain an acceptable background level.

The use of rhodamine 6G to obtain a FFA complex whichis extractable in hexane was described by Hirayama andMatsuda [101]. In the presence of fatty acids, a pinkcolour develops and its absorbance is read at 513 nm.The reference curve is linear between 20 and 200 nmol ofFFAs [102]. However, the reproducibility of rhodamine 6Gbatches is difficult to control and a reference curve has tobe performed with each new rhodamine 6G solution. Re-placing hexane by heptane improved the shelf life of therhodamine solution without entailing any loss of sensitivi-ty (Dr. J. Nari, LLE-Marseille, personal communication).

Rogel et al. [103] described a spectrophotometric lipaseactivity assay based on the displacement of parinaric acidpreviously bound reversibly to BSA, which is induced bythe oleic acid released from triolein. The lipase activity ismonitored by recording the changes in the ratio betweenthe absorbances at 319 and 329 nm. This sensitivemethod is limited, however, by the fact that detergentsand calcium ions interfere with the assay.

A plate assay to detect bacterial lipase in a medium con-taining trioleoylglycerol and the fluorescent dye rho-damine B has been described [104]. Substrate hydrolysiscauses the formation of orange fluorescent halos aroundbacterial colonies visible upon UV irradiation. The loga-rithm of lipase activity from cell-free culture supernatantsis linearly correlated with the diameter of halos, therebyallowing quantitation of lipase activities ranging from 60 to1800 mIU.

2.2.2.1.2 Fluorimetric assays

Wilton [105] described an assay which involves the dis-placement of the highly fluorescent fatty acid probe, 11-(dansylamino)undecanoic acid, in rat liver fatty acid-bind-ing protein, which is induced by the long-chain FFAs re-leased by lipases. Quantities as low as 20 pg of purifiedpig pancreatic lipase (PPL) could be detected with thismethod. However, since this highly sensitive assay isbased on the same principle as that described above us-ing BSA/parinaric acid [103], it is likely to be limited by thesame factors. Moreover, the method may be not suffi-ciently reliable when used in the case of crude biologicalmedia containing membrane fractions or albumin.

A commercially available kit (ADIFAB™, ICN pharmaceu-ticals Inc.) which can be used to quantitate FFAs was de-veloped, based on the properties of a 15 kDa mammalianintestinal fatty acid-binding protein conjugated to an acry-lodan fluorophore. Without FFAs, the indicator emits fluo-rescence at 432 nm and 505 nm when binding occurs withFFAs. As mentioned by the manufacturer, the detection ofthe FFA binding is based on a movement of the acrylodanfluorophore which occurs relative to the nonpolar bindingpocket of the protein. According to the manufacturer, thismethod can be used to detect concentrations of FFA aslow as 1 nM.

A method based on interactions between rhodamine Band the released FFAs has been described [106]. The re-action takes place in a gel containing a mixture of rho-damine B, TAGs and agarose deposited in microtiterplatewells. After adding the lipase solution, the released FFAscan be monitored by reading the fluorescence intensity(excitation 485 nm, emission 535 nm) every 10 min for 1to 2 h. The amounts of released FFAs can be estimated


using the calibration curve obtained upon adding variableamounts of FFAs to the gel.

Wolf et al. [107] used phosphatidylcholine containing pari-naric acid [9 (cis), 11 (trans), 13 (trans), 15 (cis)-octade-catetraenoic acid], a naturally fluorescent fatty acid bear-ing four conjugated double bonds, to monitor phospholi-pase A2 activity. Upon hydrolysis, the binding to albuminof the free parinaric acid released was correlated with anincrease in the fluorescence polarisation and in the totalfluorescence intensity. This method prompted us to set upa lipase assay using parinaric acid-containing TAG puri-fied from Parinari glaberrimum seed oil [108]. The purifiedTAGs are naturally fluorescent and more than half of thefatty acids from Parinari oil are known to contain parinaricacid in its esterified form [109]. Under the assay condi-tions used, the excitation and emission wavelengths ofParinari oil were 324 nm and 420 nm, respectively.

The presence of detergents (sodium taurodeoxycholate,CHAPS, Sulfobetaine SB12, Tween® 20, Brij® 35,Dobanol®, n-Dodecylglucoside) above their critical micel-lar concentration dramatically increases the fluorescenceof the free parinaric acid released by various lipases. Thisincrease in the fluorescence intensity is linear with timeand proportional to the amount of lipase added. This newmethod, performed under non-oxidative conditions, wasapplied successfully to detecting low lipase levels incrude protein extracts from plant seedlings and could bescaled down to microtiterplate measurements. Quantitiesas low as 0.1 ng of pure pancreatic lipase could be de-tected under standard conditions (pH 8). Lipase activitycan also be assayed in acidic media (pH 5) HGL [108].One drawback of this method, however, is the susceptibil-ity of parinaric acid to oxidation by atmospheric oxygen.This drawback can be overcome by adding antioxidantagents to the buffers and by performing incubation stepsunder an argon or a nitrogen atmosphere. Furthermore,this method requires the presence of a selected detergentin order to solubilise the released parinaric acid intomixed micelles; and of course, this detergent must not in-hibit lipase activity. We were not able to use this methodto measure lipase activity in the stratum corneum of hu-man skin (F. Beisson, Anal. Biochem., submitted), due tothe highly fluorescent background of the adhesive tapestrips used to collect the stratum corneum. This simpleand continuous assay is compatible, however, with a highsample throughput and might be applicable to detectingtrue lipase activities in various biological samples as wellas in directed evolution experiments.

2.2.2.1.3 Chromatographic assays

Various chromatographic techniques can be used to de-tect lipids as well as FFAs released from TAGs, namelyflorisil columns or silicic acid columns, thin-layer chro-

matography or gas-liquid chromatography [110–113]. Thelatter method requires that fatty acids are previouslytransformed into their methyl esters, however, so as torender them volatile.

After thin-layer chromatography, a quantitative analysis ofthe released FFAs can be carried out using either densi-tometric or autoradiographic methods together with radio-labelled TAGs. The specificity and the sensitivity of thesetechniques are very satisfactory, since they can be usedto detect fatty acid in quantities as small as a few pmoles[114]. However, these methods are rather time-consum-ing and they are not continuous.

A high-performance liquid chromatographic (HPLC) as-say for determining lipase activity was developed by Mau-rich et al. [115] using β-naphtyllaurate as substrate. Thespecific activity of PPL was only 1.5 IU · mg–1, which isvery low in comparison with the PPL activity measuredwith other methods and other substrates (around 10 000IU · mg–1, using a tributyrin emulsion).

With a view to determining PPL activity, Maurich et al. de-veloped a highly sensitive HPLC method, using thepalmitic and lauric esters of para-nitrophenol as sub-strates. These authors gave details as to the specific ac-tivity and reproducibility of lauric ester alone, becausepalmitic ester turned out to be a very poor substrate forPPL [116].

2.2.2.1.4 Enzymatic assays

The light emitted by some reaction products of the lu-ciferase from the marine luminous bacterium Beneckeaharveyi in the presence of fatty acids can be utilised to de-tect levels as low as 1 pmole of myristic acid and 100 to200 pmoles of palmitic or oleic acid [117]. One main draw-back of this technique is the fact that oleic, linoleic, palmi-toleic and linolenic acids inhibit the luciferase. Moreover,at pH 8, a 5-fold decrease in the luminescence is ob-served in comparison with that obtained at pH 6.5.

Due to the action of some lipoxygenases, linoleic acid, inthe presence of oxygen, generates a hydroperoxide de-rivative which can subsequently be revealed with thio-cyanate as a red complex. The formation of this complexcan either be quantatively monitored [118] or the oxygenconsumption during the reaction catalysed by the lipoxy-genase can be monitored by means of an oxygen elec-trode [119]. Some lipoxygenases do not act specificallyon free linoleate leading to overestimation of the extent ofhydrolysis. Furthermore, non-enzymatic co-oxidation maylead to erroneous results.

Acyl-CoA synthetase can be used to catalyse the for-mation of acyl-CoA from Coenzyme A and FFAs. Usingacyl CoA oxidase, acyl-CoA can then be converted into


enoyl CoA and hydrogen peroxide, which in turn is con-verted into red quinone dye by peroxidase in the pres-ence of phenol and 4-aminoantipyrine. The reaction canbe monitored by measuring the colour produced at 500nm [120]. The great drawback of this method is that manyof the enzymes present in a crude biological medium arelikely to interfere with the enzymatic reactions during theassay.

2.2.2.1.5 In situ detection by electronmicroscopyFatty acids released in animal tissues by lipases can bedetected by electron microscopy [121]. This techniquehas been used for example to detect rat lingual lipase[122] and the lipase present in the pancreatic acinar cellsof mice and rats [123, 124]. It has also been used to de-tect lipase activities in the outer epidermis: aldehyde-fixedtissues (100-µm slices) were incubated with triolein and

then exposed to lead salts to form insoluble soaps, and fi-nally processed for electron microscopy. Although thelead precipitates were difficult to detect, some sites of li-pase activity were identified in the cells. Larger precipi-tates have been obtained by using Tweens instead of tri-olein, but the former are not specific lipase substrates[125, 126].

2.2.2.2 Synthetic carboxylic esters assubstrates

2.2.2.2.1 Radioactive assays

The methods involving the use of TAGs containing radio-labelled acyl chains [127, 128] are specific and very sen-sitive lipase assays. However, they cannot be monitoredcontinuously and they need a time-consuming chromato-graphic step or an organic solvent fractionation step toisolate the released fatty acids.


Fig. 3. Principle of the resorufin ester assay.The absorption maximum of the released re-sorufin is at 572 nm at pH 6.8 and 583 nm in thealkaline range. In the case of incubations per-formed at pH < 6.8, the pH is adjusted to 6.8 byadding KOH or alcaline buffer before readingthe absorbance at 572 nm.

A method based on the binding of radiolabelled 63Ni toFFAs extracted in organic solvents has been described[129]. The estimated quantity of 63Ni bound to oleic acidwas found to be linear up to 100 nmol of fatty acid, andquantities as small as 1 nmol of FFA can be detected perassay. However, this method requires a time-consumingstep, in which the heptane phase containing the FFAs hasto be evaporated prior to adding the radiolabelled nickelsolution. It also requires the use of completely fatty acidfree glassware.

2.2.2.2.2 Colourimetric assays

When using as a lipase substrate a synthetic TAG con-taining fatty acyl chain labelled by a pyrene group in ωposition, it is possible to determine the amount of labelledFFAs released, using spectrophotometric methods with asensitivity of around one nmole. This technique is notstraightforward to use, however, since it requires perform-ing solvent extraction of lipid molecules and separatingthe labelled TAG from the released labelled products,both of which having a yellow colour [130].

Resorufin ester (1,2-O-dilauryl-rac-glycero-3-glutaricacid-resorufin ester) is a sensitive “glycerol-derived” sub-strate (Fig. 3), which is commercially available at Hoff-mann-La Roche (previously Boehringer Mannheim), andcan be used conveniently for lipase assays under specif-ic experimental conditions in the presence of a nonionic

detergent named Thesit™. This assay is widely used forthe determination of lipase activity in serum and was alsoutilised in the case of a microbial [131] and a plant lipase[132]. However, we established at our laboratory that thehydrolytic activity measured in a plant homogenate withthis resorufin ester was not in fact attributable to a true li-pase activity. The catalytic activity on resorufin ester wasnot abolished by heating the extract for 5 min at 95 °C,whereas the true lipase activity measured with radiola-belled triolein or TAGs containing parinaric acid (seechapter 2.2.2.1.2) decreased dramatically after heat treat-ment [108]. In addition, at our laboratory we have used re-sorufin ester, in absence of detergent (e.g. Thesit™), inorder to determine and compare the rates of hydrolysis ofresorufin ester and tributyrin by various pure proteins(Tab. 1). Unlike tributyrin, resorufin ester was poorly hy-drolysed by many lipases. Some lipases such as Candidarugosa, Pseudomonas glumae, Rhizomucor miehei lipas-es and human pancreatic lipase (HPL) can significantlyhydrolyse resorufin ester, showing specific activities rang-ing from 200 to 1000 IU · mg–1. However, if one looks atthe ratio between rates of hydrolysis of tributyrin and re-sorufin ester, it is striking that this ratio ranges from 5 to1000 in the case of Candida rugosa and Candida antarc-tica, respectively. A low but significant resorufin ester hy-drolysis rate was obtained with hemoglobin purified frompig liver. The data obtained using resorufin ester to mea-


Tab. 1. Initial rates of hydrolyis of tributyrin emulsions and resorufin ester by various pure proteins. Tributyrin assay condi-tions: with all the proteins assayed except RGL, the buffer used was 1 mM Tris-HCl (pH 8), 150 mM NaCl, 10 mM CaCl2.With RGL, the buffer used was 50 mM acetate (pH 6), 150 mM NaCl, 2 mM NaTDC, 1.5 µM BSA. Resorufin assay condi-tions (in the absence of Thesit™): 10 µl of a lipase sample were added to 90 µl of KH2PO4 0.1 M (pH 6.8) and 7 µl ofresorufin ester stock solution in dioxane (1 mg · ml–1). With RGL, the buffer used was 20 mM KH2PO4 (pH 6.8), 150 mMNaCl, 0.05% Triton X100.

Protein Tributyrin Resorufin ester Tributyrin/Resorufin[IU · mg–1) (IU · mg–1) ester ratio

Fungal lipasesCandida antarctica lipase B 184 0.2 1022Candida rugosa lipase 1037 214 4.8Fusarium solani cutinase 3180 32 99Pseudomonas glumae lipase 3000 401 7.5Rhizomucor miehei lipase 8240 450 18.3

Mammalian lipasesHuman pancreatic lipase + Colipase 8000 1000 9Lipoprotein lipase 250 8 31.2Rabbit gastric lipase 800 3 267

Non enzymatic proteinsHemoglobin 0 0.7 0Bovine serum albumin 0 0 –

sure lipase activities should therefore be interpreted withgreat care.

2.2.2.2.3 Fluorimetric assays

The main advantages of fluorimetric assays is their sensi-tivity and the fact that they make it possible to continu-ously monitor the reaction kinetics. We will therefore re-view here only the continuous fluorescent assay de-scribed in the literature.

Dansyl-phosphatidylethanolamine [133] and NBD-phos-pholipids [134] have been used as a substrate to detectthe phospholipase activity of lipoprotein lipase. Thesefluorescent phospholipid molecules are not, however,typical and specific substrates for other lipases.

Another important group of fluorogenic substrates is thatconsisting of pyrenic acylglycerol derivatives, which wereoriginally used for phospholipase determination [135,136]. Derivatives of this kind were used in lipase activitydeterminations in which the production of FFAs which oc-curs upon lipolysis causes a shift in the peak fluorescenceintensity. The lipolytic activity can be quantitated in termsof the increase with time in the fluorescence intensity at agiven wavelength. The use of a TAG containing fatty acylchain in which a pyrene residue is linked to the ω positionwas first described by Nègre et al. [137]. However, thissensitive assay requires the pyrene-labelled FFA re-leased to be isolated from the reaction medium. Aquencher residue (trinitrophenylamine residue) was intro-duced by Duque et al. [138] as a means of decreasing thebasal fluorescence of the TAG containing acyl-pyrene (1-O-hexadecyl-2-pyrenedecanoyl-3-trinitrophenylaminodo-decanoyl-sn-glycerol and its enantiomer). This intramolec-ularly-quenched TAG containing acyl-pyrene can there-fore be used in a continuous fluorescent assay [139]. Un-fortunately, as can be seen from the data reported byDuque et al. [138], this kind of chemically-modified TAG ispoorly hydrolysed by lipases, probably for steric reasons.We have confirmed that a poor rate of hydrolysis by hu-man pancreatic lipase occurs with this kind of labelledTAG: 5 · 10–3 IU per mg, as compared with 3000 IU permg on natural long chain TAG [108].

The overall sensitivity of a fluorescent assay using syn-thetic TAGs will obviously depend on two factors: first, onthe sensitivity of the FFA detection, i.e. on the lowestamount of fluorescent released FFAs that can be detect-ed, and secondly, on the specific activity of the lipase. Thesensitivity of the quenched pyrene-TAG based assay istherefore offset by the very poor rate of lipase hydrolysisof the corresponding synthetic TAG, as compared withthat of non chemically-modified TAGs (such as TAGs con-taining parinaric acid, see above, chapter 2.2.2.1.2).

2.2.3 Determination of the released alcohol orthiol2.2.3.1 Determination of the released glycerolfrom TAG

Direct determination of free glycerol is not commonly per-formed, since all three acyl chains of a triacylglycerol mol-ecule are rarely released by a single lipase, and thereforethe initial hydrolysis rates cannot be determined. Threetechniques have been described, however, in the litera-ture for estimating the released free glycerol:

● Periodic oxidation of free glycerol, which leads to theformation of formaldehyde that can then be assayedspectroscopically.

● Bioluminescence using the luciferin-luciferase com-plex [140].

● Phosphorylation of glycerol into glycerol-3-phosphatefollowed by conversion of the latter product into dihy-droxyacetone-phosphate, which leads to the produc-tion of hydrogen peroxide, which can be determinedspectrophotometrically at 550 nm using peroxidase(Kit lipase-PS™ of Sigma) [141].

2.2.3.2 Determination of the released colouredor fluorescent alcohol from syntheticcarboxylic esters

The hydrolysis of carboxylic esters of α-naphthol, para-ni-trophenol or 2,4-dinitrophenol leads to the release of al-cohols that can be monitored continuously and quantita-tively using a spectrophotometric method. The appear-ance of the yellow coloured para-nitrophenol (pKa 7.15)can be monitored by reading the absorbance at 405 nm[142–145]. The formation of 2,4-dinitrophenol (pKa 3.96)can be monitored from the increase in the absorbance at360 nm [146]. Since the molar extinction coefficient of aphenolic solution is generally very dependent on its ioni-sation state, dinitrophenyl esters are more convenientsubstrates than para-nitrophenyl esters when the pH ofthe assay is around 7–8. The red colour obtained with α-naphthol can be monitored after complexing the solutionwith a diazonium salt [147]. The use of acyl esters of 5-(4-hydroxy-3,5-dimethoxyphenylmethylene)-2-thioxothiazo-line-3,5-dimethoxyphenylmethylene)-2-thioxothiazoline-3-acetic acid as chromogenic lipase substrates has alsobeen described [148]. Upon hydrolysis, these substratesyield an intensely red colour which can be assayed at 505nm. However, esters of this kind do not in fact seem to bespecific substrates for determining true lipase activities.

Berg et al. [149] have reported a colourimetric assay ofHLL using para-nitrophenyl butyrate, which partitioned atthe interface of small anionic unilamelar vesicles (SUVs)with a mean diameter of around 40 nm, such as palmi-toleoyl-oleoyl-phosphatidylglycerol (POPG)-SUVs. The


hydrolysis rate measured was 100-fold higher with(POPG)-SUVs than with the monodispersed substrate orwith the substrate partitioned into zwitterionic palmi-toleoyl-oleoyl-phosphatidylcholine (POPC)-SUVs. The li-pase acivity can then be measured using colourimetricmethods.

According to the authors, POPG-SUVs fulfil the criteria fora neutral diluent, since they provide an interface for thepartitioning of both substrate and lipase and do not blockthe lipase active-site. In this well defined system, the pri-mary rates and equilibrium constants for the interfacialcatalysis by HLL have been established [149].

In our opinion, nitrophenol acyl esters should not be usedas lipase substrates for several reasons:

i. They are not at all specific lipase substrates, sincethey can be hydrolysed by non-specific esterases andproteases often present in biological samples. More-over, BSA or the non catalytic C-terminal domain ofPPL can hydrolyse para-nitrophenyl acetate at thesame rate as purified PPL [150].

ii. The catalytic turn-over number of true lipases on para-nitrophenol acyl esters is usually several orders ofmagnitude lower than that obtained with TAG. Oneshould recall that phenol esters are esters of sec-ondary alcohols, whereas the vast majority of all theknown lipases act exclusively on primary ester bondssuch as those present in the sn-1 and sn-3 positions ofTAG. As an example, the specific activity ratios (tribu-tyrin/para-nitrophenyl acetate) are 2320, 1000 and1430 when using purified PPL, HGL and rabbit gastriclipase, respectively [151].

iii. The carbonyl function of these secondary esters iselectronically activated (it bears a partial positivecharge, due to the electronic delocalisation of the aro-matic ring, which is enhanced by the electron attractiveNO2 substituent). These esters are therefore liable toundergo non-enzymatic alkaline and acidic hydrolysis.If, however, long acyl chain para-nitrophenyl esters aredissolved in an inert micellar detergent, this disadvan-tage no longer applies.

All the available methods of measuring lipase activity us-ing nitrophenyl esters therefore need to be handled withgreat care, even when pure lipases are used. A series ofpara-nitrophenyl esters with variable acyl chains havebeen frequently used, however, as reported in the litera-ture, to quickly assay the so-called chain length specifici-ty of the microbial lipases used in many biotransformationprocesses [152].

The fluorophore of fluorogenic synthetic lipase substratesis sometimes located in the alcohol moiety. This is so inthe case of 4-methylumbelliferone acyl esters, which

were first described by Jacks and Kircher [153]. Umbelli-ferone acyl esters have been found to be more sensitiveand stable [154]. The use of methylumbelliferone estersas substrates to assay the lipase activities in human epi-dermal stratum corneum has been investigated at our lab-oratory. Methylumbelliferone oleate is likely to be a spe-cific substrate for these epidermal lipase activities undergiven assay conditions, whereas methylumbelliferoneheptanoate is not a specific substrate at all (Beisson etal., Anal. Biochem., submitted).

Lauroyl pyrenemethanol has been used to assay gastriclipase, cellular lipases of hematopoietic cells and Rhizo-pus arrhizus lipase [155]. The use of monodecanoyl-fluo-rescein has also been tested as a means of assaying li-pases [156].

The carboxylic esters containing fluorogenic secondaryalcohols instead of glycerol are very sensitive and conve-nient substrates, but one should not forget that they arenot, a priori, hydrolysed specifically by lipases. Moreover,they are often prone to high spontaneous hydrolysis.

2.2.3.3 Determination of the released thiolfrom synthetic thioestersThere exist numerous thioesters which can be hydrolysedby lipases. This reaction causes the release of one orseveral sulfhydrile functions, which can subsequently bedetected using the chromogenic Ellman’s reagent (5, 5’dithiobis 2-nitro benzoate). These thioesters are, eitherpoorly or non specifically hydrolysed by lipases, however.Two thioesters (2,3-dimercaptopropan-1-ol tributyroateand 3-mercaptopropan-1,2-diol tributyroate) have beenused for lipase assays in serum samples pretreated withphenylmethylsulfonyl fluoride, a potent inhibitor of someserum esterases. Synthetic thioester TAG analogues canbe synthesised that will allow rapid analysis of the stereo-preference (sn-1/sn-3) of the investigated lipases[157–159].

2.2.4 Electric conductivityThe electric conductivity of the medium increases duringthe lipase hydrolysis reaction due to the development ofelectric charges carried by the released FFAs [160, 161].This technique has numerous drawbacks: the measure-ments are highly temperature-dependent, and the sensi-tivity is really satisfactory only when triacetin is used assubstrate. However, it has turned out that triacetin is not aparticularly suitable lipase substrate [14].

2.2.5 Acoustic wave conductanceA surface acoustic wave sensor system for assaying theactivity of pancreatic lipase has been proposed [162]. Theassay of this enzyme is based on the change in conduc-tance of the solution caused by the release of a fatty acid,


using triolein as a substrate. A linear relationship betweenfrequency response and enzyme concentration is ob-tained.

2.2.6 The oil-drop methodIn 1987, Nury et al. [163] established at our laboratorythat unique information can be gained by measuring thevariations in the oil-water interfacial tension (γo/w) as afunction of time during lipase hydrolysis. These authorsadapted the well-known “hanging-drop method” to studythe rate of lipase hydrolysis of natural long chain TAGs.

The interfacial tension is measured here by automaticallyanalysing the oil drop profile on-line, using the Laplace-Young equation. The accumulation of tensioactive hydrol-ysis products at the surface of an oil drop is responsiblefor the decrease in the interfacial tension, which in turn iscorrelated with changes with time in the drop profile [164](Fig. 4). Fully automated oil drop tensiometers based onthis principle are now commercially available at InterfacialTechnology Concept SARL (France) and have beenfound to have many advantages [165].

As compared to the other interfacial techniques, the oil-drop tensiometer presents the unique advantage of beingable to monitor lipase activities on natural long-chainTAGs at a closely controlled oil-water interface. Further-more, the surface behaviour (interfacial binding) of lipas-es and mutants at the oil-water interface can be studied

under similar conditions [166–168]. The oil-drop method-ology requires the oil to be carefully freed from any natur-al tensioactive compounds such as FFAs and di- andmonoglycerides because of the amphipathic character ofthese contaminants, which might decrease the initial in-terfacial tension. Clean materials and equipment are alsoa strict requirement for the oil-drop methodology to givereliable results, as is also the case with the Wilhelmy platemethod. At supra catalytic concentrations of lipases, theprotein adsorption to the oil droplet could also affect thesurface tension. In this case, hydrolysis kinetics shouldtherefore be interpreted with care.

3 Immunological methods

We have decided not to develop the various spectroscop-ic methods which can be used to detect adsorbed lipaseson model interfaces, such as infrared spectroscopy [169],circular dichroism [170], fluorescence microscopy [171]and ellipsometry [172].

The set of tests which go under the name of ELISA (en-zyme-linked immuno sorbent assay) forms a highly sensi-tive and specific system for detecting and quantifying li-pases [173–175]. These immunological methods can beused to detect both the active and inactive forms of a giv-en lipase. The immunological detection of lipases re-quires purifying the enzyme from a natural or recombinant


Fig. 4. Diagram of the experimental oil drop set-up. (1): Optical bench, (2): integrated sphere light source (halogen lamp),(3): thermostated cuvette, (4): syringe (Exmire type), (5): DC motor with a 500 count per revolution optical encoder,(6): telecentric gaging lens (Melles Griot, Rochester, NY, USA), (7): CCD camera 512 × 512, (8): personal computer,(9): video monitor.

source as well as raising poly- or monoclonal antibodiesusing conventional methods.

This assay essentially consists of first binding a polyclon-al antibody to a solid support (such as a PVC microtiterplate), and then causing the antigen (lipase) to interactwith the first antibody (the captor), before adding a sec-ond antibody (the detector, which is usually a monoclonalantibody) that recognises a different epitope on the anti-gen. The amount of antigen present is quantitated by es-timating the amount of the second antibody labelled witheither biotin [176] or a fluorescent probe [177, 178]. Bio-tinylation of lipases is not always straightforward andshould be carefully checked, e.g. for the level of biotinyla-tion, heterogeneity of protein material as well as for the ef-fect on the overall charge and hydrophobicity. Even whenthe effects of biotinylation on activity are absent, the bind-ing properties may be changed.

3.1 ELISA for pure lipases

Several ELISAs have been specifically developed forquantitating the amounts of pure lipases, e.g. HPL [173,179], HGL [62], lipoprotein lipase (LPL) [180–182], andhepatic lipase [181].

At our laboratory, ELISA tests were developed and usedby Aoubala et al. [62] to evaluate the surface excess ofHGL at a lipid-water interface. This ELISA assay wasadapted to the monomolecular film technique (see Chap-ter 2.1.2.3). HGL was biotinylated without any significantloss of its catalytic activity occurring and was further de-tected (detection limit 25 pg/well) by performing a sand-wich ELISA using anti-HGL polyclonal antibodies as spe-cific captors before revealing the biotin-labelled HGL, us-ing a streptavidin-peroxidase conjugate as a tracer [62].

By combining the above sandwich ELISA technique withthe monomolecular film technique, it was possible for thefirst time to measure the enzymatic activity of HGL on di-caprin monolayers as well as to determine the corre-sponding interfacial excess of the enzyme, and thus tocalculate the specific activity of the lipase at the lipid-wa-ter interface [62].

A similar sandwich ELISA test was adapted at our labora-tory by Labourdenne et al. [167] to determine the amountof HPL adsorbed at an oil/water interface, using the oil-drop technique.

3.2 Lipase immunodetection on physiologicalmedia3.2.1 Lipase immunodetection in plasma orserum

Several immunological assays for determining the pan-creatic lipase concentration in sera have been developed

and used the clinical diagnosis of pancreatitis, where theyare of particular value due to their high specificity andsensitivity [183–187]. Other immunoassays and commer-cial kits have been developed to quantitate immunoreac-tive LPL [188–192] as well as hepatic lipase [188, 193] inhuman serum and carboxyl ester lipase [194]. ELISA us-ing serum antibodies to Staphylococcus aureus lipase isa sensitive assay for serological diagnosis of staphylo-coccal infections [195–197].

3.2.2 Lipase immunodetection in otherphysiological mediaSome groups have developed ELISA for detecting andquantitating LPL in tissues or in cell culture lysates [177,189, 190, 198, 199].

The availability of specific antibodies has led many au-thors to set up sensitive and specific ELISA tests for mea-suring the amounts of lipases in physiological media suchas the duodenal contents, in which both HGL and HPL arepresent [176, 179, 200]. A radioimmunoassay has alsobeen developed for determining the concentration of HPLin the urine [183] and in the duodenal contents [200].

An ELISA test has also been described which can beused to determine pancreatic insufficiency by quantitatingthe HPL levels in the stools [201] as well as providing anindex of severity in patients suffering from cystic fibrosis[202].

3.3 Immunocytolocalisation of lipasesUsing various immunohistochemical techniques, manyauthors have mapped mammalian lipases such as gastriclipase [203], LPL [204–206], hepatic lipase [207], hor-mono-sensitive lipase [208], and microsomal triacylglyc-erol hydrolase [209].

3.4 Immunoblot analysis of lipasesImmunoblot analysis consists of combining gel elec-trophoresis with an immunochemical method of detection.It therefore provides a valuable technique for determiningthe molecular mass of lipases, as well as the specificity ofantibodies for a given lipase. Proteins are separated on asodium dodecyl sulfate polyacrylamide gel, transferred toa supporting membrane, and washed and incubated withantibodies in the presence of blocking reagents that re-duce non-specific binding.

Western blotting analysis using monoclonal antibodieshas been widely used to discriminate between sequentialand conformational epitopes. However, Bezzine et al.have recently shown first that in HPL, the unfoldingprocess might not be complete at 3% w/v SDS, and sec-ondly, that the C-terminal domain of HPL may be de-natured during treatment of the sample at 3% w/v SDS,


and then become refolded within the separation gel at0.1% w/v SDS [210].

4 Conclusions

In order to explore the relationships between the structureof the lipid bilayer and the catalytic activity of HLL, Vissinget al. [211] recently attempted to set up a “universal” li-pase substrate consisting of monomyristoylglycerol(MMG) inserted into unilamellar vesicles of dimyris-toylphosphatidylcholine (DMPC) and dimyristoylphos-phatidylglycerol (DMPG) in the proportion [DMPC/DMPG(70:30)]/[MMG](70:30). However, the HLL activity mea-sured on this “universal” substrate was found to bearound two orders of magnitude lower than that recordedon the same mixture of lipids spread at an air-water inter-face (M. Ivanova, LLE-Marseille, personal communica-tion).

The main conclusion to be drawn from the above reviewis that there exists no single universal method of lipaseassay, but rather a whole range of different techniques.The choice of a particular method will depend on theuser’s own specific requirements.

4.1 Pure lipases or crude biological media

As many non-specific esterases are often present in bio-logical samples, we recommend the use of long-chainacylglycerols as substrates rather than other esters, in or-der to detect and assay a true lipase activity. If long-chainacylglycerols are not used as the substrate, when it is pro-posed to detect lipase activities in a crude sample and/orfor the sake of convenience, great care should be taken ininterpreting the results, since the activity measured mightbe due to enzymes other than lipases. Esters other thanglycerides can be used in experiments involving purifiedlipases. Interfacial methods (monolayer and oil drop ten-siometer techniques) can be used to perform detailedstudies on the effects of various interfacial parameters onlipolysis. It is worth mentioning that monoacylglycerols,diacylglycerols and FFAs should be removed from theTAGs used as substrates in order to avoid detecting sec-ondary activities, and even more importantly, to minimiseany uncontrolled tensioactive effects occurring at the oil-water interface and due to partial glycerides and FFAs. Itshould be mentioned in addition that the choice of a syn-thetic or natural TAGs as the substrate is crucial, espe-cially when dealing with lipases showing a high degree ofselectivity for a given type of fatty acids (typoselectivity),which seems to occur in the case of some plant [18] or mi-crobial [212] lipases.

Since short acyl chains para-nitrophenyl esters can behydrolysed by non enzymatic proteins, they should not beused to assay lipase activities, even with purified lipases.

4.2 Level of lipase activity

The pH-stat method is probably the most suitable methodof measuring lipase activities greater than 0.1 IU. As men-tioned above, however, this technique can be used onlywithin a restricted pH range.

When measuring lipase activities lower than 0.1 IU, theFFAs released have to be extracted and quantitated withmethods involving the use of copper salts or rhodamine. Ifthese methods are not sufficiently sensitive, radiolabelledsubstrates can be used, and the FFAs subsequently ex-tracted. Lastly, fluorescent synthetic esters can be usedto perform sensitive continuous or discontinuous tests,but great care needs to be taken with their substratespecificity.

The method based on the use of a naturally fluorescentTAGs recently described by Beisson et al. provides a sen-sitive, specific and continuous assay. It is therefore a par-ticularly promising method for measuring lipase activitiesin crude biological media [108].

Acknowledgements

We are indebted to our colleagues at the Laboratoire deLipolyse Enzymatique du CNRS, Dr. Alain de Caro, Dr.Vincent Arondel, Dr. Frédéric Carrière and Dr. AbdelkarimAboulsalham as well as Pr. Louis Sarda (Université deProvence) for fruitful discussions. Our thanks are alsodue to our collegues Dr. Thomas Björnholm, Dr. ThomasCallissen, Dr. M. Ivanova, Dr. Liliane Dupuis, Dr. RichardLehner and Isabelle Douchet for giving permission tomention their respective unpublished data, as well as toSylvie Nevy for her contribution to the initial draft of thisreview. The assistance of Dr. Jessica Blanc in revising theEnglish manuscript is acknowledged.

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[Received: December 14, 1999; accepted: January 1, 2000]


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