[4] Monolayer techniques for studying phospholipase kinetics

[4]

P H O S P H O L I P A S E K I N E T I C S I N L I P I D MONOLAYERS

49

[4] M o n o l a y e r T e c h n i q u e s for S t u d y i n g Phospholipase Kinetics By S. RANSAC, H. MOREAU, C. RIVI~.RE, and R. VERGER

Introduction: Why Use Lipid Monolayers as Phospholipase Substrates? There are at least five major reasons for using lipid monolayers as substrates for lipolytic enzymes (readers are referred to previous reviews for details1-4): (1) The monolayer technique is highly sensitive, and very little lipid is needed to obtain kinetic measurements.This advantage can often be decisive in the case of synthetic or rare lipids. Moreover, a new phospholipase A 2 has been discovered using the monolayer technique as an analytical tool. 5 (2) During the course of the reaction, it is possible to monitor one of several physicochemical parameters characteristic of the monolayer film: surface pressure, potential, radioactivity, etc. These variables often give unique information. (3) With this technique the lipid packing of a monomolecular film of sub strate is maintained constant during the course of hydrolysis, and it is therefore possible to obtain accurate, presteady-state kinetics measurements with minimal perturbation caused by increasing amounts of reaction products. (4) Probably most importantly, it is possible with lipid monolayers to vary and modulate the "interfacial quality," which depends on the nature of the lipids forming the monolayer, the orientation and conformation of the molecules, the molecular and charge densities, the water structure, the viscosity, etc. One further advantage of the monolayer technique as compared to bulk methods is that, with the former, it is possible to transfer the film from one aqueous subphase to another. (5) Inhibition of phospholipase activity by waterinsoluble substrate analog can be precisely estimated using a "zero-order" trough and mixed monomolecular films in the absence of any synthetic, nonphysiological detergent. The monolayer technique is therefore suitable for modeling in vivo situations. I R. 2 R. 3 R. 4 G.

Verger and G. H. de Haas, Annu. Rev. Biophys. Bioeng. 5, 77 (1976). Verger, this series, Vol. 64, p. 340. Verger and F. Pattus, Chem. Phys. Lipids 30, 189 (1982). Pi6roni, Y. Gargouri, L. Sarda, and R. Verger, Adv. Colloid Interface Sci. 32, 341

(1990). 5 R. Verger, F. Ferrato, C. M. Mansbach, and G. Pi6roni, Biochemistry 21, 6883 (1982).

METHODS IN ENZYMOLOGY, VOL. 197

Copyright © 1991 by Academic Press, Inc. All fights of reproduclion in any form reserved.

50

PHOSPHOLIPASE ASSAYS,KINETICS,SUBSTRATES

[4]

!

FIG. 1. Methodfor studyingthe hydrolysisof long-chainphospholipidmonolayerswith controlled surfacedensity.A large excess of serumalbuminhas to be present in the aqueous subphase in order to solubilizethe lipolyticproducts.

Pure Lipid Monolayers as Phospholipase Substrates A new field of investigation was opened in 1935 when Hughes 6 used the monolayer technique for the first time to study enzymatic reactions. He observed that the rate of the phospholipase A-catalyzed hydrolysis of a lecithin film, measured in terms of the decrease in surface potential, diminished considerably when the number of lecithin molecules per square centimeter was increased. Since this early study, several laboratories have used the monolayer technique to monitor lipolytic activities, mainly with glycerides and phospholipids as substrates. These studies can be tentatively divided into four groups. Long-chain lipids were used and their surface density (number of molecules/cm2) was not controlled, short-chain lipids were applied, again without controlling the surface density, or shortchain lipids were used at constant surface density. As shown in Fig. 1, C. Rothen (personal communication, 1980) has developed a fourth method at our laboratory for studying the hydrolysis of long-chain phospholipid monolayers by various phospholipases A 2 involving controlled surface density. A large excess of serum albumin has to be present in the aqueous subphase in order to solubilize the lipolytic products. This step was found not to be rate limiting. The linear kinetics obtained by these authors with 6A. Hughes,Biochem. J. 29, 437 (1935).

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

51

natural long-chain phospholipids were quite similar to those previously described in the case of short-chain phospholipids using a zero-order trough with the barostat technique. 3 A shortcoming of the monolayer technique is the denaturation of many enzymes that occurs at lipid-water interfaces. This is attributable to the interfacial free energy. Fortunately, this phenomenon occurs slowly with the highly resistant phospholipases.

Triggering of Monolayer Activity of Phospholipase A2 by Electrical Field or "Vertical Compression" Thuren et al. 7 showed that the action of phospholipase A 2 can be triggered by applying an electric field across a 1,2-didodecanoyl-sn-3phosphoglycerol monolayer lying between an alkylated silicon surface and water. When the silicon wafer served as a cathode, rapid activation of porcine pancreatic phospholipase was observed and was found to depend on the magnitude of the applied potential. The degree of activation differed depending on whether pancreatic phospholipase A2 or snake or bee venom enzymes were used. Maximally, a 7-fold activation of pancreatic phospholipase A2 was observed when the applied potential was 75 V. The effective field over the lipid film was estimated to be approximately 25-175 mV, which is within the range of the membrane potentials found in cells. On the basis of these results, it was suggested that changes in membrane potential might be an important factor in the regulation of the action of intracellular phospholipases A2 in vivo. The same authors "vertically compressed" the phosphoglycerol monolayers by substituting an alkylated glass plate for air while maintaining a constant surface pressure. 8 Subsequently, the activities of phospholipases A1 and A2 toward the monolayers were measured both in the presence and in the absence of the support. While phospholipase A1 activity was increased 4-fold by the support, the activity of phospholipase A2 was reduced to 15% of the activity measured in the absence of the alkylated surface. These findings indicate that this "vertical compression" exerted on the monolayer is likely to have induced a conformational change in the phospholipid molecules, which in turn may have caused the above reciprocal changes in the activities of phospholipases Al and A2.

7 T. Thuren, A. P. Tulkki, J. A. Virtanen, and P. K. J. Kinnunen, Biochemistry 26, 4907 (1987). 8 T. Thuren, J. A. Virtanen, and P. K. J. Kinnunen, Biochemistry 26, 5816 (1987).

52

PHOSPHOLIPASE ASSAYS, KINETICS, SUBSTRATES

Hydrolytic Action of Phospholipase Transition Region

A2

[4]

in Monolayers in the Phase

Grainger et al. 9A° have characterized optically the phospholipase A2 (Naja naja) during its action against a variety of phospholipid monolayers using fluorescence microscopy. By labeling the enzyme with a fluorescent marker (fluorescein), the hydrolysis of lipid monolayers in their liquidsolid phase transition region could be directly observed with the assistance of an epifluorescence microscope. Visual observation of hydrolysis of different phospholipid monolayers in the phase transition region in realtime could differentiate various mechanism of hydrolytic action against lipid solid phase domains. Lipid monolayers were spread over a buffer subphase at temperature necessary (30° for dipalmitoylphosphatidylcholine) to reach the monolayer liquid-solid phase transition at 22 mN/m for each respective lipid. Dipalmitoylphosphatidylcholine solid phase domains were specifically targeted by phospholipase A 2 and were observed to be hydrolyzed in a manner consistent with localized packing density differences, as illustrated in Fig. 2. Dipalmitoylphosphatidylethanolamine lipid domain hydrolysis showed no such preferential phospholipase Az response but did demonstrate a preference for solid/lipid interfaces. In all cases, after critical extents of monolayer hydrolysis in the phase transition region, highly stable, organized domains of enzyme of regular sizes and morphologies were consistently seen to form in the monolayers. Enzyme domain formation was entirely dependent upon hydrolytic activity in the monolayer phase transition region and was not witnessed otherwise.

Zero-Order Trough Several types of troughs have been used to study enzyme kinetics. The simplest of these is made of Teflon and is rectangular in shape (Fig. 1), but it gives nonlinear kinetics. II To obtain rate constants, a semilogarithmic transformation of the data is required. This drawback was overcome by a new trough design (zero-order trough, Fig. 3), consisting of a substrate reservoir and a reaction compartment containing the enzyme solution.12 The two compartments are connected to each other by a narrow surface canal. The kinetic recordings obtained with this trough are linear, unlike the nonlinear plots obtained with the usual one-compartment trough. The buffer is filtered and the pH is adjusted immediately prior to use. Before 9 D. W. Grainger, A. Reichert, H. Ringsdorf, and C. Salesse, FEBS Lett. 252, 73 (1989). l0 D. W. Grainger, A. Reichert, H. Ringsdorf, and C. Salesse, Biochim. Biophys. Acta 1023, 365 (1990). II G. Zografi, R. Verger, and G. H. de Haas, Chem. Phys. Lipids 7, 185 (1971). 12 R. Verger and G. H. de Haas, Chem. Phys. Lipids 10, 127 0973).

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS liquid-analogous lipid phase

53

solid-analogous lipid 9~omains

A

B

C

hydrolized monolayer (mixture of PC, LysoPC and fatty acid)

D

protein domain

E

FIO. 2. Hydrolysis of a lipid monolayer by phospholipase A 2 (schematic): (A), monolayer in the phase transition region with solid analogous lipid domains in a fluid analogous matrix mixed with a sulforhodamine marker; (B), injection of the FITC-labeled phospholipase A2; (C), specific recognition of the substrate lipids by the enzyme and preferential attack at boundaries between the solid analogous and the liquid analogous lipid phase; (D), hydrolysis of the solid analogous lipid domains and accumulation of the hydrolysis products in the monolayer; (E), aggregation of the enzyme to domains of regular morphology. [From D. W. Grainger, A. Reichert, H. Ringsdorf, and C. Salesse, Biochim. Biophys. Acta 1023, 365 (1990).]

54

P H O S P H O L I P A SASSAYS, E KINETICS, SUBSTRATES

[4] n

om ° momo o

•

• o

•

m o mo o m o m

• omo'-

m n mm • m m m m m

• n mu mn m m mm m m n mm m

m mm mm • m m mm • • mmmmm • mmm

omom mo mo o • o • momO

m m • • m m

omom

m m m m

~d Substrate

• mn m m mnn mm m m m m mm m m m m m m m m m m m m m mm

m m mm m mmm • m mm m m

m m m

• •

• •

m m m • mm m • mmm m

m m m m m m m m

~ Nonsubstrate

I~ Solubleproducts

FI6.3. Principle of the method for studying enzymatic lipolysis of mixed monomolecular films. [From G. Pi6roni and R. Verger, J. Biol. Chem. 254, 10090 (1979).]

each experiment, the trough is carefully cleaned with a paintbrush using absolute ethanol, rinsed several times with tap water until the Teflon surface no longer retains drops of water, and then rinsed twice with distilled water. Set-up and completion of a typical kinetic experiment requires about 30 min. Unlike previous experiments, no detergent solution is employed. The surface pressure is maintained constant automatically by the surface barostat method described elsewhere. 12Fully automatized monolayer systems of this kind are now commercially available (KSV, Helsinki, Finland).

Film Recovery and Estimation of Bound Enzyme The main difference between the monolayer and the bulk system lies in the ratios of interfacial area to volume, which differ by several orders of magnitude. In the monolayer system, this ratio is usually about 1 cm -1,

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

55

depending on the depth of the trough, whereas in the bulk system it can be as high as 105 cm -~, depending on the amount of lipid used. Consequently, under bulk conditions, the adsorption of nearly all the enzyme occurs at the interface, whereas with a monolayer only one enzyme molecule out of a hundred may be at the interface. H Owing to this situation, a small but unknown amount of enzyme, responsible for the observed hydrolysis rate, is adsorbed on the monolayer. In order to circumvent this limitation, two different methods were proposed for recovering and measuring the quantity of enzymes absorbed at the interface. 13-15 After performing velocity measurements, Momsen and Brockman ~5 transferred the monolayer to a piece of hydrophobic paper, and the adsorbed enzyme was then assayed titrimetrically. The paper was pretreated by soaking for several hours in deionized water at pH 7 and then equilibrated overnight in air saturated with water vapor. When the mobile bar was about 4 cm from the end of the trough, the paper was lowered onto the surface and left in contact for about 15-20 sec. Leaving the paper on the surface up to 5 min did not affect the amount of lipase adsorbed. The paper was pulled gently over the edge of the trough, and any residual subphase droplets were shaken off. Subphase carryover at pressures of 10, 16, and 22 mN/m was found to be 20 -+ 4/zl (n = 14) when measured gravimetrically. After correcting for the blank rate and subphase carryover, the amount (moles) of adsorbed enzyme was calculated from the net velocity and the specific enzyme activity. In assays performed with radioactive enzymes, 13the film was aspirated by inserting the end of a bent glass capillary into the liquid meniscus emerging above the ridge of the Teflon compartment walls, as depicted in Fig. 4C. The other end of the same capillary was dipped into a 5-ml counting vial connected to a vacuum pump. As aspiration proceeded, the film was compressed with a mobile barrier to facilitate quantitative recovery (0.5 ml of liquid with a film of 120 cm2). The capillary was broken into pieces which were added to the vial before counting. As radioactive molecules dissolved in the subphase were unavoidably aspirated with the film constituents, the results had to be corrected by counting the radioactivity in the same volume of aspirated subphase. The difference between the two values, which actually expressed a certain excess of radioactivity existing at the surface, was attributed to the enzyme molecules bound to the film.~3 The surface-bound enzyme includes not only those enzyme molecules directly involved in the catalysis but also an unknown amount of protein 13 j. Rietsch, F. Pattus, P. Desnuelle, and R. Verger, J. Biol. Chem. 252, 4313 (1977). 14 S. G. Bhat and H. L. Brockman, J. Biol. Chem. 256, 3017 (1981). 15 W. E. Momsen and H. L. Brockman, J. Biol. Chem. 256, 6913 0981).

56

PHOSPHOLIPASE ASSAYS, KINETICS, SUBSTRATES

[4]

To vacuum pump

A

B

C

D

E

FIG. 4. Diagram of the method used for enzyme kinetic experiments after filmrespreading. (A) Enzyme injection. (B) First kinetic recording. (C) Film aspiration. (D) Film respreading. (E) Second kinetic recording.

present close to the monolayer. These enzyme molecules are not necessarily involved in the enzymatic hydrolysis of the film. Since it is possible with the monolayer technique to measure the enzyme velocity (expressed in/zmol/cm2/min) and the interfacial excess of enzyme (mg/cm2), it is easy to obtain a value of enzymatic specific activity, which can be expressed as usual (in/zmol/min/mg). As illustrated in Fig. 4, Ransac et al. 16devised a simple technique for estimating the amount of unlabeled enzyme bound to the monomolecular film. This technique involves the dual kinetic recording of enzyme velocity on two monomolecular films consecutively spread over the same enzyme solution. As usual, the enzyme was injected at a final concentration of 50 pM (Fig. 4A) into the subphase of the reaction compartment (100 cm 2) of a zero-order trough, and the first kinetic recording of enzyme velocity was performed during 10 min (Fig. 4B). The film was then collected (Fig. 4C) as described previously. The second substrate film was spread at the same final surface pressure over the remaining subphase containing the residual enzyme (Fig. 4D), and a second kinetic recording was performed (Fig. 4E). Such an experiment takes about 40 min. By comparing the enzymatic rates between the two kinetic experiments, it is possible to estimate the fraction of enzyme removed during the film aspiration. This technique turned out to be particularly well suited for determining the amount of native phospholipase A2 bound to monolayers of substrate analogs. 16

~6S. Ransac, to be published.

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

57

Lag Periods in Lipolysis

A fairly common observation reported by many authors working on the kinetics of lipolytic enzymes is the occurrence of lag periods in the hydrolysis of emulsions, liposomes, micelles, and monolayers. 17-29 The zero-order trough gave, as might be expected, linear kinetics after injection of venom enzyme under the monolayer. 21This was not the case, however, when pancreatic phospholipase Az was injected under a film of dinonanoyllecithin. In fact, the velocity, as given by the slope of the recorded curve, was found to increase with time and seemed to approach an asymptotic limit: the intercept between the asymptote and the time axis is the lag time r. This behavior contrasts strongly with the kinetics obtained after injection of pure phospholipase A2 from snake or bee venom under the same lecithin film. The exact reasons for the unusually long lag periods are still open to debate. 3° Bianco et al. 31 have observed, however, that changes in surface potential parallel the variation observed in the enzyme velocity with time. A plateau was found to occur corresponding to a decrease in potential between 20 and 70 mV. This decrease in surface potential depends linearly on the enzyme concentration in the subphase, and it probably reflects the enzyme adsorption at the interface. Demel et al. 32 and Verheij et a l . 33'34 measured the influence of the surface pressure on the lag time of several phospholipases A z . The authors classified these enzymes unambiguously depending on their penetration power. With each enzyme, there exists a characteristic critical substrate 17 H. L. Brockman, J. H. Law, and F. J. K6zdy, J. Biol. Chem. 248, 4965 (1973). is O. A. Roholt and M. Schlamowitz, Arch. Biochem. Biophys. 94, 364 (1961). 19 A. F. Rosenthal and M. Pousada, Biochim. Biophys. Acta 164, 226 (1968). 20 R. H. Quarles and R. M. C. Dawson, Biochem. J. 113, 697 (1969). 21 R. Verger, M. C. E. Mieras, and G. H. de Haas, J. Biol. Chem. 248, 4023 (1973). 22 M. K. Jain and R. C. Apitz-Castro, J. Biol. Chem. 253, 7005 (1978). 23 D. O. Tinker and J. Wei, Can. J. Biochem. 57, 97 (1979). 24 F. Pattus, A. J. Slotboom, and G. H. de Haas, Biochemistry 13, 2691 (1979). 25 G. C. Upreti, S. Rainier, and M. K. Jain, J. Membr. Biol. 55, 97 (1980). 26 G. C. Upreti and M. K. Jain, J. Membr. Biol. 55, 113 (1980). 27 B. Borgstr6m, Gastroenterology 78, 954 (1980). 2s K. Hirasawa, R. F. Irvine, and R. M. C. Dawson, Biochem. J. 193, 607 (1981). 29 M. Menashe, D. Lichtenberg, C. Gutierrez, and R. L. Biltonen, J. Biol. Chem. 256, 4541 (1981). 3o M. K. Jain and O. G. Berg, Biochim. Biophys. Acta 1002, 127 (1989). 31 I. D. Bianco, G. D. Fidelio, and B. Maggio, Biochem. J. 258, 95 (1989). 32 R. A. Demel, W. S. M. Geurts van Kessel, R. F. A. Zwaal, B. Roelofsen, and L. L. M. van Deenen, Biochim. Biophys. Acta 406, 97 (1975). 33 H. M. Verheij, M. R. Egmond, and G. H. de Haas, Biochemistry 20, 94 0981). H. M. Verheij, M. C. Boffa, C. Rothen, M. C. Brijkaert, R. Verger, and G. H. de Haas, Eur. J. Biochem. 112, 25 (1980).

58

PHOSPHOLIPASE ASSAYS, KINETICS, SUBSTRATES

[4]

packing density above which the enzyme cannot penetrate the film. This type of information can be of practical value when choosing a lipolytic enzyme for degrading the lipid moiety of biological membranes 32 or for predicting the anticoagulant properties of a given phospholipase A 2.34 Mixed Monolayers as Phospholipase Substrates

Most studies on lipolytic enzyme kinetics have been carried out in vitro with pure lipids as substrates. Actually, however, virtually all biological interfaces are composed of complex mixtures of lipids and proteins. The monolayer technique is ideally suited for studying the mode of action of lipolytic enzymes at interfaces using controlled mixtures of lipids. There exist two methods of forming mixed-lipid monolayers at the air-water interface: spreading a mixture of water-insoluble lipids from a volatile organic solvent or injecting a micellar detergent solution into the aqueous subphase covered with preformed insoluble lipid monolayers. A new application of the zero-order trough was proposed by Pi6roni and Verger 35for studying the hydrolysis of mixed monomolecular films at constant surface density and constant lipid composition (Fig. 3). A Teflon barrier was disposed transversely over the small channel of the zero-order trough in order to block surface communication between the reservoir and the reaction compartment. The surface pressure was first determined by placing the platinum plate in the reaction compartment, where the mixed film was spread at the required pressure. Then surface pressure was measured after switching the platinum plate to the reservoir compartment where the pure substrate film was subsequently spread. The surface pressure of the reservoir was equalized to that of the reaction compartment by moving the mobile barrier. The subphase of the reaction compartment, composed of a standard buffer (10 mM Tris/acetate, pH 6.0, 0.1 M NaC1, 21 mM CaC12, 1 mM EDTA, thermostatted at 25° -+ 0.5°), was stirred at 250 rpm with two magnetic bars. The barrier between the two compartments was then removed in order to allow surfaces to communicate. The pressure change during these operations did not exceed 0.25 dyne/cm. The enzyme was then injected into the reaction compartment and the kinetics recorded as described. 12 The authors 36 studied the hydrolysis of mixed monomolecular films of phosphatidylcholine/triacylglycerol by pancreatic phospholipase A2. The quantity of enzyme adsorbed on the interface was concomitantly determined with 3H-amidinated phospholipase. At phospholipid packing levels 35 G. Pi6roni and R. Verger, J. Biol. Chem. 254, 10090 (1979). t6 G. Pi~roni and R. Verger, Eur. J. Biochem. 132, 639 (1983).

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

59

above the critical penetration pressure, triacylglycerol considerably enhances phosphatidylcholine hydrolysis. On the other hand, the activity of pancreatic phospholipase A2 on a mixed film is inhibited by the action of pancreatic lipase. Using the same methodology, Alsina et al. 37 studied the enzymatic lipolysis by pancreatic phospholipase A2 and by phospholipase A2 of Vipera berus on monomolecular films of mixture of natural lipids: cholesterol-egg lecithin and triolein-egg lecithin, by adding serum albumin to the aqueous subphase in order to imitate the physiological conditions. Later on Alsina et al. 3a reported a complementary study on the lipolysis of didecanoyl phosphatidylcholine/triolein mixed monolayers by phospholipase A 2. Grainger et al. 39 have characterized physically and have studied the enzymatic hydrolysis of mixed monolayers of a natural phospholipid substrate and a polymerizable phospholipid analog. Enzyme hydrolysis showed large differences in the ability of the enzyme to selectively hydrolyze the natural phosphatidylcholine component from the monomeric as opposed to the polymeric mixtures. The results clearly show a strong influence of molecular environment on phospholipase A 2 activity, even if differences in the physical state of mixed monolayers are not detectable with isotherm and isobar measurements. Inhibition o f Phospholipases Acting on Mixed Substrate/Inhibitor Monomolecular Films

Many drugs and phospholipid analogs have been reported to act as phospholipase "inhibitors." A priori, these compounds can be said to interfere with phospholipase A 2 (PLA2) activity by interacting either directly with the enzyme or indirectly by affecting the "interfacial quality" of the substrate. A number of "membrane-active" compounds have marked effects. For instance, low concentrations of alcohols and detergents can stimulate PLA2-catalyzed hydrolysis of lipid bilayers, whereas higher concentrations are inhibitory. The effects of these compounds are related somehow to their amphiphilic nature, and it is believed that they may act as spacer molecules facilitating the penetration of the bilayer by the enzyme. An illustration has been provided by Vainio et al., 4° who 37 A. Alsina, O. Vails, G. Pirroni, R. Verger, and S. Garcia, Colloid Polym. Sci. 261, 923 (1983). 3s M. A. Alsina, M. L. Garcia, M. Espina, and O. Valls, ColloidPolym. Sci. 267, 923 (1989). 39 D. W. Grainger, A. Reichert, H. Ringsdorf, C. Salesse, D. E. Davies, and J. B. Lloyd, Biochim. Biophys. Acta 1022, 146 (1990). 4o p. Vainio, T. Thuren, K. Wichman, T. Luukkainen, and P. J. K. Kinnunen, Biochim. Biophys. Acta 814, 405 (1985).

60

P H O S P H O L I P A SASSAYS, E KINETICS, SUBSTRATES

[4l

I FIG. 5. Proposed model for competitive inhibition at interfaces. A, Total interracial area; V, total volume; E, enzyme concentration; P, product concentration; S, interracial concentration of substrate; I, interfacial concentration of inhibitor; D, inteffacial concentration of detergent. [From S. Ransac, C. Rivi~re, J. M. Souli6, C. Gancet, R. Verger, and G. H. de Haas, Biochim. Biophys. Acta 1043, 57 (1990).]

reported that human spermatozoa hydrolyzed only phosphatidylglycerol monolayers. Inhibition of the phospholipase activity by gossypol may contribute to the unknown contraceptive effects of this nonsteroid male antifertility agent. It is now becoming clear from the abundant literature on lipolytic enzymes that any meaningful interpretation of inhibition data has to take into account the kinetics of enzyme action at the lipid-water interface. As shown in Fig. 5, Ransac et al. 4~ have devised a kinetic model which is applicable to water-insoluble competitive inhibitors in the presence of detergent in order to quantitatively compare the results obtained at several laboratories. Furthermore, with the kinetic procedure developed, it was possible to make quantitative comparisons with the same inhibitor placed under various physicochemical situations, namely, micellar or monolayer states. The addition of a potential inhibitor to the reaction medium can lead to paradoxical results. Usually, variable amounts of inhibitor are added, at a constant volumetric concentration of substrate and detergent. The specific area, the interfacial concentration of detergent, and the interfacial concentration of substrate are continuously modified accordingly. In order to minimize modifications of this kind, it was proposed to maintain constant the sum of inhibitor and substrate (! + S) when varying the inhibitory molar fraction [o~ = I / ( I + S)], that is, to progressively substitute a molecule of inhibitor for each molecule of substrate. 42 This is the minimal change that can be made at increasing inhibitor concentrations. Of course, with this method, the classic kinetic procedure based on the 41 S. Ransac, C. Rivi~re, J. M. Souli6, C. Gancet, R. Verger, and G. H. de Haas, Biochim. Biophys. Acta 1043, 57 (1990). 42 G. H. de Haas, M. G. van Oort, R. D. Dijkman, and R. Verger, Biochem. Soc. Trans. 17, 274 (1989).

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

61

Michaelis-Menten model is not valid because both inhibitor and substrate concentrations vary simultaneously and inversely. However, by measuring the inhibitory power (Z) as described by Ransac et al. 41 or Si(50 ) as used by Jain et a1.,43 it is possible to obtain a normalized estimation of the relative efficiency of various potential inhibitors. Using the mixed-film technique and the zero-order trough described in Fig. 3, Ransac et al. 14 studied the hydrolysis of monomolecular films of Ldilauroylphosphatidylcholine or L-dilauroylphosphatidylglycol mixed with their corresponding phospholipid analogs bearing an amide bond instead of an ester bond at the 2-position of the glycerol backbone. These amidophospholipids, incorporated into mixed micelles, strongly and stereoselectively inhibited phospholipases A2 with an increased inhibitory power in the case of the anionic derivatives. 44 Kinetic experiments were performed at various molar fractions of inhibitor (a) present in mixed monomolecular films of substrate/inhibitor. Figure 6 shows the decrease in the relative phospholipase activities as a function of the inhibitor molar fraction at the optimum surface pressure with each substrate used. Both inhibitors of the L-phosphatidylcholine and L-phosphatidylglycol series exhibited strongly concave-shaped curves, whereas those obtained with inhibitors of the D series were rather close to the diagonal, which probably corresponds to a special case where KI* = Km*. The inhibitory powers (Z) were given by the slopes of the straight lines obtained by plotting Ro as a function of a (see insets). Inhibitors of the L series were found to be stronger inhibitors than those of the O series, and L-amido-C12-Pglycol (Z = 38) was more potent than L-amidoC12-PC (Z = 25). Figure 6A also shows that the lag times (z) due to the presence of inhibitors of the L-phosphatidylcholine series increased as a function of the inhibitor molar fraction (a). This is in good agreement with the simulation curve obtained from the kinetic model presented in Fig. 5. Enzymatic Activity of Phospholipase C (a Toxin from Clostridium perfringens) Using Phospholipid Monolayers as Substrate Several research g r o u p s 32'45'46have published kinetics studies on phospholipase C using surface radioactivity as an index to the splitting and the solubilization of the phosphorylcholine moiety in the water subphase. This method can give interesting qualitative information about the initial steps 43 M. K. Jain, W. Yuan, and M. H. Gelb, Biochemistry 28, 4135 (1989). 44 G. H. de Haas, R. D. Dijkman, M. G. van Oort, and R. Verger, Biochim. Biophys. Acta 1043, 75 (1990). 45 A. D. Bangham and R. M. C. Dawson, Biochim. Biophys. Acta 59, 103 (1962). 46 I. R. Miller, and J. M. Ruysschaert, J. Colloid Interface Sci. 35, 340 (1971).

62

PHOSPHOLIPASE ASSAYS, KINETICS, SUBSTRATES

[A]

100

Rv ,5

/

[4]

80

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\ \ \ \~x

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FIG. 6. Relative velocity, induction time (¢), and R, (inserts) as a function of inhibitor molar fraction (a). Inhibitors of the I) series ( - - ) . Inhibitors of the L series ( ). (A) LDiC12-PC used as substrate, L-amido-C12-PC ( 0 ) and D-amido-Czz-PC ( I ) used as inhibitors. Surface pressure was 10 m N / m . (B) L-DiC12-Pglycol used as substrate, L-amido-Cl2-Pglyco[ (©) and D-amido-C12-Pglycol (D) used as inhibitors. Surface pressure was 30 mN/m.

[4]

PHOSPHOLIPASE KINETICS IN LIPID MONOLAYERS

63

during phospholipase C action. However, the main limitation of this technique is that insoluble diglyceride molecules accumulate with time, perturbing the initial phospholipid film. In other words, one of the products (diglycerides) of the reaction catalyzed by phospholipase C remaining in the interface can have a profound influence and progressively reduce the enzyme velocity. In order to circumvent these product accumulation problems, Moreau e t al. 47 have developed a new method based on the subsequent rapid lipase hydrolysis of the diglyceride generated by phospholipase C. In these assays, the authors used a large excess of pancreatic lipase saturated by colipase (ranging from 0.15 to 10 nM lipase and a concentration of 40 pM of phospholipase C). It is worth noting that the pancreatic lipase/colipase system is not able to hydrolyze a pure phosphatidylcholine film. These experiments were carried out under the following experimental conditions: L-dilauroylphosphatidylcholine film spread at a surface pressure of 14 mN/m over a subphase of 20 mM Tris-HC1 buffer, pH 7.2, 0.15 M NaCI, 5 m M CaCI2, and 0.1 mM ZnO4. Using the barostat technique, 1'12it was checked that the enzyme kinetics were linear and that the velocity was directly dependent on the amount of phospholipase C added. This system can be used to study in detail the enzymatic kinetics of phospholipases C on lipid monolayers. This method is applicable to either medium-chain phospholipid monolayers or longchain phospholipid monomolecular films providing that, in the latter case, bovine serum albumin (BSA) is introduced into the subphase to dissolve the fatty acid and monoglyceride formed. Bianco e t al. 4s have recently used this new method to study the effects of sulfatide and gangliosides on phospholipase C activity.

Film Transfer Experiments Showing Role of Ca 2+ and Zn 2+ Ions in Phospholipase C Activity When EDTA (0.1 mM) was introduced into the aqueous subphase of the reaction compartment, the phospholipase C activity measured under optimal conditions was immediately and completely abolished although, under these conditions, pancreatic lipase is known to be fully active, j3 Hydrolysis was not restored after addition of either Zn 2÷ (0.5 mM final concentration) or Ca 2+ (5 mM final concentration) alone. The simultaneous presence of the two cations in the aqueous subphase was necessary for the activity of the phospholipase C to be recovered. 47 H. Moreau, G. Pi6roni, C. Jolivet-Reynaud, J. E. Alouf, and R. Verger, Biochemistry 27, 2319 (1988). 4s I. D. Bianco, G. D. Fidelio, and B. Maggio, Biochim. Biophys. Acta, in press (1990).

64

PHOSPHOLIPASE ASSAYS, KINETICS, SUBSTRATES

PHOSPHOLIPASE C

A

r INJECTION

~

I

/

"

~

¢

PHOSPHOLIPASE C INJECTION

_

n

I A

B

START TRANSFER

B A ~ N N ' ~ I ~

FILM RINSE AND TRANSFER

Za ~ INJECTION

t

C

A D/ ~ C i A,t,LAAAAAA,t~AA~,)..t/./,AIA,tm

1

2

q

C

A B C Ik,A,AAikAA,,AAA,,,k~AA,Uk

A B C h A~Ah~AAAAA~AAAAAA~ .~b

FILM RINSE AND TRANSFER

rl

A/

Ca*+ INJECTION

~. PHOSPHOLIPID

~

C

IL , AAAAAA~AAL , AA~L , AAAA ~A Am

!

3 • PNOSPHOLIPASE C

II

A /

T

C

START TRANSFER

[4]

Zn*+

1

2

~

ell"

3

FIG. 7. Transfer experiments showing the role of Ca z÷ and Zn 2+ ions in phospholipase C activity. A, B, and C represent the three barriers; barrier C was mobile during the reaction recording. The graphs on the left-hand side of panels I and II are the kinetic curves recorded before (upper graph in I and II) and after (lower graph) the film transfer. Lipase saturated with colipase was present in all the reaction compartments. [From H. Moreau, G. Pirroni, C. Jolivet-Reynaud, J. E. Alouf, and R. Verger, Biochemistry 27, 2319 (1988).]

To determine the respective roles of Ca 2÷ and Z n 2+ ions on phospholipase C activity, transfer experiments were performed as described in Fig. 7. Lipase (1 nM final concentration) saturated with colipase was added to all three compartments containing 20 mM Tris-HC1 buffer, pH 7.2, 0.1 mM EDTA, and 0.15 M NaCI. The portion of the L-dilauroylphosphatidylcholine film at a surface pressure of 14 mN/m and located over compartment 1, where phospholipase C was first injected (40 pM final concentration), was isolated by barrier B and transferred to the surface of compartment 3 with a film rinse over compartment 2. In experiment II, Zn 2÷ ions (0.5 mM final concentration) alone were first present in the three compartments, and, after film transfer, Ca 2÷ (5 mM final concentration) was injected into the aqueous subphase of compartment 3; no enzymatic activity was observed. As a control, a further injection of phospholipase C into the aqueous subphase of compartment 3 was found to initiate film hydrolysis at a rate comparable to that observed previously. In experiment I, Ca 2÷ ions (5 mM final concentration) were initially present in all three compartments. After film transfer, injection of Zn 2÷ (0.5 mM final concentration) into the aqueous subphase of compartment 3 initiated an immediate phospholipase C activity, amounting to 60% of the optimal value previously determined.

[5]

T H I O ASSAY

65

When only Zn 2+ ions were present in the aqueous subphase, phospholipase C was not associated with the substrate film and consequently was not active. In the presence of Ca 2÷ ions only, phospholipase C was associated with the film, but its activity was dependent on the presence of Zn 2÷ ions. In conclusion, Ca 2÷ ions appear to be involved in the binding of the enzyme to the lipid interface, unlike Zn 2÷ ions, which are necessary for the expression of catalytic activity. Acknowledgments Our thanks are due to Dr. C. Rothen (Bern, Switzerland) and Professor G. H. de Haas (Utrecht University, the Netherlands) for personal communications prior to publication. S.R. acknowledges fellowship support from Groupement de Recherche de Lacq du Groupe Elf Aquitaine, France (Dr. C. Gancet and Dr. J. L. Seris). The authors are grateful to Dr. J. Blanc for revising the English and to M. T. Nicolas (Marseille) for typing the manuscript.

[5] T h i o - B a s e d P h o s p h o l i p a s e A s s a y By LIN Yu and EDWARD A. DENNIS Introduction

The thio assay possesses many characteristics that recommend it as a general assay for phospholipases. The most important are that it is a continuous, spectrophotometric assay which is very convenient, it directly detects one of the products liberated upon hydrolysis, it is one of the more sensitive assays, and it is also suitable for detailed kinetic studies. 1,2 The thio assay can be used for phospholipases A~3 and A24 and with appropriate modification of the substrate would be applicable to other phospholipases.5 However, owing to the lack of commercial availability of the thiophospholipid substrate and its complicated synthesis, the thio assay has not been used extensively. We 6 have recently modified the original 4 synthetic procedure so that gram quantities of chiral thiophospholipid can be readily I H. S. Hendrickson and E. A. Dennis, J. Biol. Chem. 259, 5734 (1984). 2 H. S. Hendrickson and E. A. Dennis, J. Biol. Chem. 259, 5740 (1984). 3 G. L. Kucera, C. Miller, P. J. Sisson, R. W. Wilcox, Z. Wiemer, and M. Waite, J. Biol. Chem. 253, 12964 (1988). 4 H. S. Hendrickson, E. K. Hendrickson, and R. H. Dybuig, J. LipidRes. 24, 1532 (1983). 5 j. W. Cox, W. R. Snyder, and L. A. Horrocks, Chem. Phys. Lipids 25, 369 (1979). 6 L. Yu, R. A. Deems, J. Hajdu, and E. A. Dennis, J. Biol. Chem. 265, 2657 (1990).

METHODS IN ENZYMOLOGY, VOL. 197

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