Lipid Mobility in the Assembly and Expression of the Activity of the Prothrombinase Complex*

Transcription

1 THE JOURNAL OF BIOLOGICAL CHEMISTRY by The American Society of Biological Chemists, Inc. Vol. 260, No. 6, lasue of March 25, PP ,1985 Printed in U. S. A. Lipid Mobility in the Assembly and Expression of the Activity of the Prothrombinase Complex* (Received for publication, July 24, 1984) Deborah L. HigginsS, Peter J. Callahant, Franklyn G. Prendergastt, Michael E. NesheimS, and Kenneth G. MannSn From the $Hematology Research Section and the Department of Pharmacology, Mayo Foundation, Rochester, Minnesota A phospholipid or membrane surface is a required component of the prothrombinase complex, yet little is known about the influence of the lipid on the assembly and expression of this complex. Vesicles composed of synthetic phospholipids were used to investigate the effects of membrane fluidity on the prothrombinase complex. All vesicle types studied were capable of supporting the prothrombinase reaction which in each case was characterized by a similar apparent K,. The binding constants for the interaction of Factor Va and prothrombin with synthetic phospholipid vesicles were not significantly affected by temperature. The rate of thrombin production, however, increased with increasing temperature. The fluidity of the vesicles was assessed by measuring the fluorescence lifetimes, steady state anisotropies, and differential phase fluorometry of diphenylhexatriene embedded in the vesicles. No correlation was observed between the fluidity of the vesicles and the steady-state rate of thrombin production, even when the enzymatic activity was monitored below and above the phase transition temperature of the lipid vesicles. A distinct correlation, however, was found between the fluidity of the vesicle and the time required to reach the maximum rate of thrombin production (pre-steady-state interval). We believe that this lag time corresponds to the time required for the assembly of the prothrombinase complex. Thus, although lipid fluidity does affect the assembly of the prothrombinase complex, after the complex is assembled, this property has littleffect on the catalytic process itself. A surface-bound catalyst is essential for the proteolytic conversion of prothrombin to the blood clotting enzyme thrombin at a physiologically significant rate (1). This surface-bound catalyst, the prothrombinase complex, is composed of the serine protease Factor Xa, the protein cofactor Factor Va, Ca2+ ions, and a membrane or phospholipid surface (2-5). Although platelets are believed to supply one type of surface for prothrombinase assembly in vivo, phospholipid vesicles also provide a model system for study of complex assembly, as they can support formation of a complex with the same catalytic efficiency as seen on platelets (1). Both the substrate (prothrombin) and the enzyme (Factor * This work was supported by National Institutes of Health Grant HL17430-D. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked aduertisernent in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 11 To whom correspondence should be addressed Chairman, Department of Biochemistry, The University of Vermont, Burlington, VT Xa) bind to negatively charged phospholipids through interactions thought to involve y-carboxyglutamate residues and Ca2+ ions (6-10). The binding of Factor V and Factor Va also required negatively charged phospholipids (11, 12). However, this binding is independent of Ca2+. Factor Va also serves as a receptor for Factor Xa both on platelets (13-15) and on phospholipid vesicles (16). The role of the phospholipid surface in the prothrombinase complex has been studied primarily by investigating the binding of enzyme and substrate to vesicles of differing composition (17, 18). The effects of hydrocarbon chain length and hydrogenation on procoagulant activity have also been analyzed (19,20). More recently, however, studies using synthetic phospholipids have correlated lipid fluidity and surface charge directly with catalysis (21). It is difficult to quantify procoagulant activity from the data in these previous studies because this activity was determined from a plasma clot time assay, and this end-point (time required for a given change) assay is dependent on numerous lipid-dependent and lipidindependent interactions involving catalytic as well as binding processes. In order to deduce which interactions are influenced by lipid fluidity, we have conducted studies utilizing synthetic phospholipids and purified proteins in the coagulant assays. In addition, we have evaluated the relationship of fluidity to both catalytic and binding interactions. Vesicles were composed of synthetic phospholipids, so that the composition of the fatty acyl side chains could be varied to construct vesicles exhibiting differing fluidity characteristics. Fluidity was assessed by monitoring the temperature dependence of the fluorescence lifetimes, steady-state anisotropies, and by use of differential-phase fluorometry on probes embedded in the vesicles. These results were compared to the temperature dependence of not only prothrombin and Factor Va binding, but also to the rate of thrombin formation in the presence of purified Factor Va and Factor Xa, to help discern the basis for the rate enhancement exhibited by the prothrombinase complex in the presence of phospholipid and the role of phospholipid in the complex. EXPERIMENTALPROCEDURES Materials DMPC, DMPS, DOPC, and POPS were purchased from Avanti Polar Lipids. Cholesterol was from Nucheck Prep. Crude PC from The abbreviations used are: DPMC, ditetradecanoylglycerophosphatidylcholine; DMPS, ditetradecanoylglycerophosphatidylserine; DOPC, dioleoy!glycerophosphatidylcholine; POPS, l-palmitoyl-2- oleoylglycerophosphatidylserine; PC, L-a-phosphatidylcholine; PS, L- a-phosphatidylserine; DPH, diphenylhexatriene; dansyl, 5-dimethylaminonaphthalene-1-sulfonyl; DAPA, dansylarginine-n-(3-ethyl- 1,5-pentanediyl)amide; HEPES, N-2-hydroxyethylpiperazine-Nf-2- ethanesulfonic acid; DEGR, dansyl-glutamyl,glycyl,arginyl.

2 egg yolk and PS from bovine brain were purchased from Sigma. A11 lipids were used without further purification. DPH was purchased from Aldrich. DAPA was synthesized from dansylarginine and 4-ethylpiperidine as described previously (22). Purification of Proteins Bovine Factor V was isolated and assayed as described previously (5, 23). The protein preparations typically exhibited an activation quotient in replacement bioassays of at least 30 (activity of thrombinactivated Factor V(Va)/activity of Factor V which had not been intentionally activated) and a specific activity of greater than 1500 units/mg when fully activated and analyzed in human Factor V- deficient plasma uersus a bovine plasma standard. The protein was stored as Factor V at -20 "C in 50% glycerol, 0.01 M Tris, 0.01 M borate, 1 mm CaC12, ph 8.3, until use. Under these conditions Factor V preparations have been stored for up to 2 years without noticeable deterioration of either activity or electrophoretic behavior. Factor V was activated to Factor Va by the folfowing procedure. Factor V was diluted to 0.5 mg/ml in 0.02 M imidazole, 5 mm CaCI,, ph 7.5. The Factor V was activated hy adding bovine a-thrombin to a final concentration of 2 NIH units/ml, and the solution was then incubated at 37 "C until an assay showed complete activation (approximately 5 min). The solution was made 1 mm in diisopropyl fluorophosphate, and the ph was brought to 6.5. Thrombin was removed by passage over a column of SPC-50, and the Factor Va was purified from the activation peptides by application to a QAE (quaternary ~inoethy1)- cellulose column. The Factor Va was eluted using a gradient of 0.02 M imidazole, 5 mm CaCI2, ph 6.5, to 0.02 M imidazole, 5 mm CaCh, 0.3 M NaCl, ph 6.5. The Factor Va was precipitated with 75% (NHs),S04. Prior to use (within 24 h of purification) the Factor Va was dialyzed against the appropriate buffer. Bovine prothrombin and Factor X were prepared as described by Bajaj and Mann (24). Factor X was activated to Factor Xa using electrophoretically homogeneous Factor X activator from Russell's viper venom, immobilized on cyanogen bromide-activated agarose (25). The Factor X activator was supplied by Dr. Walter Kisiel of the University of Washington at Seattle, WA. All proteins were electrophoretically homogeneous based on polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (26) and are stable in activity and electrophoretic behavior for at least 1 year if stored in glycerol-h,o (5050) at -20 "C. The molecular weights and extinc~ion coef~cients (t:zm,mnm) of the respective bovine proteins were taken as follows: Factor v, 330,0~, 9.6 (23); Factor Va, 168,000, 12.2'; Factor Xa, 55,100, 12.4 (27, 28); and prothrombin, 72,000, 14.4 (29, 30). Methods F~sp~l~pid Vesicle Prepurat~n-Single bilayer phospholipid vesicles of homogeneous size were prepared by modification of the method of Barenholz et al. (31). Aliquots of the various lipids were mixed in appropriate amounts for a final total weight of 5 mg. The lipid solution was dried under a stream of nitrogen, The lipids were then dispersed in 5 ml of 0.02 M HEPES, 0.15 M NaCl, ph 7.4, and sonicated using a direct probe with a Heat Systems Ultrasonics sonicator (model W-375) with continuous sonication for 30 min at 3- output control. The sample was main~ined an ice bath (except for DMPS/DMPC mixtures which were maintained at 37 "C) and under a constant stream of nitrogen during sonication. Following sonication, the vesicle dispersion was centrifuged for 30 min at 35,000 rpm in a Beckman model L3-50 ultracentrifuge (SW 50-1 swinging bucket rotor) to remove particles and large multilamellar vesicles. Then the dispersion was centrifuged at 40,000 rpm for 3 h, and the clear supernatant region I11 (31) was removed, This supernatant containing a homogeneous vesicle dispersion was used within 24 h of preparation. The phospholipid concent.ration of the stock solution was based upon organic phosphate determined by a protocol developed by the Mayo Clinic Lipids Laboratory for serum phospholipid determination (32) using a w/w conversion factor of 25 (phospholipid/p). The size of the phospholipid vesicles was determined using quasielastic light scattering (33). The apparatus employed consisted of an argon laser from Spectra-Physics (model 165) with a model 265 exciter. The high voltage power supply (model 215) was from Malvern _I * Determined by differential refractometry by M. E. Nesheim and K. G. Mann (unpublished results) Scientific, as was the photomultiplier. A Nicomp Instruments autocorrelator (model 6862) with a Tectronix T921 oscilloscope and a Malvern Scientific (model RR56~~emperature controller were used. Measurements were made at 488 A and 25 "C. Quantitative Binding Measurements-Relative 90" light-scattering measurements were made in a thermostated cell compartment in an SLM-Aminco (model 4800) fluorometer. Excitation and emission wavelengths were both set at 320 nm. The samples contained 2.0 ml of 0.02 M HEPES, 0.15 M NaCl, 2 mm Ca2+, ph 7.4, and a phospholipid concentration of 15 p ~ Protein. samples were added to the cuvette, and the light scattering from the protein-phospholipid complex was compared to the scatter due to the protein added to buffer solution. The light-scattering data were analyzed by the method developed by Nelsestuen and Lim (34). The value of the change in refractive index with concentration (&/&) for phospholipid was The refractive indexes for Factor V, Factor Va, and the components derived from these proteins were assumed to be This appears to be a reasonable assumption, since the amino acid and carbohydrate compositions of Factor V and Factor Va are not appreciably different from those of prothrombin and Factor X (35). The refractive index of a protein-lipid complex is assumed to be a weighted average of the protein and lipid refractive indexes. Fluorescence Lifetime Measurements-Phospholipid vesicles were labeled with DPH by placing 10 p1 of a DPH stock solution (1 mm DPH in acetonitrile) in a test tube. After the acetonitrile was evaporated under Nz, 2 ml of vesicles in 0.02 M HEPES, 0.15 M NaCi, ph 7.4, were added to the test tube. The sample was placed in a water bath above the phase transition of the tipids for 5-10 min and then sonicated in a bath sonicator for approximately 2 min. The cycle of heating and sonicating was repeated until the DPH had been sufficiently incorporated into the vesicles. Measurements of fluorescence lifetimes (71, steady-state anisotropies, and differential phase fluorometry (AT) were performed on an SLM subnanosecond fluorometer. Fluorescence lifetimes were measured by the phase-modulation method of Spencer and Weber (36) with a polarizer oriented to 54.7" in the emission path to negate effects of Brownian motion on 7 (37). Measurements of AT were made as described by Mantulin and Weber (38), Lakowicz andprendergast (39), and Lakowicz et al. (40). An excitation wavelength of 360 nm (slits, 0.5 mm) was used to excite the fluorescence of DPH. Fluorescence emission was isolated from Rayleigh and Raman scattering by placement of a Schott. KV 418-nm filter in the emission path. Steadystate anisotropies were measured by operation of the instrument in a "T" format in the manner described by Weber and ~ a b l o (41). u ~ ~ Again KV 418-nm filters were placed in the emission path to isolate fluorescence emission. Caiculations on Fluorescence Data-Steady-state anisotropies (r-) were calculated from the definition where Is and IL represent the intensities of fluorescence emission polarized vertically and horizontally, respectively, when the excitation is vertically polarized. Limiting hindered anisotropies as measured by differential phase fluorometry (rm, AT) were calculated from the relation where and m, = (1 -F 2rm)(l - r-1 3 = 2 + rc - r,(4r0-1) (40). As pointed out elsewhere (42), r,, AT1 and r,, (the latter determined by time-resolved measurements) are the same under the conditions of the experiments described here but are not necessarily synonymous. The fluorescence order parameter, S, was then determined from the relation r,/ro = S' (3) (according to Jahnig (43, 44); Heyn (45); Lipari and Szabo (46); and Engel and Prendergast (42)). This reiation holds providing that the substrate in which the probe is embedded does not contribute significantly to the depolarization of the fluorestence, i.e. does not exhibit

3 3606 a rotational correlation time similar to the fluorescence lifetime of the probe. The order parameter has a maximal value of one for a perfectly ordered system (e.g. for phospholipids in the gel phase of a lipid with fatty acyl chains rigid) and zero for a perfectly disordered system. As discussed by Jahnig (43, 44), Heyn (45), and Engel and Prendergast (42), changes in the order parameter for DPH describe the extent of disorder in the bilayer more precisely than does the fluorescence steady-state anisotropy. We have made no attempt to calculate microviscosities from the fluorescence data for reasons outlined by Engel and Prendergast (42), although, in principle, a microviscosity parameter can now be obtained from steady-state anisotropy data by use of the semi-empirical method of Hare (47). The qualitative picture of membrane disorder provided by the order parameter is, however, sufficient for our present purposes. Finally, it is important to note that although we have not provided any data, the same q~litutiue inte~retations were made when the fluorescence probe trimethylammonium DPH (48) was used. The latter is a fluorophore that resides at the membrane interface because of its cationic charge. Therefore, its location in the bilayer at temperatures greater than the phase transition temperature is known, in contrasto what is observed for DPH (42). Prothrombime Activity Experiments Monitored with DAPA-The activity of the prothrombinase complex was measured as described previously (1). This method is based of the fluorescence increase when the DAPA included in the reaction mixture binds to newly formed thrombin. A substrate solution composed of approximately 0.1 mg/ml bovine prothrombin in 0.02 M HEPES, 0.15 M NaC1, ph 7.4,3 X lo4 M DAPA, and 2 M CaClz was prepared at least 30 min prior to the start of the assays so that the Ca'+-dependent change in the conformation of prothrombin (49,501 occurred prior to the start of the reaction. A Perkin-Elmer MPF 44B fluorescence spectrophotomer contain- ing a thermostated cuvette holder was used for all assays. The excitation and emission wavelengths were 335 and 565 nm, respectively. The slit widths used were 10 and 15 nm, respectively. A 430- nm cutoff filter was used in the emission path. The sensitivity of the fluorometer was normally adjusted to give a deflection of 90% of the full scale upon completion of the reaction. Following the addition of 1.5 ml of the substrate solution to the cuvette, the base-line was adjusted to zero. Microliter quantities of the vesicle preparation of interest were added to yield a final phosphate concentration of 15 FM. Normally Factor Va was then added in microliter quantities to give a final concentration of 1.5 X lo-' M. At least 15 min was allowed for the temperature of the reaction mixture to reach the desired temperature. The reaction was then initiated by the addition of Factor Xa to a final concentration of 2 X lo-* M, and the subsequent time course of thrombin formation, observed as enhanced fluorescence intensity, was continuously recorded. Initial rates of thrombin production were calculated from the initial slope of the recorded data. In some instances a "lag" was seen to exist prior to the establishment of a steady-state rate. In these instances, the maximum initial slope of the recorded data was used to calculate the rate of thrombin production. In order to obtain the apparent K, and V,, for the reaction, the data were analyzed by the integrated Henri-Michaelis-Menten equation (51) as described by Nesheim et ul. (1). Measurement of Pre-steady-state Intervals by Kinetics-Prothrombin activation was initiated at the desired temperature, and the course of the reaction was continuously recorded from the fluorescence intensity of the DAPA-thrombin complex. Final concentrations of the components of the reactions were: prothrombin, Factor Xa, 2.0 nm; Factor Va, 15.0 nm; DOPC/DOPS or DMPC~MPS, 10 F~M; Ca2+, 2.5 mm; and DAPA, 3.0 FM. The final volume was 1.5 mi, and the buffering medium was 0.02 M HEPES, 0.15 M NaCl, ph 7.4. Reactions were initiated with a small aliquot of concentrated prothrombin;factor Xa, Factor Va, or premixed Factor Va plus Factor Xa, to the remainder of the components in the cuvette used as a reaction vessel. All concentrated components were equilibrated in 2.5 mm Caz+-containing buffer prior to addition, In order to increase the sensitivity of the measurements, an excitation wavelength of 280-nm was used, which both decreased background fluorescence from free DAPA and provided energy transfer from thrombin to DAPA in the DAPA-thrombin complex (52). The sensitivity of the fluorometer was adjusted such that a change in signal corresponding to a full scale deflection on the chart recorder corresponded to conversion of approximately 10% of the avaiiable prothrombin to thrombin. The pre- steady-state interval was defined as the time corresponding to that measured from the origin to point of intersection of a line drawn horizontally from the origin and another indicated by the linear portion of the record of the reaction profile. Time Course of the Assembly of Prothrombime Measured by the Fluorescence Intensity of DEGR-Xa-Bovine DEGR-Xa was prepared from Factor Xa by modification of Factor Xa at the active site with dansyl-glutamyl-glycinyl-arginyl chloromethyl ketone as described previously (16). A 50 nm solution of it (1.5ml final volume) WAS prepared in 0.02 M HEPES, 0.15 M NaCl, ph 7.4, 2.5 mm Ca2+, and 30 &M DMPC/DMPS. The solution was equilibrated at either 2 or 37 "C, and the initial fluorescence intensity (X, = 280 nm, X, = 565 nm) was recorded. A small aliquot of concentrated Factor Va then was added (final concentration 100 nm) and the time course of the change in intensity accompanying the incorporation of DEGR-Xa was recorded continuousiy. The total change in intensity was approximately 18% of the initial value. In efforts to correlate the kinetics of prothrombin activation with prothrombinase assembly, similar solutions were prepared at 2.0 "C containing prothrombin (1.39 p ~ and ) 2.0 nm unmodified Factor Xa (plus 50 nm DEGR-Xa) and DAPA (3.0 p ~ ) Prothrombin. conversion was then initiated by adding an aliquot of concentrated Factor Va (final concentration 100 nm). Prothrombin activation was monitored continuously by the fluorescence intensity of the DAPA-thrombin complex (A,= = 280nM, = 565nM). Instantaneous rates were measured at various intervals throughout the course of the reaction, and corresponding kt values were calculated from the relationship, L=- - rial "I) (4) in which Km was assigned a value of 1.0 pm (1). The concentration of Factor Xa incorporated into the prothrombinase complex was assumed proportional to the measured values of Lt. The total Factor [Xa] concentration was taken as 2.0 nm, which corresponds to the level of active Factor Xa in the reactions. During the course of the reaction the maximum extent of substrate depletion was approximately 30% of the initial value; thus the term in parentheses in the above equation changed only marginally over the interval for which the measurements were made. RESULTS The Ability of Synthetic F ~ s p Vesicles ~ l to ~ Support ~ the Prothr~~bi~~ CompEex-To investigate the influence of membrane fluidity on the activity of the p~thrombin~ complex, we chose phospholipid vesicles composed of synthetic phospholipids which either would or would not undergo a phase transition in the range of 0-37 "C. Table I demonstrates that vesicles composed of highly purified lipids containing defined acyl side chains or of naturally occurring PC and PS can support prothrombinase activity equally well. The TABLE I The effect of DhosDholiDid on the prothrombinase comdkx Relative" rate M X ioe PCIPSb DOPClpOPS' DOPC/POPS/choiesterold DMPCIDMPS" No liuid Relative rate of thrombin production at 25 "C in 0.02 M HEPES, 0.25 M NaCI, ph 7.4, containing 1.39 pht prothrombin, 2 nm Factor Xa, 15 nm Factor Va, 15 PM phospholipid, and 2 mm ea2+. Vesicles composed of 80% phosphatidylcholine from egg and 20% phosphatidylserine from brain. E Vesicles composed of 80% dioleoylphosphatidylcholine and 20% 1-palmitoyl-2-oleoyl phosphatidylserine. Vesicles composed of 60% dioleoylphosphatidylcholine, 20% 1- palmitoyl-2-oleoyl phosphatidylserine, and 20% cholesterol. e Vesicles composed of 80% dimyristoylphosphatidylcholine and 20% dimynstoylphosphatidylsenne. I(,

4 Lipid Mobility and Prothrombinase 3607 decrease in the relative rate of thrombin production seen with the DMPC/DMPS vesicles composed of phospholipids with short saturated acyl side chains is insignificant when compared to the relative rate (0.2) of the reaction in a mixture totally devoid of phospholipid. The apparent Michaelis constants of the reaction are also similar regardless of the composition of the vesicles. Thus, all vesicles studied seemed to supply a suitable surface for complex assembly. When no lipid was added to the reaction mixture (Table I), the Michaelis constant of the reaction increased approximately 5-fold to 5.7 p ~ This. value is significantly lower than the K,,, reported (131 p ~ in ) the presence of Factor Xa alone (53). Presumably, the presence of Factor Va causes the 23- fold decrease in the apparent K, of the reaction. As previously reported by Resnick and Nelsestuen (54), temperature had little or no effect on the binding parameters (Kd and n) of Factor 11 to the various lipid mixtures. Light scattering measurements over a temperature range of 9-48 "C indicated that prothrombin interacts with vesicles of DOPC/ POPS (80:20) and DOPC/POPS/cholesterol (70:20:10) with Kd = 1.1 f 0.3 p~ and n = 180 f 10, and Kd = 1.2 f 0.2 p~ and n = 150 f 10, respectively. When the binding of prothrombin to DMPC/DMPS (80:20) vesicles was monitored, very little interaction could be observed. It was impossible to calculate reliable values for Kd or n, due to aggregation of the vesicles in the presence of Ca2+ at concentrations required for the measurements. The lack of detectable binding under the experimental conditions (60 pm phospholipid, p~ prothrombin) could have resulted from either an increase in the dissociation constant or a decrease in the number of prothrombin molecules bound per vesicle. A decrease in the protein bound at saturation would result in an increase of n which is defined as the number of lipids/protein molecule at saturation. The reason for the difference in the interaction of prothrombin with DMPC/DMPS compared to the vesicles composed of longer chain unsaturated fatty acids (DOPC/ POPS) is unclear. The apparent Michaelis constant (Table I) of the reaction increases only slightly when DMPC/DMPS vesicles are substituted for DOPC/POPS. The vesicles are approximately the same size as determined by quasi-elastic laser light scattering. The diameter of DOPC/POPS and DMPC/DMPS vesicles was 315 and 305 A, respectively. The incorporation of DMPS into the DMPC/DMPS vesicles, however, may not be quan- titative, and these vesicles may actually contain less than 20% DMPS and, thus, support much less binding. The binding of bovine Factor V and Factor Va to DMPC/ DMPS vesicles could not be measured accurately due to aggregation of the vesicles in the presence of Ca2+. The binding to all other vesicle types was also independent of temperature over the range measured. With all vesicle types the apparent Michaelis constant for the prothrombinase reaction remained relatively constant throughout the temperature range studied, whereas the typical alteration in maximum velocity and initial rate which is characteristic of the temperature dependence of enzyme-catalyzed reactions was seen. The Effect of Vesicle Fluidity on the Maximum Initial Rate of Thrombin Production-Various amounts of cholesterol (from 0-20%) were added to DOPC/POPS vesicles to decrease the fluidity of those vesicles. Fig. 1 indicates that the order parameter (S+), which reflects the rotational freedom of the probe embedded in the membrane and thus the fluidity of the membrane, increases with increasing amounts of cholesterol and also with decreasing temperature. When these vesicles were measured for their ability to support the prothrombinase s (+I 0.8 t I I YT O K - ~ X 103 FIG. 1. The temperature dependence of the order parameter of DOPC/POPS vesicles containing various amounts of cholesterol. Vesicleswerecomposed oe 0, DOPC/POPS (80:20); 0, DOPC/POPS/cholesterol (75:205); A, DOPC/POPS/cholesterol (702010); A, DOPC/POPS/cholesterol (602020) '/T OK"X lo3 FIG. 2. The effect of temperature on the initial rate of the prothrombinase reaction with vesicles containing various amounts of cholesterol. The rate of thrombin formation was determined by monitoring the increase in DAPA fluorescence in a solution of 0.02 M HEPES, 0.15 M NaCI, ph 7.4, containing 2 nm Factor Xa, 15 nm Factor Va, 1.39 p~ prothrombin, 2 mm Ca2+, 15 p~ phospholipid, and 3 phi DAPA. Vesicles were composed oe 0, DOPC/POPS (80:20); 0, DOPC/POPS/cholestero1(75:205); A, DOPC/POPS/cholesterol (702010); A, DOPC/POPS/cholesterol (602020). complex at any given temperature, they all gave similar initial rates (Fig. 2) indicating that the 2-fold increase in order parameter had little or no effect on the ultimate rate of catalysis, suggesting that the membrane fluidity over the range studied is not important to the enzymatic expression of the prothrombinase complex. The only alteration seen when the order parameter was increased by the addition of cholesterol to the vesicles or at low temperatures was a lag between the addition of Factor Xa and the time when the maximum initial rate was observed. These lags or pre-steady-state intervals were only seen at low temperatures and with vesicles containing cholesterol. The duration of the pre-steady-state interval appeared to be correlated with higher order parameters and led to the suggestion that the lags may reflect the time required for complex assembly. Less fluid vesicles produced larger lags. To further investigate this phenomenon, vesicles composed of DMPC/DMPS, which exhibit a broad phase transition centered at approximately 25 "C, were studied. The data for

5 3608 Lipid Mobility and Prothrombinase the order parameter (S+), shown in Fig. 3, demonstrate the marked difference in the disorder of the phospholipid preparation at temperatures 525 "C compared to that of the DOPC/ POPS vesicles. Further, the DMPC/DMPS vesicles exhibit a clear phase transition which has a midpoint at approximately 28 "C. In contrast, the order parameter observed for DOPC/ POPS vesicles as temperatures increases, decreases almost monotonically. A phase transition in this temperature range is expected for the DMPC/DMPS vesicles since the phase transition temperature for DMPC is 23 "C and that for DMPS is 38 "C (55). The transition profile is broad in part because the vesicles are small (i.e. there is a high radius of curvature) and in part because there is good mixing of the two types of phospholipid in the bilayers. The minimal temperature-dependent changes in the order parameter of the DOPC/POPS vesicles are also expected since all the measurements were made in a temperature range greater than the phase transition temperatures for both lipids (52). The initial (postlag) rates of prothrombin conversion are compared in Fig. 4a for the two types of vesicles. Both types of vesicles gave parallel results on Arrhenius plots. No marked discontiuity was evident in the region of the phase transition of the DMPCjDMPS vesicles, consistent with the conclusion that membrane fluidity has no dramatic effect on the prothrombinase catalytic reaction per se. The Arrhenius plots from data gathered with all types of lipid mixtures (Figs. 2 and 4, for example) were nonlinear. For reactions carried out in the absence of lipid, the Arrhenius plots were not only nonlinear but had a positive slope at high temperatures, probably due to enzyme inactivation at high temperatures, >30 "C (Fig. 4b). In the presence of lipid, how- ever, the prothrombinase complex appeared stable up 42 to "C. It is possible that this instability may make a contribution to the nonlinearity of the Arrhenius plots higher at temperatures in the presence of lipid. The Pre-steady-state Process (Lag) and Enzyme Assembly- Although the fluidity of the lipid on which the prothrombinase complex is assembled has little effect on the ultimate rate of catalysis of peptide bond cleavage in prothrombin, it does seem to affect the rate of complex assembly. A pre-steadystate interval was only seen with DMPC/DMPS vesicles at temperatures below the phase transition. When the temperature was raised to 37 "C or if DOPC/POPS vesicles were substituted for DMPC/DMPS vesicles at 2 "C, no pre-steady- state interval was observed, regardless of which component was added to initiate the reaction. Order parameter data , I I I I I I 0.2 /"-I i if t-c /-r OK" X lo3 FIG. 3. The temperature dependence of the order parameter of DOPC/POPS and DMPC/DMPS vesicles. Vesicles were composed of: 0, DOPC/POPS (80:20) or 0, DMPC/DMPS (80:20). Log initial rate Log initial rate '/T O K - ~ X lo YT OK-' X lo3 FIG. 4. The effect of temperature on the initial rate of the prothrombinase reaction. The rate of thrombin formation was determined by monitoring the increase in DAPA fluorescence in a solution of 0.02 M HEPES, 0.15 M NaC1, ph 7.4, containing 2 nm Factor Xa, 15 nm Factor Va, 1.39 pm prothrombin, 2 pm Ca2+, 15 p~ phospholipid, and 3 pm DAPA. a, phospholipid vesicles were composed of: 0, DOPC/POPS (80:20); 0, DMPC/DMPS (80:20). b, no phospholipid vesicles were added. indicate that, in contrast to oleoyl-containing the lipids, myristyl-containing lipids underwent a phase transition at -25 "C. Further, this pre-steady-state interval was observed only when the reaction was initiated with either Va, Xa, or a mixture of Va and Xa. These observations are represented in Fig. 5. The vertical axis on the chart in Fig. 5 is fluorescence intensity for thrombin-bound DAPA, whereas the horizontal axis is time, with each increment on the graph paper the equivalent of 5 s. When the reaction was initiated by the addition of prothrombin to equilibrated Factor Va, Factor Xa, phospholipid, and calcium, the reaction proceeded with no lag at a reasonably constant initial rate (Fig. 5A). When the reaction was initiated with Factor Va (panel B), with Factor Xa (panel C), or with a mixture of Factor Va and Factor Xa (panel D), a significant increase in reaction rate with time was observed. This lag, or pre-steady-state interval, could be estimated quantitatively by extrapolating the subsequent quasi-linear portion of the curve to 0 fluorescence intensity. For 5 consecutive experiments performed by the addition of Factor Xa last, a lag value of 46 f 2 s was observed, while for Factor Va (three experiments), a lag value of 47 * 3 s was observed. A similar extrapolation for prothrombin gave an apparent lag of 52.5 s. Data for pre-steady-state intervals for the different lipid systems are summarized in Table 11. In quantitative terms, these numerical values for these lags

6 Lipid Mobility and Prothrombinase c 50 - FIG. 5. Prothrombin activation measured by fluorescence intensity of the DAPA-thrombin complex. Prothrombin activation was measured at 2 "C by fluorescence intensity ( L = 280 nm;, X, = 565 nm). The concentrations of the reactants were: prothrombin, 1.39 p ~ Ca2+, ; 2.5 mm; DAPA, 3.0 pm; DMPC/DMPS, 10 pm; Factor Va, 15 nm; and Factor Xa, 2.0 nm. The reactions were initiated with prothrombin (A), Factor Va (B), Factor Xa (C), or Factor Va + Factor Xa (D). A change in intensity corresponding to a complete deflection of the indicated scale (vertical anis) reflects conversion of approximately 10% of the available prothrombin to thrombin. Each of the divisions indicated on the horizontal axis represents 5-s intervals. When the reaction was initiated with the addition of prothrombin to the other equilibrated components (A), no pre-steady-state interval (lag) was evident, in contrast to reactions initiated with the other indicated components. TABLE I1 methyl ketone-modified Factor Xa (DEGR-FactorXa). Our The influence of various components of the prothrombinase complex laboratory has previously validated DEGR-Factor Xa as a on the pre-steady-state intervals marker for the assembly of Factor Xa into Va-lipid complexes Pre-steady-state interval (16). When DEGR-Factor Xa binds to Factor Va/lipid, Final component changes in both fluorescence polarization and intensity values added" DMPC/DMPS DMPCfDMPS at DOPC/POPS at at 2 "Cb(8020) 37 "c' (8020) 2 "Cd(8020) are observed. We have made use of the latter (intensity) to interpret the influence of nonfluid lipid upon the binding of s Factor Xa. Fig. 6, A and B, represents measurements of the Prothrombin 0" 0 0 rate constants for prothrombin conversion (Fig. 6A) and Factor Xa 46 & fluorescence intensity change of DEGR-Xa with time (Fig. Factor Va 47 -t Factors Xa:Va B). The experiment indicated by Fig. 6A was carried out at 2 "C with 1.39 PM prothrombin, 3.0 PM DAPA, 30 p~ DMPC/ "All other components of the prothrombinase complex were in- DMPS, 50 nm DEGR-Xa (which is inactive), and 2.0 nm cubated together for 15 min at the desired temperature before the unmodified Factor Xa. The reaction was initiated with Factor addition of the final component. *The order parameter for this vesicle type was Va (final concentration 100 nm), and the progress of the The order parameter for this vesicle type was reaction was monitored by the fluorescence intensity of the The order parameter for this vesicle type was DAPA-thrombin complex. The experiment of Fig. 6B was e 0 corresponds to 52.5 s. similarly performed, except prothrombin, DAPA, and unmodified Factor Xa were excluded. Assembly of Factor Xa was monitored by the change in intensity of the included DEGRare not easily interpretable, since they obviously represent Xa, both at 2 (closed circles) and 37 "C (open circles). The complex events; however, from the qualitative standpoint, the closed circles of the upper panel can be compared with those data of Fig. 5 clearly indicate that the lag is dependent upon of the lower panel. This comparison suggests that time-de- Factor Va and Factor Xa addition and not on substrate pendent changes in the rate constant observed below the (prothrombin) addition. Thus, the lag, which occurs for lipids phase transition temperature are consistent within experibelow the phase transition temperature, appears to represent mental error with processes involving the incorporation of some fluidity-dependent process involving enzyme complex DEGR-Factor Xa into the Factor Va-phospholipid complex. (Factor Xa-Factor Va) assembly or expression, rather than Similar incorporation experiments were conducted with substrate (prothrombin) binding. DEGR-Xa using DOPC/DOPS vesicles at 2 "C. For this ex- The simplest interpretation of the lag data of Fig. 5 is that perimental system no lag in fluorescence change associated the nonfluid lipid condition inserts another measurable time- with DEGR-Factor Xa incorporation was observed. dependent rate process in the assembly and expression of the enzymatic Factor Va/Factor Xa complex. To further assess DISCUSSION this interpretation further, fluorescence intensity measure- Phospholipids play a critical role in the prothrombinase ments were conducted using dansyl-glu-gly-arg chloro- complex by providing a surface for the assembly of the en-

7 3610 maximum initial rate of the reaction steadily increases with increasing temperature, showing no obvious break at the phase transition temperature despite the obvious change in lipid order (fluidity) evident from the fluorescence anisotropy data. However, a pre-steady-state interval (lag time) is required prior to the establishment of the steady-state rate. ' I.o ' 15 This lag ~ disappears above the phase transition and most likely 0 I 2 3 Time (min) FIG. 6. The time-dependent change in the rate constant for prothrombin conversion compared to the incorporation of DEGR-Xa into the prothrombin^ complex, Panel A indicates the rate constant (vertical axis) of p~th~mbin conversion with time (horizontal axis) upon initiation of the reaction (2 "C) with Factor Va. Rate constants were estimated from the slopes of the reaction profile of DAPA-thrombin fluorescence measured at the indicated times. Panel B represents the change in fluorescence intensity of DEGR-Factor Xa measured in similar experiments at 2 "C (0) and 37 "C(0). The change in intensity of EDGR-Factor Xa was attributed to assembly of the prothrombinase complex, for which the time required was appreciably greater at 2 "C compared to 37 "C. In addition, the time course of assembly of prothrombinase correlated well withthechange in kat observed in thekinetics of prothrombin activation. zyme, cofactor, and substrate. Although acidic phospholipids are known to be required for the binding interactions (6-E), little else is known about what other characteristics of the phospholipids promote catalysis. Phospholipids containing short chain fatty acids have been reported to show diminished activity when compared to those with long chain fatty acids (11). Other reports suggest that when natural phosphatidylethanolamine is mixed with synthetic saturated phosphatidylserines, the coagulant activity is maximal with DMPS, with an increase or decrease in the hydrocarbon chain length causing decreased activity (19). Hydrogenation of naturally occurring PC and PS decreases procoagulant activity, which led Sterzing and Barton (20) to suggest that optimal procoagulant activity is related to membrane "fluidity." Tan et az. (21) recently tested this hypothesis using synthetic phospholipids. Differential scanning calorimetry was employed to determine the position of the phase transition, and a onestage prothrombinase assay (i.e. clotting of fibrinogen by thrombin produced after Ca" addition) was used to monitor activity of the prothrombinase complex. They concluded that the transformation from the solid to gel phase was accompanied by a sharp increase in the coagulant activity. Our studies have made use of continuous monitoring of thrombin production rather than clotting assays, and make it possible to separate enzyme assembly from expression of enzyme activity. We do not see a dramatic alteration or discontinuity in the expression of enzymatic activity associated with the phase transition of D~PC/DMPS vesicles. The reflects enzyme complex assembly on the lipid surface. Enzyme assembly, rather than enzyme activity, is implicated, since: 1) the lag is only observed for Factor Xa or for Factor Va addition and is not observed for prothrombin; and 2) the change in rate constant associated with the lag is correlated with the rate of Factor Xa binding into the complex. Below the phase transition temperature, complex formation is noticeably slowed. In molecular terms, this may reflect the temperature dependence of the recruitment of lipid molecules from the bulk surface mixture by the proteins as they adsorb to the surface. It is logical to conclude that the recruitment process might be retarded in the gel phase when the lateral diffusion rate of the phospholipid molecule is reduced. The molecular interactions required for the maximal rate of thrombin production include both the assembly of the prothrombinase complex, Step 1. Xa + Va + PCPs F? XaVaPCPS Step 2. XaVaPCPS + I1 * XaVaPCPSII and the hydrolysis of two bonds in prothrombin to yield thrombin, Step 3. XaVaPCPSII + XaVaPCPS f Ha + Fragment 1.2. The rate at which thrombin is produced is dependent upon the pre-steady-state assembly of the catalyst, the binding of substrate, and the steady-state catalytic rate ultimately achieved. We have been able to separate the various processes involved in thrombin generation because our model system is composed of purified protein and lipid components and also because we have analyzed thrombin production continuously. A one-stage assay will not differentiate betweencomplex assembly and the reaction rate obtained with the assembled complex. The observable end point (clot formation) depends on binding and the assembly of the enzyme complex, the catalysis of prothrombin to thrombin, the hydrolysis of a second substrate, fibrinogen, and fibrin monomer polymerization. When Factor Va and Factor Xa are incubated together with lipid and Ca2+ prior to the addition of substrate (prothrombin), no pre-steady-state interval is observedfor nonfluid lipids. Thus the fluidity appears to have little, if any, influence on the ability of prothrombin to present itself as a substrate for the complex. This observation seems to exclude prothrombin binding elsewhere on the vesicle and then moving to the complex by virtue of lipid movement as being a rate-limiting step in the reaction. Under the conditions used in this study, the prothrombinase complex exhibits an apparent K,,, of about 2 PM regardless of the fatty acyl chains in the phospholipid. The apparent K, of the reaction in the absence of lipid (but in the presence of the cofactor, Factor Va) is 5.7 PM (Table I, no lipid). This value reflects greater than a 20-fold decrease in the apparent Km when compared to the value of 131 PM obtained by Rosing et al. (53). The most likely explanation for this observation is an interaction between the Va-Xa complex and prothrombin in solution even in the absence of a lipid surface. Interestingly, the same apparent Km (5-10 PM) was observed with prothrombin species which were isolated from coumarin-treated cows (56) and which contained various numbers of y-carboxyglu-

8 Mobility Lipid and Prothrombinase 3611 tamic residues. These prothrombins cannot bind to acidic phospholipids (56), suggesting that the Factor Va-Factor Xa complex, even in the presence of phospholipid, interacts with prothrombin lacking the phospholipid binding determinants on fragment 1. Unfortunately, the proposed prothrombin- Factor Va interaction is poorly defined. The association may be relatively weak and, thus, difficult to detect. Esmon and co-workers3 found that the heavy chain of Factor Va (the 94,000-dalton subunit) binds to prothrombin immobilized on agarose. Furthermore, they found that when Factor Va was inactivated by activated protein C-mediated hydrolysis of the heavy chain, the heavy chain fragments could not interact with prothrombin. The decreased apparent K,,, of the reaction in the presence of Factor Va, but the absence of lipid, also suggests a Factor Va-prothrombin interaction. By modeling the prothrombinase reaction as occurring in a shell over the surface of the vesicle, Nesheim et al. (57) were able to account for the 2.78 x 105-fold enhancement in the rate of thrombin production which is seen experimentally with the intact complex. Implicit in this model is the importance of the ability of phospholipid vesicles to concentrate the enzyme and substrate at the interface between the lipid vesicle and bulk solution. At physiological concentrations of prothrombin (1.4 p ~), the prothrombin concentration in the interface shell is 1.8 mm, well above the apparent K, of the reaction (57). This leads to the assumption that the apparent K, of the reaction relates to the density of the substrate at the vesicle surface (53, 57, 58). Nelsestuen and co-workers (59, 60) suggest, alternatively, that the phospholipid may provide tighter binding by virtue of the additive binding energies of protein-protein interactions and protein-lipid interactions and that the apparent K, of the reaction is a function of the free prothrombin concentration. Although the experiments described here do not differentiate between the two hypotheses, they do suggest that substrate binding at a site distant to the enzyme-cofactor complex and then moving toward the complex by virtue of the lipid movement cannot be a rate-limiting step in the reaction. The substrate may be concentrated at the vesicle surface by virtue of its lipid binding interactions. Constant prothrombin-lipid dissociation and association would predict that the substrate wouldhave an increased probability of binding to the lipid-enzyme-cofactor complex and that protein-protein interactions may favor the binding of thrombin to the complex as opposed to the lipid vesicle at a distant site. The apparent K,,, observed for the prothrombinase reaction may be the sum of a lipid-binding term which reflects the concentration of prothrombin at the vesicle surface, as well as a term reflecting the influence of Factor Va interactions. The results described here suggest that the fluidity of the lipid surface affects the rate of complex assembly. Once the complex is assembled, however, lipid fluidity has little effect on the rate of thrombin production. Acknowledgments-We wish to thank Sandra Schumann and Jeanne Nemitz for their assistance in the preparation of this manuscript. REFERENCES 1. Nesheim, M. E., Taswell, J. B., and Mann, K. G. (1979) J. Biol. Chem. 254, Davie, E. W., and Fujikawa, K. (1975) Annu. Reu. Biochem. 44, Mann, K. G. (1976) Methods Enzymol. 45, Suttie, J. W., and Jackson, C. M. (1977) Physiol. Reu. 57, Nesheim, M. E., Katzmann, J. A., Tracy, P. B., and Mann, K. G. C. T. Esmon, personal communication. (1981) Methods Enzymol. 80, Paphadjopoulos, D., and Hanahan, D. J. (1964) Biochim. Biophys. Acta 90, Bull, R. K., Jevons, S., and Barton, P. G. (1972) J. Biol. Chem. 247, Stenflo, J., and Suttie, J. W. (1977) Annu. Reu. Biochem. 46, Nelsestuen, G. L., Zytkovicz, T. H., and Howard, J. B. (1974) J. Biol. Chem. 249, Howard, J. B., and Nelsestuen, G. L. (1975) Proc. Natl. Acad. Sci. U. S. A. 72, Subbaiah, P. V., Bajwa, S. S., Smith, C. M., and Hanahan, D. J. (1976) Biochim. Biophys. Acta 444, Bloom, J. W., Nesheim, M. E., and Mann, K. G. (1979) Biochemistry 18, Miletich, J. P., Jackson, C. M., and Majerus, P. W. (1978) J. Biol. Chem. 253, Dahlback, B., and Stenflo, J. (1978) Biochemistry 17, Tracy, P. B., Nesheim, M. E., and Mann, K. G. (1981) J. Biol. Chem. 256, Nesheim, M. E., Kettner, C., Shaw, E., and Mann, K. G. (1981) J. Biol. Chem. 266, Zwaal, R. F. A. (1978) Biochim. Biophys. Acta 515, Colman, R. W. (1976) Prog. Hemostasis Thromb. 3, Grisdale, P. J., and Okary, A. (1965) Can. J. Biochem. 13, Sterzing, P. R., and Barton, P. G. (1973) Chem. Phys. Lipids 10, Tans, G., van Zutphen, H., Comfurius, P., Hemker, H. C., and Zwaal, R. F. A. (1979) Eur. J. Biochem. 95, Nesheim, M.E., Prendergast, F. G., and Mann, K. G. (1979) Biochemistry 18, Nesheim, M. E., Myrmel, K. H., Hibbard, L., and Mann, K. G. (1979) J. Biol. Chem. 254, Bajaj, S. P., and Mann, K. G. (1973) J. Biol. Chem. 248, Downing, M. R., Burkowski, R. J., Clark, M. M., and Mann, K. G. (1975) J. Biol. Chem 250, Weber, K., and Osborn, M. (1969) J. Biol. Chem. 244, Jackson, C. M., Johnson, T. F., and Hanahan, D. J. (1968) Biochemistry 7, Fujikawa, K., Legaz, M. W., anddavie, E. W. (1972) Biochemistry 11, Heldebrant, C. M., Butkowski, R. J., Bajaj, S. P., and Mann, K. G. (1973) J. Biol. Chem. 248, Owen, W. G., Esmon, C. T., and Jackson, C. M. (1974) J. Biol. Chem. 249, Barenholz, Y., Gibbs, D., Litman, B. J., Goll, J., Thompson, T. E., and Carlson, F. D. (1977) Biochemistry 16, Gomori, G. (1942) J. Lab. Clin. Med. 27, Bloomfield, V. A., and Lim, T. K. (1978) Methods Enzymol. 48, Nelsestuen, G. L., and Lim, T. K. (1977) Biochemistry 16, Nesheim, M.E., and Mann, K. G. (1979) J. Bid. Chem. 254, Spencer, R. D., and Weber, G. (1969) Ann. N. Y. Acad. Sci. 158, Spencer, R. D., and Weber, G. (1970) J. Chem. Phys. 52, Mantulin, W. W., and Weber, G. (1977) J. Chem. Phys. 66, Lakowicz, J. R., and Prendergast, F. G. (1978) Biophys. J. 24, Lakowicz, J. R., Prendergast, F. G., and Hogen, D. (1979) Biochemistry 18, Weber, G., and Bablouzian, B. (1966) J. Biol. Chem. 241, Engel, L. W., and Prendergast, F. G. (1981) Biochemistry 20, Jahnig, F. (1979) Proc. Natl. Acad. Sci. U. S. A. 76, Jahnig, F. (1979) J. Chem. Phys. 70, Heyn, M. (1979) FEBS Lett. 108, Lipari, G., and Szabo, A. (1980) Biophys. J. 30, Hare, F. (1983) Biophys. J. 42, Prendergast, F. G., Haugland, R. P., and Callahan, P. J. (1981) Biochemistry 20,

9 3612 Lipid Mobility and Prothrombinase 49. Prendergast, F. G., and Mann, K. G. (1977) J. Bid. Chem. 252, 56. Malhotra, O., Nesheim, M. E,, and Mann, K. G. (1985) J. Bid Chem. 260, J' w.' and K. G' (1978) ''' Nesheim, M. E., Eid, S., and Mann, K. G. (1981) J. Biol. Chem Segel, I. H. (1975) Enzyme Kinetics, pp , Wiley-Intersci- 256, ence, New York 58. Lindhout, T., Grovers-Riemslag, J. W. P., van dewaart, P., 52. Hibbard, L. S., and Mann, K. G., (1980) J. Biol. Chem. 255,638- Hemker, H. C., and Rosing, J. (1982) Biochemistry 21, Rasing, J.7 Tans, G., Govers-Riemslag, J. w. p.9 Zwaal, R. F. A Nelsestuen, G. L. (1980) in Regulation of Coagulation (Mann, K. and Hemker, H. C. (1980) J. Biol. Chern. 255, Resnick, R. M., and Nelsestuen, G. L. (1980) Biochemistry G., and Taylor, F. B., Jr., eds) pp , Elsevier/North- 19, Holland, New York 55. Szoka, F., Jr., and Papahadjopoulos, D. (1980) Annu. Rev. Bio- 60. Pusey, M. L., and Nelsestuen, G. L. (1983) Blochern. Biophys. phys. Bioeng. 9, Res. Comrnun. 114,