FRET between non-substrate probes detects lateral lipid domain formation during phospholipase A2 interfacial catalysis

Archives of Biochemistry and Biophysics 480 (2008) 1–10

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FRET between non-substrate probes detects lateral lipid domain formation during phospholipase A2 interfacial catalysis Alba A. Vallejo, Marta S. Fernández * Department of Biochemistry, Centro de Investigación y de Estudios Avanzados, del I.P.N. (CINVESTAV), P.O. Box 14.740, 07000 México D. F., Mexico

a r t i c l e

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Article history: Received 25 August 2008 and in revised form 19 September 2008 Available online 30 September 2008 Keywords: FRET Lipid domains PLA2 Interfacial activity

a b s t r a c t The probes C12-NBD-FA (12-[(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino)] dodecanoic acid) and C18-R (octadecyl rhodamine B chloride) have been used as donor and acceptor, respectively, in FRET studies on liposomes subjected to pancreatic PLA2 action. Neither of these fluorophores is a substrate for the enzyme but one of them, C12-NBD-FA, is an analog of the fatty acid reaction product. The fluorophores were incorporated into 1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine (DPPC) liposomes and FRET was studied following the fluorescence of the donor, C12-NBD-FA. Working with a molar ratio of acceptor to donor (A/D) of 1, we have found that FRET efficiency (E) decreases during DPPC hydrolysis. After 60 min, the decrease is equivalent to a reduction of more than five times in the effective A/D ratio, as estimated by interpolation in an efficiency vs. A/D reference curve. Using a more complete, empirical approach, the efficiency data, calculated from experiments at variable A/D proportions and constant donor concentration, were fitted by a rectangular hyperbolic function. The parameter K of this function, representing the A/D ratio at half-maximum transfer efficiency, increases more that five times after 60 min hydrolysis. This agrees with the reduction of the effective acceptor density sensed by the donor after hydrolysis, detected by the interpolation procedure. The heterogeneous distribution of acceptor and donor induced by hydrolysis can be attributed to the formation of product domains in the phospholipid membranes and is consistent with the preferential segregation of the donor, which is an analog of the fatty acid reaction product, in those domains. In conclusion, FRET between non-substrate probes detects the heterogeneities generated in phospholipid membranes by PLA2 action. Ó 2008 Elsevier Inc. All rights reserved.

Introduction The phospholipase A2 (PLA2)1 superfamily comprises a wide variety of enzymes that promote the hydrolysis of the sn-2 ester bond of phosphoglycerides. The mammalian PLA2 from group IB belongs to this superfamily [1,2]. Since it is synthesized abundantly in the pancreas, it is known as pancreatic PLA2. However, it is also expressed, to a lesser extent, in different tissues from organs such as esophagus, stomach, small intestine, colon or lung, among others [3,4]. Expression and induction of this secretory enzyme has also been reported in brain as well as in neurons in primary cultures [5]. In addition to its catalytic activity [6], pancreatic PLA2 can also behave as ligand for receptors that control cell proliferation and vascular smooth muscle contraction [7–9]. This duality of functions makes evident

* Corresponding author. Fax: +52 55 5747 3391. E-mail address: [email protected] (M.S. Fernández). 1 Abbreviations: DPPC: 1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine; C12NBD-FA: 12-[(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)]dodecanoic acid; C18-R: octadecyl transfer B chloride; PLA2: phospholipase A2; FRET: fluorescence resonance energy transferl; A: acceptor; D: donor; FD: fluorescence of donor; FDA: fluorescence of donor in the presence of acceptor. 0003-9861/$ - see front matter Ó 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2008.09.015

the critical importance of understanding in detail, the regulatory mechanisms of the enzymatic activity. The pancreatic enzyme has been taken as representative of the secretory PLA2 family and as a biophysical model to elucidate the mechanisms of activation and heterogeneous catalysis at the lipid-water interface [6,10–17]. In these regards, the porcine pancreatic enzyme has been one of the most extensively studied. There is abundant information concerning its molecular structure, biochemical characteristics and mechanism of action [6,12–17]. Nevertheless, the physicochemical changes in the lipid membranes [13,15,16,18–21] and the lateral organization of substrate and products during hydrolysis by this and other secretory phospholipases, are still under investigation. An interesting feature of phospholipase A2 is its high catalytic activity on self-associated phospholipids as compared with monomeric substrates. This phenomenon, called interfacial activation, is extremely dependent on the physicochemical properties of the aggregated substrate [10]. In our laboratory, we have been concerned with studies of the effect of liposome physical state on phospholipase A2 activity [10,12,14]. Sonicated DPPC vesicles in gel state, are hydrolyzed immediately after addition of PLA2 whereas for the substrate in liquid-crystalline phase, the steadystate rate of hydrolysis is preceded by lag periods of very low

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hydrolytic rate and variable duration, depending on the experimental conditions [12,14–17,23]. The modification of the physical properties of lipid bilayers and red cell membranes by the presence of cholesterol also affect their susceptibility to PLA2 hydrolysis, as recently demonstrated by Heiner et al. [24]. Interestingly, Leidy et al. have reported that the interfacial activation is dependent not only on the macroscopic physical state but also on the fine morphological details of the organized substrate such as the heterogeneities associated to the ripple state [25]. The PLA2-mediated hydrolysis of phospholipids generates fatty acids and lysophospholipids, which may affect the properties of the aggregated substrate [26]. Sequestration of the fatty acid product by delipidated bovine serum albumin (BSA) induces a lag phase in the PLA2-catalyzed hydrolysis of gel phase DPPC liposomes [12]. This BSA-induced latency can be reversed by addition of external free fatty acids. Besides, the fatty acids released by hydrolysis are important in the generation of a negative surface electrostatic potential, which makes the interfacial concentration of calcium ions different from the concentration in the bulk aqueous phase [15,16]. The interfacial calcium concentration modulates the duration of the latency phase. Hydrolysis products can also generate other important changes in membranes by undergoing lateral phase segregation [21,22,27–29]. In this respect, it is interesting that a fatty acid product analog, C12-NBD-FA, decreases its fluorescence and acts as monitor of PLA2 interfacial activation. The formation of product domains that include the C12-NBD-FA probe and lead to self-quenching of the monitor may be a possible cause of the decrease in fluorescence [17]. Domains, or membrane lateral heterogeneities, play important roles in many biological processes including the function of enzymes, transporters and channels or the location and redistribution of receptors, among others. These heterogeneities reflect non-random distribution of integral proteins or phase segregation of lipids in localized membrane areas [30,31]. Some membrane heterogeneities may arise from enzymatic action on the bilayer [32]. In this regard, the pancreatic phospholipase A2, as representative of the secretory phospholipase A2 family, can be used to investigate the formation of lipid domains during hydrolysis of lipid bilayers. Phospholipase A2 interfacial activity as well as the physical properties of membranes including the lateral organization of lipids can be studied using fluorescence monitors [14,23,33–37]. In particular, fluorescence resonance energy transfer (FRET) is a highly informative approach to the study of membranes, enzymes and interfacial phenomena [38–40]. Henshaw et al. used FRET between tryptophans of a venom PLA2 and Laurdan, in a study of the effect of products on the susceptibility of dipalmitoylphosphatidylcholine bilayers to the enzymatic action [35]. FRET between donorand acceptor-labeled PLA2, has been applied to detect the possible formation of enzyme oligomers during hydrolysis at the lipidwater interface [41]. Other FRET approaches to PLA2 activity have been concentrated mainly on the use of phospholipid substrate molecules containing both, the donor and the acceptor of a FRET pair such that the fluorescence of the donor, which is attached to one of the acyl chains, is quenched by the acceptor, attached to the other [42–44]. In this way, intramolecular FRET can monitor phospholipase A2 activity since hydrolysis of the double-labeled phospholipid relieves the quenching. Under this experimental design, FRET is affected directly by the catalytic activity of PLA2. In this work, we aim to apply a FRET approach to the detection of lipid bilayer microheterogeneities that may arise as a consequence of phospholipase A2 activity. In particular, we investigate whether the activity of phospholipase A2 on liposomes affects the intermolecular FRET between two non-substrate fluorescent probes inserted in the lipid vesicles. Our study is focused on the change in fluorescence of C12-NBD-FA in the presence of C18-R.

The rationale is that FRET between these two monitors could be perturbed as a result of hydrolysis. There could be no direct effect of the catalytic process on the fluorophores since neither of the two is a substrate for the enzyme. Therefore, any variation in FRET efficiency that might appear during the enzymatic action should be attributed to a secondary effect of the hydrolysis. Materials and methods Materials The synthetic phospholipid 1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine (DPPC) was from Sigma Chemical Company. Its purity, reported as higher than 99%, was verified by thin-layer chromatography using silica gel plates and chloroform/methanol/ acetic acid/water (140:40:16:8 v/v) as developing solvent. No impurities were observed in the plates when illuminated with UV light after being revealed with 0.1% ANS (8-anilino-1-naphthalenesulfonic acid, sodium salt) aqueous solution [45]. Porcine pancreatic phospholipase A2 from Sigma was subjected to reversed-phase high-performance liquid chromatography (RPHPLC) following the procedure of Tomasselli et al. [46]. The fluorescent compounds C12-NBD-FA (12-[(7-nitrobenz-2-oxa-1,3diazol-4-yl)amino)] dodecanoic acid) and C18-R (octadecylrhodamine B chloride) were purchased from Molecular Probes. Spectroscopic grade solvents, analytical grade reagents and double distilled water further purified through a Milli-Q Plus ultrafiltration system, were used throughout. The NaOH solution (about 0.01 N) employed in the pH-stat titrations of the fatty acid released by PLA2 catalysis was prepared by diluting the supernatant of the alkali-saturated solution (100 g NaOH/100 mL). The exact normality of the solution was determined using potassium biphthalate as standard [12–17]. Liposome preparation Liposomes were prepared as described previously [12,14,17,26,37], by sonication and further annealing at 51 °C, i.e., about 10 °C above the gel/liquid crystal phase transition of DPPC. The procedure is described in brief. Appropriate volumes of chloroform/methanol (9:1, v/v) solutions of DPPC and the fluorescent probes at the required proportions, were thoroughly mixed. Thereafter, the organic solvents were evaporated with a nitrogen stream. To remove any trace of retained chloroform, the sample was redissolved in cyclohexane and redried exhaustively with nitrogen. Afterwards, it was resuspended in an adequate volume of 10 mM NaCl, 5 mM CaCl2, 10 mM Tris–HCl, pH 8.0, and sonicated with a 20-kHz B30 Branson sonifier cell disruptor in the pulsed mode at 70% duty cycle. Sonication was followed by 1 h annealing. Unless otherwise noted, the final phospholipid concentration was 0.35 mM. Liposomes destined for the pH-stat experiments were prepared similarly, except that Tris–HCl was omitted from the aqueous solution. Fluorescence studies Fluorescence studies were carried out in a Perkin Elmer Luminescence LS50B Spectrophotometer with slit widths set at 2.5 nm for both, the excitation and emission monochromators. The sample was thermostatted and magnetically stirred and its temperature was recorded continuously using a microprobe thermocouple inserted directly into the cuvette. FRET studies of the time-course of PLA2 hydrolysis, were performed using 0.35 mM DPPC liposomes labeled with 1.75 lM C12-NBD-FA and variable C18-R concentrations to give different C18-R/C12-NBD-FA molar ratios, ranging from 0 to 1.6. C12-NBD-FA fluorescence, in either the presence or

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absence of C18-R, was measured at 531 nm upon excitation at 450 nm, which is about 10 nm blue-shifted from the donor excitation maximum. This setup minimizes the direct excitation of the acceptor as shown by Struck et al. in their pioneering study of FRET between phospholipids labeled with the NBD and rhodamine fluorescent groups [39]. Fixed wavelength data as well as spectra were analyzed by means of the FL Winlab software. Appropriate blanks were employed to correct measurements for any light scattering contribution. All experiments were carried out at 34 °C. pH-stat measurements Time-course of hydrolysis of DPPC liposomes catalyzed by PLA2 were carried out using as substrate 0.35 mM DPPC liposomes either alone or labeled with 1.75 lM C12-NBD-FA and 1.75 or 3.5 lM C18-R. The fatty acids released by hydrolysis were titrated by means of a pH-stat station from Radiometer consisting of a TTT-80 titrator, a PHM84 pHmeter, an ABU-80 automatic burette, and a REC-80 recorder. Titration was performed at pH 8.00 with 0.010 N NaOH [12–17]. Results and discussion Fluorescence of C12-NBD-FA in the presence of C18-R: FRET between non-substrate probes during the interfacial activity of PLA2 The chromophores used in this work are NBD and rhodamine. The significant overlap between the NBD emission and rhodamine excitation spectra has made this set of fluorescent probes one of the most employed FRET pairs in investigations of physicochemical properties of membranes [39,47,48]. In the present work we use the chromophores attached to two different molecules which are not substrates for phospholipase A2. The donor, an analog of the fatty acid reaction product, is C12-NBD-FA (12-[(7-nitrobenz-2-oxa-1,3diazol-4-yl)amino)] dodecanoic acid) and the acceptor, C18-R (octadecyl rhodamine B chloride). Both compounds are lipophilic and suitable to be inserted in membranes. Although the NBD fluorescent group is attached to C12, it tends to ‘‘loop up” to the interface due to its polarity and the flexibility of the acyl chain [49]. Recently, it has been reported that this propensity to ‘‘loop up” to the membrane surface seems to be diminished in the gel phase [50]. Concerning C18-R, the acceptor, it is anchored to the bilayer through its long aliphatic chain which places the rhodamine group, linked to C1, close to the interface [51]. Fig. 1a shows the emission spectra of C12-NBD-FA, C18-R and the pair C12-NBD-FA/C18-R, incorporated into DPPC liposomes. Fluorescence excitation wavelengths have been 450 nm for C12-NBD-FA and for the FRET pair [39] and 560 nm for C18-R. In these experiments, spectra were normalized at 531 nm for C12-NBD-FA and the FRET pair and at 585 nm for C18-R. The emission spectral analysis shows that together, the two probes constitute a FRET pair, with C12-NBD acting as the donor fluorophore (D) and C18-R, as the acceptor (A). It is clear that C12-NBD-FA fluorescence is quenched and C18-R, despite of not being directly excited, fluoresces. This is comparable to the behavior of compounds such as NBD-DPPE and Rho-DPPE [39,47,48] which share similar fluorescent groups. Hydrolysis of DPPC liposomes affects the spectral behavior of the FRET pair. Fig. 1b shows that after 60 min of PLA2-catalyzed hydrolysis of DPPC liposomes, there is partial dequenching of C12-NBD-FA fluorescence. Comparison of the spectra, which were normalized at the acceptor emission maximum (585 nm), shows an increase in intensity at the C12-NBD-FA emission wavelength (531 nm) upon phospholipid hydrolysis. The partial reversal of FRET detected by the spectral analysis presented in this figure, needs to be quantified and measured kinetically. To this purpose a procedure at fixed wavelengths is described below.

Fig. 1. (a) Emission spectra of C12-NBD-FA (– – – –), C18-R () and the pair C12) in DPPC liposomes. The fluorophores constitute a FRET NBD-FA/C18-R ( pair with C12-NBD-FA acting as donor (D) and C18-R, as acceptor (A). Phospholipid concentration was 0.35 mM. Liposomes were labeled with either 1.75 lM C12-NBDFA (– – – –), 1.75 lM C18-R () or 1.75 lM C12-NBD-FA plus 1.75 lM C18-R ) to yield an acceptor to donor molar ratio (A/D) of 1. Fluorescence ( excitation wavelengths were 450 nm for C12-NBD-FA and the FRET pair and 560 nm for C18-R. Normalization of spectra was done at 531 nm for C12-NBD-FA and the FRET pair, and at 585 nm for C18-R. In the figure, the spectrum of the latter fluorophore was artificially shifted up in wavelength by 2 nm to facilitate its visualization. (b) FRET spectral change upon PLA2-catalyzed hydrolysis of DPPC liposomes labeled with the pair C12-NBD-FA (donor) and C18-R (acceptor). Emission ) and after 60 min hydrolysis (----) of spectra were recorded before ( 0.35 mM DPPC liposomes labeled with C12-NBD-FA and C18-R at 1.75 lM of each probe, to give an A/D ratio of 1. Liposomes were prepared as described above. PLA2 concentration was 2.5 lg/mL. C12-NBD-FA fluorescence (emission maximum 531 nm) was excited at 450 nm, which is about 10 nm blue-shifted from the donor excitation maximum. This setup minimizes the direct excitation of the acceptor [39]. Spectra were run at 400 nm/min and normalized at C18-R emission maximum (585 nm). T = 34 °C.

One of the procedures that has been employed repeatedly to quantify FRET is based on determining the quenching of donor fluorescence by the acceptor [38,39] as described by Eq. (1), which is presented later on in this paper. In this procedure, all fluorescence measurements have to be performed at the excitation and emission wavelengths corresponding to C12-NBD-FA, the donor.

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The recordings of kinetic data during PLA2 hydrolysis have to be done on separate samples of DPPC liposomes either labeled with donor alone or with donor and acceptor or unlabeled, to be used as a blank for light scattering corrections. Thereafter, the degree of quenching and the FRET efficiency are calculated employing data obtained exactly at the same time of hydrolysis for donor alone and donor in the presence of acceptor, according to Eq. (1). In the following we present the experiments to obtain the data necessary for computing FRET efficiency. Fig. 2 depicts the behavior of C12-NBD-FA fluorescence, in the absence or presence of acceptor, during the hydrolysis of DPPC liposomes. The study was carried out at different acceptor to donor ratios keeping C12-NBD-FA concentration constant. The figure shows that the fluorescence of C12-NBD-FA decreases during PLA2 interfacial activity as reported in a previous publication of this laboratory [17]. In that paper we discussed the possibility that C12NBD-FA, which is not a PLA2 substrate but a product analogue, could segregate with hydrolysis products. Another alternative was that the fluorescence decrease could be due to the increase in the polarity of the interface generated by the hydrolysis products [52]. The present work could help clarify this matter. In the figure we can see the effect of C18-R, the acceptor of the FRET pair, on the time-course of the fluorescence change of C12-NBD-FA, the donor in the FRET pair. The fluorescence of C12-NBD-FA is reduced by C18-R and the magnitude of the reduction is different upon hydrolysis of the DPPC liposomes into which the fluorophores are incorporated. It appears that upon hydrolysis of DPPC, the degree of quenching of C12-NBD-FA fluorescence by C18-R is less than in the basal situation prior to adding the enzyme. As discussed above, the observed quenching of C12-NBD-FA fluorescence corresponds to non-radiative energy transfer to C18-R. As a complement to these results, and with the purpose of ruling out any effect of the fluorophores on the enzyme activity, we included a control experiment

Fig. 2. Fluorescence of C12-NBD-FA (donor), in the presence of different proportions of C18-R (acceptor), during hydrolysis of DPPC liposomes catalyzed by PLA2. The reaction was followed by the 531 nm fluorescence of C12-NBD-FA excited at 450 nm. Phospholipid concentration was 0.35 mM. Liposomes were labeled with a single concentration of donor (1.75 lM C12-NBD-FA) and variable concentrations of octadecylrhodamine to yield acceptor to donor molar ratios (A/D) of: 0 (a, s); 0.1 (b, r); 0.4 (c, ); 1.0 (d, e) and 1.6 (e, 4). Results from additional experiments conducted at intermediate A/D ratios fall in between but are omitted to avoid cluttering the figure. The enzyme was added at time zero to a final concentration of 2.5 lg/mL. Temperature was 34 °C. Results shown are representative of six different experiments. Inset: pH-stat titration of the fatty acid released in 10 min by PLA2catalyzed hydrolysis of DPPC liposomes labeled with the FRET probes C12-NBD-FA (D, donor) and C18-R (A, acceptor). Liposomes were prepared at 0.35 mM DPPC (control bar) and labeled with 1.75 lM C12-NBD-FA (bar D) or with 1.75 lM C12NBD-FA together with 1.75, 2.1 or 3.5 lM C18-R to give acceptor to donor molar ratios (A/D) of 1, 1.2 or 2 as indicated below each bar. Liposomes were prepared in 10 mM NaCl, 5 mM CaCl2. The enzyme was added at time zero to a final concentration of 2.5 lg/mL. Titration was performed in 3 mL lipid dispersion at pH 8.00 using 0.010 N NaOH. Bars show the means (±S.D.) of three experiments.

as an inset in the figure. The inset shows the pH-stat titration of the fatty acid released in 42 min by PLA2-catalyzed hydrolysis of DPPC liposomes labeled with the FRET probes C12-NBD-FA and C18-R. Liposomes were prepared from DPPC (control bar) and labeled with C12-NBD-FA alone (bar D) or together with C18-R at variable concentrations to give acceptor to donor molar ratios (A/D) of 1, 1.2 or 2 as indicated below each bar. It can be observed that the pH titration of the released fatty acids is barely affected by C12NBD-FA alone or in combination with C18-R. This lack of effect is seen even at an A/D ratio of 2, which markedly exceeds 1.6, the highest A/D ratio used in the fluorescence studies. Calculation of FRET efficiency From the fluorescence results of Fig. 2 we can calculate the efficiency of FRET between C12-NBD-FA and C18-R as a function of time and for each A/D ratio. We present an example of the details of the calculation in Fig. 3. The figure shows the dependence on hydrolysis time of the fluorescence of donor alone (FD) or in the presence of acceptor (FDA) at an A/D ratio of 1, taken from Fig. 2, as well as the efficiency calculated from both. The efficiency of the fluorescence resonance energy transfer process (E) can be quantified using the following equation [38,39]:

E¼1

FDA FD

ð1Þ

where FD is the fluorescence of the donor, and FDA, the fluorescence of the donor in the presence of the acceptor. In our case, FD and FDA should be the fluorescence of C12-NBD-FA in the absence or presence of C18-R, respectively. We can use this equation for the analysis of the FD and FDA curves of Fig. 3 that were recorded during the time-course of PLA2 action. In this experiment, the FDA curve corresponds to an acceptor to donor ratio of 1. Application of the equation to different times of hydrolysis gives the efficiency curve shown in the same figure. The efficiency decreases as hydrolysis proceeds and levels off after 40 min hydrolysis. The initial efficiency was 0.877 and dropped to 0.51 after 60 min hydrolysis, The rest of

Fig. 3. Effect of PLA2-catalyzed hydrolysis of DPPC liposomes on the fluorescence of ), C12-NBD-FA either alone (FD – – – – –) or in the presence of C18-R (FDA and on the transfer efficiency (E) of the FRET pair formed by the two fluorophores (– – O – –). In the FRET pair, C12-NBD-FA functions as donor (D) and C18-R, as acceptor (A). Phospholipid concentration was 0.35 mM. Liposomes were labeled with 1.75 lM C12-NBD-FA (– – – – –) or 1.75 lM C12-NBD-FA and 1.75 lM C18-R ) to yield an acceptor to donor molar ratio (A/D) of 1. The enzyme was ( added at time zero, to a final concentration of 2.5 lg/mL. The reaction was followed by the 531 nm fluorescence of C12-NBD-FA excited at 450 nm either in the absence (FD) or presence of acceptor (FDA). Temperature was 34 °C. The efficiency (E) at any given hydrolysis time, was calculated by inserting in Eq. (1) the FD and FDA values obtained from the corresponding time-courses of fluorescence depicted in the figure. Results shown are representative of six different experiments.

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the fluorescence curves of Fig. 2, obtained at other A/D ratios can be processed in the same way to obtain the corresponding FRET efficiencies shown in Fig. 4. It is important to note that FRET efficiency between unassociated acceptors and donors uniformly distributed in membranes depends on the acceptor surface density. This has been first described by Fung and Stryer [38] who adapted the Förster theory [53], originally developed for acceptors and donors in bulk solutions, to bidimensional systems. The observed dependence of transfer efficiency on the acceptor surface density was in good agreement with calculations using the spectral overlap integral (J), the quantum yield of the donor in the absence of acceptor, the refractive index of the medium and the dipole–dipole orientation factor (j2). This factor was averaged to 2/3 because for fluorophores inserted in bilayers, the rotational freedom randomizes orientations. The calculated values of R0, the distance between donor and acceptor for 50% transfer efficiency, were consistent with values .derived from experimental transfer efficiencies in liposomes. The conclusion was that the surface density of amphipathic fluorophores evenly distributed in bilayers, can be accurately determined by FRET efficiency measurements. Based on the above demonstration further confirmed by other authors [54], in the following, we analyze our FRET efficiency measurements in terms of the acceptor to donor molar ratios. Since we are working at constant donor concentration, this parameter is directly related to the acceptor surface density. Implications of the reduction of FRET efficiency induced by PLA2catalyzed hydrolysis of liposomes: redistribution of the fluorescent probes as evaluated by two different procedures To evaluate the implications of the reduction in efficiency we followed two procedures. The simplest one is based on the interpolation of the decreased efficiency after hydrolysis in an efficiency vs. A/D reference curve constructed at a single donor concentration and variable proportions of acceptor with data obtained prior to hydrolysis (Fig. 5). Although data might have been fitted by a rectangular hyperbola (see Fig. 6a, below, constructed with the same experimental data of Fig. 5 and fitted by Eq. (2) with

Fig. 4. FRET efficiency of the pair C12-NBD-FA (donor, D) and C18-R (acceptor, A) in DPPC liposomes, at variable acceptor to donor (A/D) ratios, during hydrolysis of DPPC liposomes catalyzed by PLA2. Details of the experimental conditions were described in the legend to Fig. 2. The efficiency (E) at any given hydrolysis time, was calculated as explained for Fig. 4, by inserting in Eq. (1) the fluorescence values of the donor in the absence (FD) and in the presence of acceptor (FDA). These values were obtained from the corresponding time-courses of fluorescence at different A/D ratios, depicted in Fig. 2. Liposomes were labeled with a single concentration of donor and variable concentrations of acceptor to yield A/D molar ratios of: 0 (a, s); 0.1 (b, r); 0.4 (c, h); 1.0 (d, e) and 1.6 (e, 4). Results from additional experiments conducted at intermediate A/D ratios (not shown) fall in between. Data are the means (±S.D.) of six different experiments.

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R2 = 0.9936), in this case the reference curve was drawn without assuming any theoretical model, just by connecting the data points with straight line segments. We observe that efficiency increases with the acceptor to donor ratio, very markedly at first, then more gradually, to finally level off asymptotically for A/D larger than about 1.6. The figure includes an example of the application of this procedure to the experiment of Fig. 3 which was conducted at an analytical A/D ratio of 1, for which the initial efficiency Ei was 0.877 (point i). This efficiency dropped to 0.51 (Et) after 60 min hydrolysis, If the efficiency value after hydrolysis (Et) is interpolated in the reference curve (point t), the resulting A/D value, 0.18, will be the effective ratio for the FRET process after hydrolysis for time t. It should be noticed that the initial A/D ratio can also be considered as an effective ratio. Thus, the effective A/D value drops about 5.6 times, from 1 to 0.18. Since we work at constant donor concentration, this indicates that the effective acceptor concentration sensed by the donor becomes 5.6 times smaller. These changes are an indirect consequence of the enzymatic action, and can be attributed to the lateral reorganization of the substrate and the reaction products [21,22,27–29,55]. To the best of our knowledge, at the moment there is no evidence suggesting that, after the enzymatic action, the orientation factor of Förster theory could be different from the average, randomized value. The interpolation procedure only takes into account the possible redistribution of donor and acceptor after hydrolysis and does not consider other changes that could affect the surroundings of the probes. This oversimplification is implicit in referring the efficiency after hydrolysis, to a calibration curve constructed with data obtained in the absence of hydrolysis such that the system consists in acceptor and donor incorporated into substrate-only liposomes. However, since after hydrolysis liposomes contain products, we

Fig. 5. Transfer efficiency for the FRET pair C12-NBD-FA (donor, D) and C18-R (acceptor, A) in DPPC liposomes, as function of the ratio of acceptor to donor (A/D). The curve, drawn by connecting data points with straight line segments, was constructed with data calculated according to Eq. (1) from donor fluorescence measurements at 531 nm upon excitation at 450 nm. Phospholipid concentration was 0.35 mM. Liposomes were labeled with 1.75 lM C12-NBD-FA and variable concentrations of C18-R to yield the analytical A/D values shown in the abscissa. Temperature was 34 °C. Results are the means (±SD) of six experiments. Points signaled as i and t correspond to the initial (Ei) and final (Et) transfer efficiency values, respectively, determined in Fig. 4. Inset: effective acceptor to donor ratio (A/ D) corresponding to the points i and t for FRET efficiency values before (Ei = 0.877) and after 60 min of enzymatic action (Et = 0.51) as obtained from Fig. 4.

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Fig. 6. FRET efficiency as function of the analytical acceptor to donor ratio (A/D) for the pair C18-R/C12-NBD-FA at different times of PLA2-catalyzed hydrolysis of DPPC liposomes. Hydrolysis times were: (a) t = 0 (control before PLA2 addition); (b) t = 1 min; (c) t = 5 min; (d) t = 10 min; (e) t = 15 min; (f) t = 20 min; (g) t = 30 min; (h) t = 40 min; (i) t = 50 min (not shown); (j) t = 60 min. FRET efficiencies at the aforementioned hydrolysis times, were calculated according to Eq. (1), from the donor fluorescence in the absence or presence of acceptor at several A/D ratios, in experiments similar to those of Figs. 2–4. Data are the means (±S.D.) of six experiments. The lines represent best fits to data of the two- parameter hyperbolic function E = EMX(A/D)/[(K+(A/D)] (Eq. (2)), with EMX set to 1. The value of the corresponding best fit K parameter is shown in each panel together with the standard error of the estimate (K ± S.E.E). In the case of the 50 min hydrolysis curve (curve i, omitted) the value of K is 0.847 ± 0.303. Coefficients of determination (R2) for this nonlinear regression analysis of data are: (a) 0.9936; (b) 0.9974; (c) 0.9942; (d) 0.9463; (e) 0.9277; (f) 0.9281; (g) 0.9092; (h) 0.9129; (i) 0.9277; (j) 0.9414.

also used another approach to carry out the quantitative analysis of FRET under more realistic conditions, as described below. This second procedure consisted in a more complete analysis of the efficiencies (E) and analytical A/D ratios at different times of hydrolysis. Data from curves similar to those of Fig. 4 were employed to obtain the dependence of FRET efficiency on the analytical acceptor to donor ratio (A/D) at several times of hydrolysis as shown in Fig. 6. Following the empirical treatment of Zacharias et al. [56] we fitted our data by a two-parameter hyperbolic equation:

E¼

EMX  ðA=DÞ K þ ðA=DÞ

ð2Þ

In this equation, where the parameter K represents the analytical acceptor to donor ratio necessary to achieve half-maximum transfer efficiency, E tends to zero as A/D approaches zero and levels off at EMX as A/D increases to infinity. We have modeled the data setting EMX to 1 in consonance with Eq. (1), where the complete quenching of donor fluorescence by the acceptor through the energy transfer process would make FDA = 0 and give a maximum efficiency of 1. The lines in the different panels of the figure show the best fitting curves obtained through the non-linear regression analysis of data according to Eq. (2). It can be observed that as the hydrolysis time increases, the curves are displaced toward higher A/D ratios. The displacement corresponds to an increase in the value of the parameter K, the apparent constant that represents the A/D ratio for half-

maximum transfer efficiency. We observe that the longer the hydrolysis time, the larger the value of K. In fact, K increases from 0.169 before hydrolysis, to 0.870 after 60 min of enzymatic action which means that the A/D analytical ratio needed to reach halfmaximum FRET efficiency is 5.15 times higher. Comparison of the results obtained by the two procedures To compare the results obtained by the two procedures we imagine an ideal situation in which a reference curve at time zero of hydrolysis and a curve at time t of hydrolysis, both rectangular hyperbolas, pass exactly through the experimental E vs. A/D data. The curves represent Eq. (2), with EMX set to 1 as in Fig. 6. We can take the curve in panel A at time zero of hydrolysis, characterized by Ki (control before PLA2 addition) and any other curve at time t of hydrolysis characterized by Kt. In general, actual data points almost never lie exactly on a line fitted to data through a theoretical equation. That is also the case of the experiments in Fig. 7. However, what we analyze is an ideal situation in which the points lie exactly on the hyperbolas. The analysis is represented schematically in Fig. 7. The initial point i, which corresponds to the analytical concentration (A/D)i and the efficiency Ei, will be located in the reference curve, constructed with data before hydrolysis (t = 0) and characterized by Ki. The relationship between (A/D)i and Ei, the coordinates of point i will be:

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Fig. 7. Schematic, ideal comparison of the two procedures followed to estimate the change in A/D ratio after hydrolysis of DPPC liposomes from the variacion of the FRET efficiency of the pair C12-NBD-FA (donor, D) and C18-R (acceptor, A). Path I corresponds to the interpolation procedure of Fig. 5 and path II to the complete procedure of Fig. 6. The coordinates of point i are the initial efficiency (Ei) and the analytical concentration (A/D)i. The efficiency decreases to (Et) after hydrolysis for time t > 0. If path I is followed Ei drops to Et along the same hyperbola at time zero and defines the point t and the interpolated value (A/D)eff,t which is the effective ratio of acceptor to donor after hydrolysis. If the second procedure is followed (path II), the efficiency Et drops from the hyperbola at time zero to another hyperbola at time t defining the point t for which the analytical acceptor to donor ratio is (A/D)i. The curves correspond to Eq. (2) (E = EMX.(A/D)/[(K+(A/D)]) withK = Ki for hydrolysis ). As shown in the text, time t = 0 (– – –) and K = Kt for any other time t ( after hydrolysis for time t, Ai/Aeff,t = Kt/Ki (Eqs. (6) and (7)).

Ei ¼

EMX  ðA=DÞi Ki ðA=DÞi

ð3Þ

After the sample has been hydrolyzed for time t, the efficiency decreases to Et. Then, we can follow either the simple interpolation or the complete procedure through paths I or II, respectively. According to the simple procedure (path I), the efficiency Et is inserted in the reference curve and the interpolated A/D value is taken as the effective acceptor to donor ratio (A/D)eff,t. The coordinates of the point t should be related by Eq. (4):

Et ¼

EMX  ðA=DÞeff;t Ki þ ðD=AÞeff;t

ð4Þ

The other possibility is to follow the second, more complete procedure (path II). In that case, the system after hydrolysis would be described by another hyperbola characterized by Kt and containing the point t whose coordinates are (A/D)i and Et, the decreased efficiency after hydrolysis for time t. The relationship between Et and (A/D)i at point t would be:

Et ¼

EMX  ðA=DÞi Kt þ ðA=DÞi

ð5Þ

Since EMX is set to 1, application of simple algebra to Eqs. (4) and (5) gives:

ðA=DÞi Kt ¼ ðA=DÞeff;t Ki

ð6Þ

Considering that in all the experiments at variable A/D ratios only the acceptor concentration was modified while the donor concentration remained constant, Eq. (6) can also be written as:

ðAÞi Kt ¼ ðAÞeff;t Ki

ð7Þ

In Fig. 8 we compare the results from both procedures through the equality of Eq. (6). The figure depicts a plot of [(A/D)i]/ [(A/D)eff,t] vs. Kt/Ki at different analytical A/D molar ratios. The left

7

Fig. 8. Analysis of FRET parameters data according to Eq. (6) by plotting [(A/D)i]/ [(A/D)eff,t] vs. Kt/Ki at different (A/D)i molar ratios. (A/D)i ]/[(A/D)eff,t ratios were calculated from data drawn from the reference curve of Fig. 5 by interpolation of efficiencies obtained from Fig. 3 curves at different times of hydrolysis (t). Kt and Ki at different times of hydrolysis are from Fig. 6. Experimental data were subject to linear regression analysis. Parameter and determination coefficient values for the best fitting straight lines (– – –) are: (A/D)i = 0.1 (d): intercept b0 = 0.8240; slope b1 = 0.095; r2 = 0.8231; (A/D)i = 0.2 (s): b0 = 0.0.4786; b1 = 0.4291; r2 = 0.9410; (A/ D)i = 0.6 (M): b0 = 0.2532; b1 = 0.8738; r2 = 0.9824; (A/D)i = 1 (h): b0 = 0.0414; b1 = 1.099; r2 = 0.9935); (A/D)i = 1.6 (N): b0 = 0.5400; b1 = 0.6617; r2 = 0.9693); (A/ D)i = 2 (not shown): b0 = 1.2390; b1 = 0.3989; r2 = 0.7496. If [(A/D)i]/[(A/D)eff,t] = Kt/ ) with b0 = 0 and b1 = 1. Ki (Eq. (6)) one obtains the ideal, full line (

side of Eq. (6) [(A/D)i]/[(A/D)eff,t], represents the simple, interpolation procedure. For the calculations, the efficiencies taken from curves similar to those of Fig. 4 at several analytical A/D ratios were interpolated in the reference curve of Fig. 5 to obtain the effective A/ D ratio after different times of hydrolysis. The right side of Eq. (6), Kt/Ki, represents the second, more complete, hyperbolic procedure. The values of Ki and Kt for several times of hydrolysis, were taken from the curves of Fig. 6. Data were subjected to linear regression analysis. The values of the parameters and determination coefficients for the best fitting straight lines can be compared with the ideal straight line representing the equality of Eq. (6): [(A/D)i]/ [(A/D)eff,t] = Kt/Ki. From the comparison, it is obvious that the consistency of the simple interpolation procedure with the complete procedure holds for single A/D ratios in the central part of the reference curve, from about 0.6 to 1, but not for values in the extreme segments. At very low or very high A/D ratios, consistency is lost. Fret efficiency parameters and pH-stat titration during PLA2-mediated hydrolysis of DPPC liposomes Fig. 9 illustrates the dependence on hydrolysis time of Kt/Ki and [(A/D)i]/[(A/D)eff,t] (Eq. (6)). It is evident that for A/D = 1, the results of both procedures are close to the equality stated by Eq. (6). Under this condition, Kt/Ki and Ai/Aeff,t increase with hydrolysis time and then, after about 40 min, remain almost constant to reach values of 5.15 and 5.6, respectively, at 60 min hydrolysis. In the case of Kt/Ki, this means that after 60 min hydrolysis, the apparent constant Kt is 5.15 times larger than Ki or that the analytical A/D ratio needed to reach half-maximum efficiency is 5.15 times larger. Thus, the actual concentration of acceptor sensed by the donor should be approx. 19.4% of the initial value. This is in good agreement with the results obtained from the interpolation method, in which, after

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Fig. 9. Dependence on hydrolysis time of the relative increment of Kt with respect to Ki(Kt/Ki) (—h—) and of [(A/D)i]/[(A/D)eff,t] at (A/D)i = 1 (– – j – –) (Eq. (6)). For the calculation of the Kt/Ki ratios the values of the parameter K at different times of hydrolysis were taken from Fig. 6. (A/D)i]/[(A/D)eff,t ratios were calculated from data drawn from the reference curve of Fig. 5 by interpolation of efficiencies obtained from Fig. 4 at different times of hydrolysis (t). Inset: time-course of PLA2-catalyzed hydrolysis of 0.35 mM DPPC liposomes followed by pH-stat titration at pH 8.00. Liposomes were prepared in 10 mM NaCl, 5 mM CaCl2 either in the absence (. . .. . .. . .) or presence (– – –) of the FRET pair (1.75 lM C12-NBD-FA and 1.75 lM C18-R, A/D = 1). The enzyme was added at time zero to a final concentration of 2.5 lg/mL. The pH was kept constant at 8.00 with 0.010 N NaOH; temperature was 34 °C. Each titration was recorded continuously and is representative of three experiments. PLA2 catalysis was studied in 3 mL of 0.35 mM DPPC dispersion such that a consumption of 1.05 lEq of NaOH titrant would represent 100% hydrolysis of the available phospholipid, at the sn-2 ester bond. Thus, the plateau corresponds to approx. 60% hydrolysis.

60 min hydrolysis, Ai/Aeff,t = 5.6 (Eqs. (6) and (7)) and the effective acceptor density in the surroundings of the donor decreases to 18% of the initial value (Fig. 6). Another issue that deserves consideration is the comparison of the temporal behavior of the FRET parameters with the time-course of hydrolysis of DPPC liposomes followed by pH-stat titration. As shown in the inset of the figure, hydrolysis of sonicated DPPC liposomes at 34 °C does not present a latency phase [12–14,57]. Also, labeling of vesicles with the FRET pair C12-NBD-FA/C18-R, does not affect the titration profile. The absence of a latency period has been attributed to different causes including the conformation of the sn-2 acyl chain, which may be favorable for hydrolysis, or the possible presence of defects in the liposomal structure [57,14]. Use of the excimer-forming dipyrenephosphatidyl-choline fluorescent substrate inserted at a very low molar ratio (1/400) in liposomes of a non-hydrolyzable diether analog of DPPC, enabled the study of hydrolysis by PLA2 in virtual absence of released products (<1/400) [14]. Under such condition it was possible to detect significant and immediate hydrolysis after addition of the enzyme at 34 °C in the gel phase but very low hydrolysis at 45 °C, in the liquid-crystalline state. In both cases, there was further activation of the hydrolysis by addition of external products. From Fig. 7 of Ref. [14] it can be estimated that for this early phase of hydrolysis at 34 °C, in virtual absence of products, the reaction rate amounts to about 29% of the rate obtained after addition of 12 mol% external products. These findings explain why the time-course of hydrolysis by pH-stat titration shows immediate activity upon addition of the enzyme (inset of Fig. 9). As for the ratio Kt/Ki, in the absence of heterogeneous distribution of acceptor and donor, it would remain equal to unity. Thus, some accumulation of products must be necessary for the heterogeneities to develop and the uneven distribution of acceptor and donor to take place. In fact, a subtle delay in the rising of Kt/Ki is observed. At first, and during a short period, Kt/Ki rises slowly and then more rapidly to finally level off after increasing more than five times. It is possible that the relatively slow phase of Kt/Ki increment could

correspond to the accumulation of products. This phase is not detected by pH-stat titration since, as discussed above, even in the absence of products, the hydrolytic activity is immediate and quite significant. In contrast, the relative increment of Kt is dependent on some accumulation of products. It is interesting that the maximum value of Kt/Ki is reached only after about 40 min hydrolysis while in the pH-stat curves the plateau is attained in about 10 min. It seems that each method detects different, special features of the events associated to hydrolysis. An additional consideration is that the formation and persistence of domains could be favored by the low diffusion rate of products in the gel phase. The diffusion constant in the gel membranes is about 70 times smaller than in the fluid phase [58]. The molar ratio of phosphatidylcholine between the inner and outer monolayers of liposomes similar to those employed in the present work is approx. 65/35 [59]. The inset to Fig. 9 shows that hydrolysis reaches a plateau at approx. 60% hydrolysis (see figure caption). Thus, it can be suggested that the inner monolayer is not accessible to the enzyme hydrolytic activity. This is in agreement with findings of Jain et al. indicating that the outer monolayer can be hydrolyzed without affecting the liposome integrity [60,61]. The result is also congruent with ultrafiltration experiments showing that pig pancreatic PLA2-mediated hydrolysis does not induce the disruption of liposomes [17]. At 34 °C, the DPPC sonicated liposomes used in this work, are in the gel phase. More precisely, and if the fine morphological details are considered, vesicles are in the neighborhood of the ripple state since the temperature reported for the pretransition is 34 °C [62]. However, values as low as 28 °C have also been reported [63]. Recently, using the generalized polarization spectra slope (GPS) of Laurdan we have constructed a partial phase diagram showing that sonicated DPPC liposomes at 34 °C are in a region of coexistence of phases which extends to temperatures lower than 34 °C. This is consistent with the proposal that a small fraction of La phospholipids coexists with gel phospholipids (Lb0 ) in the ripple state [64]. Therefore, it appears that sonicated DPPC liposomes at 34 °C are either indeed in the ripple state (Pb0 ) or in the middle of the transition between the ripple and gel phase. In any case, there should be a certain fraction of ripples in those vesicles. Ripples can be considered as one-dimensional defects of fluid lipid molecules into a matrix of gel state lipids [63] which could explain the absence of a lag period in the time-course of PLA2-mediated hydrolysis of DPPC liposomes at 34 °C. General discussion Changes in FRET efficiency are indicative of membrane heterogeneities. If the lipid membrane remained homogeneous, efficiency could not be expected to change. Fig. 10 illustrates what would happen had the efficiency not changed upon hydrolysis (panels a and b) as compared with the real behavior (panels c and d). We used Eq. (1) to simulate the time-course of the fluorescence of donor in the presence of various proportions of acceptor in the hypothetical situation in which the efficiency, at any given hydrolysis time, were equal to the efficiency before hydrolysis. Fig. 10a shows that the simulated profiles of fluorescence during hydrolysis look very different from the real ones (Fig. 10c). In the real situation, hydrolysis induces a decreased quenching of the donor fluorescence by the acceptor, in contrast to the simulated scenario, where the degree of donor fluorescence quenching by the acceptor remains constant. Obviously, and by definition, the constancy in efficiency is reflected in a time-invariant parameter K (Fig. 10b). This is in marked contrast with the actual results shown in Fig. 10d in which K becomes more than five times larger after 60 min hydrolysis. As already analyzed above, this increase means that the effective acceptor molar concentration surrounding the donors, has

A.A. Vallejo, M.S. Fernández / Archives of Biochemistry and Biophysics 480 (2008) 1–10

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Fig. 10. Effect of PLA2-catalyzed hydrolysis of DPPC liposomes on donor fluorescence and FRET efficiency of the C12-NBD-FA/C18-R pair. Comparison between simulated data for the hypothetical, homogeneous distribution of the probes (top panels a and b) and real results consistent with non-random distribution of donor and acceptor (bottom panels c and d). Enzyme was added at time zero. (a) Simulated behavior of C12-NBD-FA fluorescence in the presence of variable proportions of C18-R (FDA) if, during hydrolysis, FRET efficiency remained constant at the value obtained before adding the enzyme. Estimations of FDA at a given time (– – – – –) were done according to Eq. (1) (E = 1-FDA/FD) ) and the efficiency obtained before hydrolysis for the A/ by inserting the corresponding, experimental value of FD, the donor fluorescence in the absence of acceptor ( D ratio under consideration (Figs. 2 and 5). The acceptor to donor molar ratios (A/D) from up to down were: 0; 0.1; 0.2; 0.4; 0.6; 0.8; 1.0; 1.2; 1.4 and 1.6. (b) Hypothetical FRET efficiency as function of the A/D ratio representing the speculative assumption that efficiency at any hydrolysis time (t) is equal to the initial efficiency (Et = Ei). The horizontal and vertical dotted lines denote, respectively, the half-maximal efficiency and the parameter K of Eq. (2), representing the A/D value for half-maximal efficiency. (c) Fluorescence of C12-NBD-FA (donor), in the presence of different proportions of C18-R (acceptor), during hydrolysis of DPPC liposomes catalyzed by PLA2. The traces represent results of experiments similar to those shown in Fig. 2 (see its legend for details). The acceptor to donor molar ratios (A/D) from up to down were: 0; 0.1; 0.2; 0.4; 0.6; 0.8; 1.0; 1.2; 1.4 and 1.6. (d) Family of curves of the form E = EMX(A/D)/[(K+(A/D)] (Eq. (2)) showing the relationship between FRET efficiency and acceptor to donor ratio (A/D) at different times of PLA2-catalyzed DPPC hydrolysis. Curves represent best fitting of data shown in each of Fig. 6 panels, by the hyperbolic function described by Eq. (2) with EMX set to 1. The hydrolysis time from up to down is: 0 (control before PLA2 addition); 1 min; 5 min; 10 min; 15 min; 20 min; 30 min; 40 min; 50 min (not shown in Fig. 7); 60 min. Vertical, dotted lines indicate the values of A/D for half-maximal efficiency (which is denoted by the horizontal, dotted line) at time zero (K0 = 0.169) and after 60 min hydrolysis (K60 = 0.870).

been reduced to 18% of the initial value before hydrolysis (Eq. (7)). It can be concluded that the experimental data are far from what would happen if the lipid membrane remained homogeneous during hydrolysis. The non-uniform distribution of the FRET probes is a clear indication of the emergence of lipid membrane heterogeneities. These heterogeneities can be ascribed to the formation of domains by the released products. Since C12-NBD-FA, the donor fluorophore, is an analog of the fatty acid reaction product, it could function as monitor of the behavior of the fatty acids released by the hydrolytic process. Burack et al. [55] have shown that 1-pyrenyldecanoate, another fatty acid analog, cosegregates with products of phosphatidylcholine hydrolysis. Thus, the preferential segregation of C12-NBD-FA with products, besides being consistent with the experimental data, is also quite likely. In the last years, a majority of biochemical, biophysical and cell biology studies on membrane heterogeneities has been devoted to the cholesterol and sphingolipid enriched domains known as rafts. Yet, as remarked by Shaikh and Edidin, ‘‘membranes are not just rafts” [65]. Rafts, however important, are a class among other kinds of heterogeneities that may induce structural perturbations in membranes. It is worth to underscore that biological amphipathic

molecules different from cholesterol or sphingolipids, such as free fatty acids and lysophospholipids, are also capable of phase segregation in membranes. Therefore, opening up the focus of attention to other mixtures of amphipathic compounds which generate ‘‘non-raft” domains might help enrich investigations and discussions on the properties of biological interfaces and their repercussions on the biophysics of membranes. Dynamic heterogeneities may play a role in important membrane functions such as those related to the activation of interfacial enzymes, receptor redistribution, membrane trafficking or signaling, among others [31]. The detection of domains during the interfacial action of phospholipase A2 by means of FRET, suggests the possibility of applying a similar approach to other studies on the dynamic emergence of heterogeneous regions in membranes. Acknowledgments Results of this work are included in the Doctor of Science Thesis of Alba A. Vallejo, to be presented to Cinvestav (IPN). Marta S. Fernández is a member of the National System of Investigators (SNI), México.

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