Chemistry

and Physics of Lipids, 59 ( 199 I ) I67-

Elsevier Scientific

Publishers

Ireland

I8 I

167

Ltd.

Aggregation behavior of lipid IV* in aqueous solutions physiological pH. 1: Simple buffer solutions Michael Hoferaqb, Randolph

at

Y. Hamptonc, Christian R.H. Raetzc and Hyuk Yub

“Joanneurn Research, Opticul Sensor Institute. Sleyrergasse 17. A-8010 Gru: (Ausfriu). hDepartmenr of Chemistry, University Wisconsin-Madison, Madison. Wisconsin 53706 and “Merck, Sharp and Dohmr, Rahwav. New Jersey 07065 (U.S.A.) (Received

November

We have investigated the aggregation

27th.

1990; revision received March

behaviour

charide) in aqueous solutions in the physiological surface pressure. electron microscopy

of lipid IV,

small unilamellar ..,__ iS wsv

bilayered

Keywords:

^*..A:_” JLUUICS.

endotoxin;

studies. The sonication

range of IO-‘-IO-’ ‘H-NMR

spectra might be attributed

\I,.. I....,^ r_..r”.:..e,.. WC II~VG rsr~rauvsmy

__..^...^A pvpuszu

lipopdysaccharide;

ro: Michael

0009-3084/91/.%03.50 Published and Printed

Hofer,

Steyrergasse

0

Joanneum 17, A-8010

Research, Graz,

1991 Elsevier Scientific

in Ireland

in PBS, Tris and Hepes proThe vesicle size

to a rigid surface structure. This structure

_P...:“.4L” VI wbrxc3

“_ A,..* LIIIUIVI

,.rl.nr wucn

nnn..a,.ntar ca~pc~ca~~ca

..,;+l. *I.w~ku km.

observations.

lipid aggregation

The aim of this work is to clarify the aggregation behaviour of lipid IV, in aqueous solutions in the physiological pH range using a variety of physicochemical techniques. In this paper we report res~ults for buffpr solutions sluch as nhnnnhateI-----l-----buffered saline (PBS), Tris or Hepes. A forthcoming publications will deal with the behaviour of lipid IV* in more physiological systems such as the eukaryotic cell growth medium F-12 in the presence and absence of protein. Lipid WA (also called lipid IA) is a bioactive precursor of lipid A [1], the latter being the lipid anchor of lipopolysaccharide (LPS) on the outer surface of the outer membrane of Gram-negative bacteria. LPS (or endotoxin) is a potent physiological agent in many eukaryotic organisms, causing fever, systemic hypotension and puimonary hypertension [2,3]. Endotoxic shock is fatal Correspondence

” ,.,.*r:e+^_na a CUGAI~LSIIC~

in order to explain some of the experimental

Introduction

tical Sensor Institute,

of lipid IV,

M, possibly even at lower concentrations.

only to the pH and to some extent to the ionic strength. The long time stability of the

vesicles at higher lipid concentrations lipid IV,;

(a bioactive precursor of lipid A and the lipid anchor of lipopolysac-

and force field simulation

vesicles as well as the structureless

_..___..._A L.. .I_^ “:-..t..r:__ suppw LCU uy LIIC ~m~ukz~w~~

1991)

pH range using dynamic light scattering, nuclear magnetic resonance, fluorescence,

duces vesicles which are stable in the concentration is not sensitive to the nature of the buffer.

13th. 1991; accepted June l4th,

of

Op-

Austria.

Publishers

Ireland

in over 50% of patients despite modern medical therapy. An understanding of the molecular mechanism of cell interaction will require knowledge about the physico-chemical behaviour of LPS and related compounds. Unfortunately, the isolation of LPS from bacterl=cllltQ.nlmnrt pm.ria ._ ._lv-.” .. .. ..“. a!Ways in 2 hetPm@fienlJs duct [3]. On the other hand, lipid WA can be isolated from Gram-negative bacteria mutants in milligram amounts of high purity [4,5]. Lipid IVA (and lipid A) displays many of the physiological effects of LPS in both animal and isolated cells [5]. Interestingly, recent work in our and other [6] laboratories indicates that in some species lipid IVA is a potent antagonist of LPS. The potential use of this observation in the development of pharmacological agents based on lipid IV, adds to the need for an understanding of the physical properties of this lipid. Briefly, we review some observations on the physico-chemical behaviour of compounds related to lipid IV* in the literature. Lipka et al. [7] investigated the phase behaviour of lipid X, Ltd.

168

chemically a 2,3-bis (3-hydroxymyristoyl) a-Dglucosamine-l-phosphate. The structure of this lipid corresponds to the reducing end of lipid IVA and would result from hydrolysis of lipid IV* at the bond indicated by an arrow in Fig. 1. Their main finding was that lipid X forms micelles in water at pH L 7 and physiological saline. This is somewhat surprising, since diacyl and dialkyl glycerophospholipids with similar fatty acid chain length usually do not form micelles but bilayers. The reported results on lipid IV* and LPS are not always consistent, possibly due to the heterogeneity or impurities of the isolated product. Mild acid hydrolysed LPS from Salmonella minnesota Re59.5 has been proposed to form a “shed roof-like” structure on the membrane surface with an angle of about 45” between the hydrophilic sugar groups and the hydrocarbon chains [Xl. In soluinverted tion, micelles transforming in thermodynamic equilibrium to the hexagonal H,, phase have been proposed [9]. For LPS from Escherichia coli strain D2 1f’2, non-rigid irregular tubes or ribbons were found to be consistent with neutron scattering data at pH 7.5 [lo]. This type of structure was visualized in electron micrographs and found for the sodium and the Tris salt [ll]. However, a large diversity of structures depending on temperature and ions have been found in this

Oligosaccharide k OH

study. A polymorphic phase behaviour of LPS was also suggested by Coughlin et al. [12] with nonlamellar or hexagonal phases at pH extremes. Materials and Methods Lipid IV* was prepared as described in detail elsewhere [13]. Briefly, a strain of mutant Salmonella typhimurium (STi 50) that accumulates lipid IVA at non-permissive temperatures was used as the lipid source. The bacteria were extracted on the kilogram scale with an organic solvent. Lipid IVA was isolated and purified by sequential ion exchange, silicic acid and in some instances by reverse phase column chromatography with octadecylsilane silica gel (G.T. Baker Chemical Co.) and a two-solvent system [14]. Most of the experiments herein were conducted using lipid IVA that had been through the first two column steps. This is referred to as Biosil lipid IV*. Reverse phase purified and Biosil lipid IV* showed an identical aggregation behaviour as probed by quasi-elastic light scattering (QELS) experiments (see below). The Biosil lipid IV, is approximately 98% pure as assessed by thin layer chromatography and one-dimensional nuclear magnetic resonance (NMR), whereas no impurities could be detected in the reverse phase lipid using these two methods. Lipid IV* was stored as the dry pyridinium salt, desiccated at -20°C. PBS, Tris and Hepes were from Whittaker M.A. Bioproducts (Walkersville, MD), dimyristoyl phosphatidic acid (DMPA) was from Sigma Chemical Company (St. Louis, MO). All were used as received without further purification.

3:I$k$Jo-p_ \p Simulations

HYoH

wyo CH3

,o

Hi-OH

\

FH2

T-H

OH

f=o

O-PLO

(CH,zho 7th

CY

OH

“7””

GH,ho H7-oH CH3

(CH,zho

CH3 Fig. 1. Structure of lipid IV,. then represents the structure

Molecular modeling was performed on a Evans and Sutherland PS 300 with the program package Macromodel@ (version 1.5) using Allinger’s 1986 parameterization for the force field between atoms ]151.

Split at the arrow. of lipid X.

the right half

Surface pressure

Lipid IV* samples were spread on the aqueous subphase (10 mM Tris at pH 7.4) with the spreading solutions made of reverse phase lipid and a binary mixture of chloroform and methanol

169

(3: 1) on a Langmuir trough (28.5 cm x 11.1 cm x 1.0 cm), which is milled out of annealed Teflon, enclosed in a Plexiglas box. The relative humidity of the enclosure was maintained at around 70% with moist filter papers as the vapor wicks. A 1.0 cm x 2.5 cm sandblasted platinum plate attached to a Cahn 2000 electrobalance was used as the Wilhelmy plate. The film was compressed by moving a Teflon barrier at the air/subphase interface at a rate of 1.11 cm*/min. The subphase temperature was controlled to within O.l”C by circulating thermostated water through a glass coil placed at the bottom of the trough and the air within the Plexiglas enclosure by a light bulb. Two independent runs are made to ascertain their reproducibility within 0.1 mN/m. Quasi-elastic light scattering (QELS) Quasi-elastic light scattering was used as the major investigation tool for rapid and accurate assessment of the aggregation behaviour of lipid IV* in all media. It is a fast method to determine diffusion coefficients of lipid aggregates [ 16,171, although with a somewhat limited resolution. The technique has recently been reviewed by an edited monograph of Pecora [ 181. QELS experiments were performed on two different commercial instruments: a Malvern RR 103 selected goniometer with a equipped photomultiplier tube (ITT FW 130) and an argon ion laser (Lexel 7%.2) operating at 488 nm and a Brookhaven BIZOOSM goniometer with a 10 mW He/Ne laser (Melles Griot) operating at 632.8 nm. On both instruments correlation functions were recorded with a 64 channel autocorrelator (Malverm K 7025 and Brookhaven BI 2030, respectively) as a function of scattering angle 19between 30” and 130”. To obtain the decay constants P, which can be related to the hydrodynamic radius, a fit of the measured correlation functions has to be performed. This data treatment, however, was different on the two systems. Since the Brookhaven goniometer was connected on-line to an IBM PC/AT, data analysis was performed with the program package provided with the instrument (cumulant fit, double exponential lit and non-negative constraint inverse Laplace transformation). On the Malvern system,

data were transferred to a VAX 8650 and the correlation functions were fitted using a Marquart type non-linear regression routine with different model functions (single exponential, cumulants, double exponential). Using the same sample in both apparatus, the results obtained by the evaluation procedures were identical within experimental error. The correlation functions were acquired at different scattering angles, and the calculated decay constants P plotted against the square of the scattering vector q* to determine the apparent diffusion coeflicient Dart, (q = 4m A-’ sin (e/2), where n is the refractive index of the scattering medium, 0 the scattering angle and h the wavelength of the incident light). To amplify deviations from the q* dependence of P, we also plot I’lq* vs. q*. In this representation, the data ideally fall on a horizontal line with the intercept being the apparent diffusion coefficient extrapolated to zero scattering angle. The values obtained by the two methods should agree if a freely diffusing, non-interacting system is being investigated. Figure 2 shows a typical example of such plots. We finally convert the apparent diffusion coefficient DaPP to an apparent hydrodynamic radius RHEA,,using the Stokes-Einstein relationship r/q* = Dapp = kT/6a~R,,app

(1) 11000

10000 I

c 7

6000 -

ov. 0

2

q2 x

4

10’l”

* 6

’

8

’

IO0

[cm-2l

Fig. 2. Decay constants r and diffusion coeffkients r/y* as a function of the square of the scattering vector q2. Lines represent least squares fits to the data. D,, evaluated from the slope is 8.76 x IO-’ cm*/s, from the intercept 8.83 x IO-’ cm%. Sample: IO-’ M lipid IV, in PBS at pH 7.4, sonicated for IO min.

170

where 7 is the viscosity of the medium, k the Boltzmann constant and T the absolute temperature of the sample. This conversion is justified for spherical particles only, i.e., for spherical micelles and vesicles. Our studies indicate that this assumption is valid for lipid IV*. In light scattering experiments, reproducibility and accuracy of the measurements are strongly dependent on the purity of the sample. The term “purity” refers to the extent of true scattering due only to the solute. It is therefore important that all samples are prepared in a consistently clean way, especially if polydispersity analysis is the goal. For this reason, all buffers used in QELS experiments were first filtered through 20 nm Anotop filters (Anotec Separations, New York, NY). We have adopted the following procedure for the preparation of the lipid samples. Lipid IVA powder (the pyridinium salt) was placed into Pyrex test tubes prerinsed with filtered buffer. Then buffer was added to achieve the desired concentration, the tube was sealed and sonicated in a bath type sonicator (Lab Supplies, Hicksville, NY). The concentration of this stock solution was usually about l-3 mM lipid IV,. Final concentrations were made by dilution of the stock solution into filtered buffer in precleaned Pyrex tubes. For each measurement, about 1 ml was needed for the light scattering experiment, the other 2 ml were used to rinse a precleaned quartz cell (Wilmad, NY). Ail samples were prepared 30 min before the experiment and contained l-5 mM EDTA. The addition of EDTA to the buffers has two effects: first, it chelates divalent ions which might introduce fusion and/or precipitation of the structures present in solution [19]; second, it inhibits bacterial growth rather effectively. A separately prepared control solution of lipid IV, was always measured with each set of experiments in order to ensure proper dispersion of the lipid by the sonicator. The concentration of lipid and the duration of sonication used for this “standard” were: c = 10e5 M lipid IV, in PBS at pH 7.4, sonicated for 10 min. Fluorescence

The purpose of the fluorescence experiment was to determine the critical concentration of lipid aggregates in aqueous medium. The IVA

fluorescence measurements were made on an Aminco SLM 4800 fluorometer with Rhodamine B in water as the reference sample. All spectra were recorded with 4 nm slit widths on all slits in the range of 560-750 nm with an integration time of 1 s per step. A polarizer in front of the detector optics was used to measure vertically polarized light only. We used Nile Red (Molecular Probes, Eugene, OR) to label the hydrophobic interior of the aggregates in water. This dye has already been used to study phosphatidylcholine vesicles [20]. It shows strong, environment-dependent blue shift, high quantum yield and low fluorescence in water. A detailed analysis of the spectral properties together with some applications can be found in Refs. 20 and 21. The samples were prepared in the following way. A stock solution of lipid IVA which had been sonicated for 20 min was diluted into an ethanol cleaned, dry quartz cell. A second cell was tilled with the solvent (PBS). A known amount of Nile Red in acetone (usually about l-3 ~1 to 2 ml sample volume; final concentration of Nile Red: - 1 nM) was added to both samples which were then heated for 10 min at 40-45°C in the dark in order to saturate the lipid phase with the dye. The spectra of both were taken immediately after, without prior exposure of the dye to light. Finally, we subtracted the solvent spectra from the solution spectra in order to eliminate any effect of dye adsorption on the quartz cell walls. Nuclear magnetic resonance (NMR)

The experiments where performed on a Bruker 500 MHz instrument in the Fourier transform mode. All phosphorus spectra were proton decoupled and recorded in a 1 cm tube with lipid IVA dissolved in 10 mM Hepes buffer at pH 7.4 and a small amount of D20 added to lock on the deuterium nucleus. The lipid concentration was always about 2 mg/ml, the lowest concentration that gave acceptable signal-to-noise ratios for the “P spectra in an overnight experiment. The proton spectra were measured in 10 mM phosphate buffer at pH 7.4 using D,O (100.0 atom%. Aldrich, Milwaukee, WI) as the solvent, saturating the HDO line at about 4~63 ppm. Typical acquisi-

171

tion times for proton spectra were of the order of 30 min and 12-15 h for the phosphorus spectra. The solutions used for the NMR experiments were first examined by QELS to verify proper dispersion of the lipid. Electron microscopy Electron microscopy is widely used for studying the aggregation of lipids in aqueous systems [22-241. For that purpose, a high voltage transmission electron microscope (HVEM) and the low voltage scanning electron microscope (LVSEM, SM 800, Hitachi) were used. The latter was equipped with a special ultra high resolution device [25]. Typical accelerating voltages used on the HVEM were of the order of 1 x lo6 V and on the LVSEM between 1.5 and 5 kV. Lipid IV* dispersions were spread on carbon coated Formvar grids (for TEM) or on microscope slides (for SEM) and freeze dried. Solutions not containing serum albumin did not spread well, resulting in a non-uniform distribution of the aggregates on either the grid or the glass. However, this did not impede obtaining an estimate of the size distribution we were interested in to compare with QELS measurements. Sucrose was added to the solution (I:5 molar ratio lipid/sucrose) before freezing to stabilize the particles against dehydration [26], although a comparison of samples prepared with and without sucrose did not show significant differences in terms of particle size and shape. Samples of lipid WA were frozen in slush nitrogen, freeze dried at -60°C for 2 days to sublimate the ice and then slowly warmed up to room temperature under vacuum. All samples for the SEM (and several samples for the TEM) were sputter coated with a Pt/Au film of about 4 nm thickness whereas standard samples for the HVEM were used without further treatment. Results

parameters stipulating the aggregation behaviour of lipids in aqueous solutions 127,281, we have simulated the structure of lipid IV* using an energy minimizing computer algorithm. Figure 3 shows the minimized geometry of this simulation with the molecule viewed from above the glucosamines (thick lines), along the long axis of the acyl chains (thin lines), which extend to the back of the figure. Hydrogens of the fatty acid chains and the sugars have been omitted for clarity. Comparing the structure of lipid A proposed in Ref. 8 with the simulated structure of lipid IVA in Fig. 3, one finds clear similarities. Despite the difference in the number of hydrocarbon side chains and the absence of the 3-keto-D-mannooctalosonic acid (KDO) in lipid IV*, the tilt angle of the di-glucosamine head group to the fatty acids remains almost unchanged (43” found vs. 45” in Ref. 8). The hydrocarbon chains are almost parallel, although not quite, giving the molecule somewhat the form of a “four-barrel derringer”. It is hence possible that lipid IVA also forms a “shed roof-like” structure on the surface of a bilayer, leading to a rigid exterior through strong interac-

Fig. 3. Simulated three-dimensional simulation

was done on a E&S

Macromodel

(version

The

I .5) with Allinger’s

1986 parameteriza-

tion. The lipid is viewed from the glucosamine

top (thick lines.

45” inclined

the acyl chains

to the plane of representation),

(thin lines) extend backwards.

Conformation simulation studies The chemical structure of lipid IV* is shown in Fig. 1. This type of projection does not, however, reveal the three-dimensional configuration of the lipid. Since packing constraints are important

structure of lipid IV,.

PS 300 using the program

representation.

They

are in the all-/runs

show a slight curvature groups rotate Hydrogens

almost normal

to the plane of

conformation

more or less freely

in the simulation

studies.

of the acyl chains (included in the calculation)

been omitted

for clarity

packing condition”

and

along the chains. The two phosphate

but the picture

have

represents a “closed

of the fatty acids chains.

172

tion of the sugars by hydrogen bonding. However, it must be remembered that the simulations were done without solvent in vacuum at 0 K, i.e. in the absence of any thermal energy and may therefore not reflect the minimum energy structure of lipid IVA on the membrane surface or in aqueous aggregates. Monolayer properties

We report the experimental results for monolayer behaviour of lipid IV* on the air-water interface. The point of the experiment is to determine the molecular projection area/molecule. The minimum area per molecule at the air/water interface can in turn be used for an estimate of the critical packing parameter [27]. Figure 4 shows the surface pressure @)/area ( A2)diagram measured at a constant temperature (isotherm) with a subphase of 10 mM Tris at pH 7.4. The form of the n/A2diagram is characteristic for a liquidexpanded monolayer. The collapse occurs at a surface pressure of 50 mN/m, the corresponding area per molecule is 92 A2. Lipid X shows similar behaviour at pH = 7.4 [7] with a limiting area/molecule of 52 A’, but not lipid A, for which a liquid and a solid-analogous phase have clearly been observed [29] with a limiting area/molecule of variation of the 130 A2. An important area/molecule with pH, ionic strength and the

nature of the ions present due to different charges on the phosphate group has been reported for lipid X [7]. Dephosphorylation in lipid A also leads to decrease in the limiting area/molecule from 130 A2 to 90 A2191. Aggregation behaviour 1. Apparent hydrodynamic radius. Quasi-elastic

light scattering was used to study the aqueous aggregation behaviour of lipid IVA. The pyridinium salt of lipid IVA is a white powder which does not dissolve in aqueous solutions without additional energy input. Even after 2 days at 37’C, no detectable scattering signal from the subphase could be obtained at low angles. We were able to obtain clear suspensions of lipid IV, after a brief sonication (10 s) with a bath sonicator at room temperature. The dependence of the mean apparent hydrodynamic radius on the duration of sonication is shown in Fig. 5 for freshly prepared (circles) and for frozen and thawed solutions (squares). The open circles in the figure correspond to solutions which have been sonicated at a lipid concentration of 2 mM and diluted to IO PM in the isotonic medium. The tilled circles represent measurements where the lipid has been sonicated only for about IO s at 2 mM, diluted to IO PM, and further sonicated to the indicated time. This was done in order to verify the independence of the 60

A E

50.

.

t

E -

50

.

40-

401 b

-i

30 .i

cr'

205

. ‘. .

!? ; m

F

.

30-

.

p\Lf

I

t?? Q a ii e

l. .

20-

.

IO-

0.

i?

o+..z”n”-.‘-*

60

100

120

10' 0

l*. ‘0.. 140

300

600

sonication l

. 160

area / molecule [A 2l Fig. 4. Surface area per molecule vs. surface pressure isotherm of lipid IV, as measured 30 min after spreading of the lipid dissolved in a chloroform/methanol (3: I) mixture.

900

time

1:

30

[s]

Fig. 5. Mean apparent hydrodynamic radius Ruapp as a function of sonication time for three different lipid IV, preparations: circles correspond to freshly prepared solutions. squares to frozen and thawed ones. Filled circles represent sonication at low5 M lipid. open circles sonication at 2 mM lipid with dilution afterwards. The line is arbitrary but emphasises the rough first order reaction kinetics of the fragmentation process.

173

preparation pathway to the final solutions. In both cases, the aggregates break down rather fast, roughly following a first order reaction kinetics. After 10 min sonication, initially different preparations reach the same apparent hydrodynamic radius within experimental error. The large error bar at the beginning (after 10 s sonication) is a sign of a broad size distribution of the aggregates. Correlation functions have been analyzed with the second-order cumulant fit in order to get information about the mean hydrodynamic radius. This model represents the size distribution of the dispersion with only four parameters. If the initial polydispersity is broad, a four parameter lit cannot describe this distribution adequately resulting in large error bars of the mean hydrodynamic radius. Samples which were frozen, thawed and resubjected to sonication showed similar decrease of the mean RHapp as the fresh samples (open squares). However, the initial mean hydrodynamic radius and the associated polydispersity varied from almost no change after freezing to highly aggregated samples. Usually, thawed samples were cloudy, which indicates an aggregation on the length scale of microns. The data points given by the open squares therefore represent the exception rather than the rule. At constant pH, ionic strength and power setting of the sonicator, the duration of sonication appears as the major variable controlling the size of the aggregates. To explore further the aggregation behaviour of the lipid IVA, we measured the concentration dependence of RHapp. A concentrated sample (2 mM lipid) was sonicated for IO min and then diluted under isotonic condition. Figure 6 shows the result for two different solution conditions: the filled circles represent the samples sonicated at pH 7.4 in PBS and open squares the samples at pH 8 in 10 mM Hepes. We investigated the behaviour in Hepes because it was the buffer used for the “PNMR experiments. Figure 6 shows that in either solvent there exist two flat regimes, although the radii obtained in PBS are consistently higher than the ones in Hepes. The transition between these flat portions takes place around 6-8 x 10e5 M and is not sensitive to pH in the vicinity of pH 7.4. Although the change in Ruapp is rather small, it is reproducible, lies outside of the experimental error

c of lipid [PM] Fig. 6. Mean apparent hydrodynamic tion

of

lipid

concentration.

radius

Filled

RHapp as a func-

squares

represent

measurements in PBS at pH 7.4, open squares measurements in Tris

at pH 8. In both cases the sonication

time was IO min at

2 mM lipid IV,.

dilutions

were made from this stock solution

after sonication.

Lines are guides to the eye. not least squares

tits.

and appears in the same concentration range, IO-100 iM. The large error at 5 PM, still only on the order of a few percent, is due to the low intensity scattered by the highly dilute aggregates, making measurements at angles beyond 60” impossible. This concentration was the lowest for which reliable and reproducible QELS results have been obtained. Thus similarly sized aggregates of lipid IV, can be observed in the low micromolar regions, at widely varying ionic strength. The aggregates were found to be stable at room temperature in terms of their mean hydrodynamic radius in 150 mM PBS for at least several days, although rapid aggregation (within minutes) was observed at temperatures higher than 45°C. The time dependence of the mean hydrodynamic radius at room temperature was difficult to determine by QELS because the samples usually picked up dust before there was a reproducible change in radius. Up to 300 mM ionic strength. no fusion and/or precipitation of the lipid aggregates could be observed within several hours. At 600 mM NaCI. aggregation was rapid, leading to an average hydrodynamic radius of about 400-600 nm within minutes (data not shown). Another important parameter of the system is

174

the pH at which the solutions are prepared, although, as stated in the Introduction, we were mainly interested in the characterization of the aggregation behaviour of the lipid in the physiological pH range. Figure 7 depicts the variation of the mean apparent hydrodynamic radius with pH in the range of 6-9. The data points represented by open squares stand for measurements in PBS and those by filled squares for measurements in Tris. In both cases the ionic strength was 150 mM, the duration of sonication was 10 min and the concentration of the lipid was 10e5 M. At pH 7.4, four separate measurements are shown to illustrate the reproducibility of the preparation; two from the pH series and two from the reference samples, one in Tris, the other one in PBS. Samples at pH = 6 appeared more polydisperse, expressed again through the larger error bars of the second cumulant fit to the QELS decay profiles. We interpret the results to indicate that the mean RHappof the lipid IV, aggregates is almost independent of pH at pH 1 7.4. 2. Critical concentrations. Lipid aggregates in aqueous solution have a thermodynamically well defined critical concentration below which no supramolecular structures are present [27,28]. For

15

’

I

5

6

7

6

9

10

PH Fig. 7. Mean apparent hydrodynamic

radius RHapp as a func-

tion of pH. Open squares correspond

to measurements

mM PBS.

tilled squares to measurements

stock solution Methods,

was prepared as described in Materials

A and

but only sonicated for about IO s, in order to break

up undissolved mMI

in I SO

in 150 mM Tris.

solid lipid. Aliquots

were then diluted

into filtered

of the stock solution buffer

desired pH to a tinal lipid concentration suspensions

were subsequently

solutions

(- I

with the

of 10 PM. The diluted

sonicated for 10 min.

micelles, this concentration, commonly designated as “critical micelle concentration” (CMC), typically lies in the range of 10e3 to 10e5 M. For vesicles, the “critical vesicle concentration” (CVC) is in the region of lo-” to low8 M. In this concentration range most experimental techniques can not detect the presence of aggregates. Therefore, CVCs are in general difficult to determine whereas CMCs can easily be determined by several methods [30]. We have examined lipid IV* using several experimental techniques which are able to distinguish between bilayers and micelles. A frequently used method involves detection of the aggregates by the measurement of the fluorescence intensity enhancement of a dye incorporated in the aggregates (see for example Ref. 31). We have already established the presence of aggregates down to a concentration of about 5 PM by QELS. The fluorescence experiments, using the dye Nile Red, were started at 10 PM lipid IVA to overlap with the light scattering experiments. Aqueous suspensions of the lipid were prepared as described in Materials and Methods. The fluorescence intensity decreases linearly with the lipid concentration at the wavelength of the maximum fluorescence of Nile Red. This position shifted slightly from 624 nm in the 10 PM sample to 618 nm in 100 nM sample. Both values are comparable to the peak position in the spectrum of Nile Red in phosphatidylcholine vesicles [20]. The lowest concentration at which a spectrum could be recorded was 0.1 PM lipid IVA. The spectra are independent of the dye concentration as well as of the amount of acetone used to dissolve the dye. We conclude that the lipid aggregates persist at a total lipid concentration of lo-’ M. 3. Thermal transitions. Nuclear magnetic resonance is a frequently applied technique in the study of lipid aggregation. We have used NMR in two ways to examine the aggregation behaviour of lipid IV,: first, to study the evolution of the spectrum with varying temperature to detect “phase transitions” [32] and second to test for the presence or absence of sequestered aqueous space by quenching experiments with paramagnetic ions such as Mn’+ [33-351. Figure 8 shows the proton spectra of the lipid aggregates at 2YC and at 55°C. The insert depicts

PPm

6

2

0

25 “C

55 “C

Fig 8. ‘H-NMR and “P-NMR spectra of lipid IV,. The two proton spectra at 25°C and 55°C illustrate the “chain melting” of the four fatty acids of the lipid. The insert shows two “P spectra: top, before the addition of I mol% Mn2+ to the lipid sample; bottom. after the addition of EDTA to the Mn"' containing sample. Insert (cl shows the “P spectrum of the lipid sample in the presence of 1 mol% Mn’+ on an expanded scale. Details of the sample preparation are given in the text.

s

lipid WA + 1 molgb Mn2+ + EDTA in excess

176

three “P spectra of lipid IV,: before the addition of 1 mol% Mn2+ (top), after further addition of EDTA (bottom) and in between (insert c). The “P spectra were measured with a 2 mM lipid solution, sonicated for 30 min in 10 mM Hepes at pH 7.4 (about 10,000 acquisitions). DzO (10%) was added for locking purposes but no internal or external phosphoric acid standard was recorded. Therefore, the ppm scale may be slightly off upfield or downfield. The spectrum before the addition of Mn2+ (top) can be completely recovered (bottom) after the addition of EDTA to the lipid solution containing Mn’+ as a broadening agent. Therefore, no significant amount of Mn2+ can be traded in a protected space. The spectrum after the addition of Mn2+ is shown in insert (c). The remaining small peak at about -2 ppm is evidence for the presence of a protected space. The “P spectrum of the lipid in aqueous media contains three distinct lines (possibly four) whereas the spectrum in a chloroform/methanol mixture consists of only two lines corresponding to the two phosphorous in the lipid IVA molecule (data not shown). The line width of the phosphorous resonances in chloroform/methanol (where the lipid is dispersed molecularly) is only slightly smaller than in the aqueous dispersion. It is therefore difficult to use these findings as a conclusive argument in favor of one type of structure. However, increasing radius of the aggregates (shorter sonication time) leads to a broadening of the lines. This is in good agreement with a vesicular nature of the lipid aggregates. The data treatment on the instrument did not allow further refinement of the “P spectra in terms of a linear prediction or maximum entropy analyses [36-381. The ‘H spectra were measured in a solution of 1 mM in lipid, 0.1 mM in EDTA (solvent: PBS at pH = 7.4) and we accumulated about 200 scans. A strong increase of the contribution of the fatty acid protons (0.5-2.8 ppm) is observed when the indicating a “chain temperature increases, melting” of the fatty acids, We have performed additional experiments at various temperatures using TSP (3-(trimethylsilyl) propanesulfonate) as an internal integration standard. Figure 9 compares the integrated intensity of the methyl and methylene protons (except the a-methylene group) of lipid

20

30

40

temperature Fig. 9. Number lipid IV,

of integrated

50

60

70

[“Cl protons (0.5-2.8

(open squares) and DMPA

(WI

ppm) of

I .5 mM

circles) suspensions

sonicated for IO and 30 min at 40” and 55°C. respectively. The solvent was 10 mM 10,000 atom% temperatures

Na3P04.

DzO. adjusted

90 mM

NaCI.

O.lmM

EDTA

in

with DCI to pH 7.4. Transition

as shown by the arrows represent the inflection

points of the curves.

IVA (dotted line) and cr-t_-dimyristol phosphatidic acid (DMPA; dashed line) as a function of temperature in 10 mM sodium phosphate in 100.00 atom% DzO at pH 7.4. Both samples were 1.5 mM in the lipid, 0.1 mM in EDTA, 90 mM in NaCl and contained 0.75% TSP. The EDTA lines are far downfield (3.3-3.8 ppm) and thus do not interfere with the integration of the fatty acid protons. The DMPA was sonicated for 30 min at 55°C lipid IV* 10 min at 40°C. The mean hydrodynamic radius obtained by QELS directly in the 1 cm NMR tube after sonication was 25.4 f 1.5 and 15.4 f 0.6 nm, respectively. The main phase transition of DMPA at pH 7.4 has been reported to be around 50°C depending on the ionic strength of the solution [39]. Our results with DMPA (full circles) are in accordance with this, giving a transition temperature of about 52°C. When the same experiment is performed with lipid IV* (open squares), a transition temperature is also observed, at about 37°C. Lipid transition temperatures are known to differ depending on the radius of curvature of sonicated vesicles; strong curvature leads to a broadening of the transition temperature of about 5°C [40]. Below and above the phase transition, the observed NMR spectrum is mostly flat besides the doublet ascribed to ED-

TA and the lines due to the pyridinium counterion (7.4-8.5 ppm). 4. Morphology. Electron microscopy was used to complement what we have examined so far. Figure 10 shows a scanning electron micrograph of lipid IVA sonicated for 10 min in PBS at pH 7.4 in the presence of a 5 molar excess of sucrose. The picture was taken at an accelerating voltage of 1.5 kV. The distance between the dots on the bottom of the figure is 50 nm. The size distribution appears to be rather homogeneous, and the mean diameter of about 50 nm is in good agreement with the mean hydrodynamic diameter of 41 f 1 run (QELS) taking the coating for the SEM (- 4-5 nm) into account. Figure 11 shows a high voltage electron micrograph of lipid IV,, initially sonicated for 2 min, and allowed to remain at 22°C over 3 weeks at a concentration of IO-’ M. The initial RHapp measured by QELS was 28 f 1.5 nm. After 21 days, the sample appears still rather monodisperse with a mean hydrodynamic radius of 55 f 10 run (determined by QELS), confirmed by this electron micrograph, where one can recognize some multilayered structures.

Fig. IO. Low voltage scanning electron micrograph sion of lipid IV,

in 150 mM

of a suspen-

PBS at pH 7.4. sonicated for IO

min at 10m5 M in the presence of a 5 mol excess of sucrose. freeze dried and coated with Pt/Au. dots on the bottom

The distance between the

of the figure represents 50 nm.

Fig. I I. High voltage electron lipid IV,

in 150 mM

microgrdph

of a suspension of

PBS at pH 7.4. sonicated for IO min at

IO-’ M. The suspension was allowed to anneal for 3 weeks at room temperature

before freeze drying. The bar represents 100

nm.

Discussion

The objective of this work was to elucidate the aggregation behaviour of lipid IV, in simple buffer solutions around pH 7.4. The results of the different experimental techniques are consistent with a vesicular structure of the lipid aggregates. We have shown evidence for these structures in total lipid concentrations from 4 x IO-’ M to 1 x IO-’ M. Electron micrographs show aggregates similar to those in Fig. 10 at concentrations < lo-’ M. Direct evidence for the vesicular structure of the aggregates is provided by the following experimental observations. (1) Fluorescence of the lipophilic dye Nile Red could be detected down to a lipid concentration of 0.1 PM. This concentration lies well below any reported CMC of micelle forming lipids, typically in the range between 10 PM and 10 mM. The observed blue shift of the maximum in the fluorescence intensity of Nile Red is very close to the one found in phosphatidylcholine vesicles [20]. (2) Transmission as well as scanning electron microscopy show round particles with an average diameter of the order of 50 nm (after IO min sonication) in good agreement with the values

determined by QELS upon taking into account the coating for the SEM. After several weeks of annealing, TEM shows sometimes large, multilamellar structures with an average radius of 55 nm, which correlates nicely with the mean hydrodynamic radius of 60 nm found by QELS. (3) The hydrodynamic radius of the aggregates decreases with increasing duration of sonication and levels off at sonication times above 20 min with a value of about 18 nm. This radius is too large to be accounted for by spherical micelles and the observed dependence of the hydrodynamic radius on the sonication time is typical for vesicleforming lipids [4 11. (4) NMR experiments show a “chain melting” in the hydrocarbon region in the temperature range typical for vesicle forming dimyristoyl lipids (20-60°C). whereas micelles (i.e. lipid X) do not show transitions at these temperatures [7]. This transition has also been verified by calorimetry. (5) The increasing size of the aggregates with decreasing pH is in agreement with theoretical predictions for liposomes [27,28]. The surface area per lipid molecule should decrease as the pH decreases, due to the titration of the phosphate charges, leading to a decrease in the repulsive head group interaction. Geometrical packing constraints will therefore force the lipids into bilayer shells of lesser curvature than before, thus resulting in larger vesicles. This has been observed experimentally. The same reasoning applies to increasing salinity, where we have observed similar increase in overall dimensions (data not shown). Studies on the geometric packing of the lipid IV* molecules using the structure of the simulations indicate that bilayers are a reasonable aggregation state for the lipid IV, molecules. Further, comparing the area/molecule as monolayer on the air/water interface of lipid X and lipid IV, (52 and 92 A 2, respectively) and assuming that the hydrocarbon chain volume of lipid lVA with four myristoyl side chains is twice that of lipid X (two myristoyl side chains), the lipid lVA molecule should have a more rectangular shape than the wedge shaped lipid X, and thus be more suited to pack into bilayers. This is consistent with the simulation studies, which show a molecule of rectangular shape with an inclined

l

head group from the side view. The critical packing parameter (volume of the lipid molecule divided by the surface area per molecule times the critical hydrocarbon chain length) of lipid IV* was calculated using the measured area/molecule of 92 A2, a hydrocarbon chain length of 14.4 A (11 carbons) evaluated from the cross section of the bilayer measured in X ray experiments [42] and a volume of 24 A-‘/CHl, a literature value for DMPC [43]. The calculated value of 0.8 falls right into the range for bilayer forming lipids, 0.5- 1.O. Geometric packing constraints dominate the lipid aggregation behaviour mainly in the case of moderate head group structures and/or interactions. The application of this concept to lipid IV, might therefore seem somewhat problematic. For the next stage of our study, we attempted to probe the mechanical rigidity of the lipid vesicles by means of their osmotic responses. We found that the osmotic responses, neither the swelling by a hypotonic osmolarity nor the deswelling by a hypertonic osmolarity, could be elicited as detected by QELS in terms of a variation in the diffusion coefficient of the vesicles. More specifically, the vesicles sonicated in 150 mM PBS for 10 min neither showed any detectable change in Dapp after addition of sucrose (up to 500 mM) or NaCl (up to 300 mM) nor upon dilution of the salinity by water. Such an insensitive osmotic response is to be expected with small unilamellar vesicles (SUV), as has been reported in the literature (441. Briefly, high/low osmolarity in the solution will shrink/swell large unilamellar and multilamellar vesicles, but not SUV [45]; their bilayer is already “stressed“ to a maximum and the work necessary to accomplish swellingshrinking is too high as to be furnished by the osmotic pressure. On the other hand, SUV have a strong bilayer curvature and thus a greater capacity to fuse (461. They usually aggregate and fuse rather rapidly in the order of hours, especially in the presence of divalent ions [47]. However, conflicting conclusions on the fusion of SUV above and below the phase transition temperature are reported [48,49], making it difficult to establish what is to be the expected behaviour. The observed long time stability of our samples at room temperature could be ac-

179

counted for by a very rigid structure in the polar head group and/or strong interactions through hydrogen bonding between head groups. According to our simulation studies and the arguments offered by Labischinski et al. [8], the “shed rooflike” structure would favor strong hydrogen bonding among polar head groups, leading to an almost “frozen” structure of the whole molecule below the transition temperature. This would explain the long term stability of the vesicles at room temperature in spite of the large curvature energy. The low concentration of the lipid in most of the experiments and the high surface charge (3-4 charges per head group) of the vesicles should reinforce the long time stability. We now turn to a set of observations which are not entirely consistent with a vesicular structure for lipid IVA aggregates. (1) The “P spectrum in aqueous solution shows a triplet (possibly quadruplet), whereas the spectrum in chloroform/methanol consists of only two lines, corresponding to the two phosphates in the molecular structure. Since lipid IVA is assumed to be in a molecularly dispersed form in the mixed organic solvent, the additional peaks are specific to its aggregation in water. These peaks could be attributed to the differences in the curvature between the inner and the outer lipid layer which leads to a splitting of the phosphorus resonances. However, the typical shift in such a case is of the order of 0.1-0.02 ppm [50], at least 5 times smaller than the one in Fig. 9 (- 1 ppm). The two main peaks of the 3’P spectrum disappeared after addition of 1 mol% Mn2+, a paramagnetic ion likely to broaden the exterior phosphorus resonances [33] (insert c) in Fig. 8). We found that the spectrum could be completely recovered upon addition of EDTA. We therefore take this observation to infer that the lipid is strongly asymmetrically distributed and concentrated in the outer layer of the vesicle. The asymmetric distribution of lipids in SUV is an established fact (see, for example, Ref. 51). Assuming a distribution of 3O%J70% of the lipid between inner/outer layer [51], it seems possible that the phosphorus resonance of the inner layer is very weak. (2) The proton spectrum of lipid IV, aggregates exhibits almost no structure at room temperature

besides a flat hydrocarbon chain signal. Even above the “phase transition“ of the fatty acid chains no sharp resonances appeared. This is in contrast to other SUV, which show pronounced structure in their proton spectra even below the phase transition [52] but would agree with a very rigid head group structure. (3) In the temperature experiment, the number of protons contributing to the hydrocarbon signal of DMPA and lipid IVA compared to the number of protons present in the fatty acids of these lipids is very different; almost 50 of the 54 protons of the two myristoyl residues of DMPA contribute to the signal whereas only about 62 of the 92 (not considering the cxand P protons) do so in the case of lipid IV*. This would mean that if rigidity is the reason for this behaviour, it must extend well into the hydrocarbon region. (4) The ~~~~~ of the aggregates shows a small, but reproducible increase at values around 6 to 8 x 10m5M lipid. This increase is found in Tris, PBS and Hepes, and is independent of the ionic strength of the medium. The Debye screening length in 150 mM PBS is small (a few nanometers) as is the concentration of the vesicles ( < lo-’ M). This excludes any kind of interparticular interactions, which might enhance the apparent diffusion constant in a QELS experiment to be responsible for this effect. However, error bars are high, especially at the lowest concentrations, making it hard to reach a definite conclusion. One explanation of these observations could be that the lipid forms additional micelles and/or that other types of aggregates (lamellar phases) are formed at high concentrations. The four hydrophobic fatty acid chains impede the lipid to dissolve in the aqueous subphase without additional energy input on a reasonable time scale. However, once in solution, this energy input may disrupt the structures present and kinetically trap the lipid in the vesicular aggregation. Micelles, if they exist, should have a CMC and therefore are unlikely to exist at low concentrations. The decrease of the mean apparent hydrodynamic radius above 6-8 x lo-’ M, a typical CMC for such lipids [7], could be due to the presence of micelles at high concentrations. This model would also tentatively explain the additional peaks in the

180

3’P spectrum and the behaviour upon addition of Mn’+. At the moment, we only have two experimental hints that such a situation might exist. The electron micrographs of a sonication series at high concentration ( 10e3 M) show the presence of aggregates of micellar size together with big structures after a short period of sonication and their progressive disappearance as sonication is continued. QELS on the same sample shows the presence of a peak at about 6-8 nm diameter in the distribution analyses, although this could be attributed to an artifact of the inversion procedure and thus having no clear physical significance. In summary, we can say that the sonication of lipid IV* at pH values around 7.4 in PBS, Tris and Hepes produces bilayer vesicles which are stable at very low concentrations. These vesicles show a relatively long time stability, which could be attributed to the inclined head groups that favor hydrogen bonding between sugar groups thus rigidifying the structure. The vesicle size is not sensitive to the nature of the buffer, only to the ionic strength and to the pH below - 7. A pH shift to values around 6 has more dramatic consequences on the size of the aggregates than a pH shift to values around 9, which essentially does not change the mean size of the vesicles. We have tentatively proposed a coexistence of micelles and/or other aggregates with vesicles at higher lipid concentrations.

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