Characterization of liposomes formed by lipopolysaccharides from Burkholderia cenocepacia,...

13574 Phys. Chem. Chem. Phys., 2010, 12, 13574–13585 This journal is c the Owner Societies 2010

Characterization of liposomes formed by lipopolysaccharides from

Burkholderia cenocepacia, Burkholderia multivorans and Agrobacteriumtumefaciens: from the molecular structure to the aggregate architecture

Gerardino D’Errico,ab

Alba Silipo,cGaetano Mangiapia,

abGiuseppe Vitiello,

ab

Aurel Radulescu,dAntonio Molinaro,

cRosa Lanzetta

cand Luigi Paduano*

ab

Received 30th March 2010, Accepted 12th July 2010

DOI: 10.1039/c0cp00066c

The microstructure of liposomes formed by the lipopolysaccharides (LPS) derived from

Burkholderia cenocepacia ET-12 type strain LMG 16656, Burkholderia multivorans strain

C1576 and Agrobacterium tumefaciens strain TT111 has been investigated by a combined

experimental strategy, including dynamic light scattering (DLS), small-angle neutron scattering

(SANS) and electron paramagnetic resonance (EPR). The results highlight that the LPS molecular

structure determines, through a complex interplay of hydrophobic, steric and electrostatic

interactions, the morphology of the aggregates formed in aqueous medium. All the considered

LPS form liposomes that in most cases present a multilamellar arrangement. The thickness of the

hydrophobic domain of each bilayer and the local ordering of the acyl chains are determined not

only by the molecular structure of the LPS glycolipid portion (lipid A), but also, indirectly, by the

bulkiness of the saccharidic portion. In the case of a long polysaccharidic chain, such as that of

the LPS derived from Burkholderia multivorans, liposomes coexist with elongated micellar

aggregates, whose population decreases if a typical phospholipid, such as dioleoyl

phosphatidylethanolamine (DOPE) is introduced in the liposome formulation. The effect of

temperature has also been considered: for all the considered LPS an extremely smooth transition

of the acyl chain self-organization from a gel to a liquid crystalline phase is detected around

30–35 1C. In the biological context, our results suggest that the rich biodiversity of LPS molecular

structure could be fundamental to finely tune the structure and functional properties of the outer

membrane of Gram negative bacteria.

1. Introduction

Bacteria are unicellular organisms devoid of a membrane-

delimited nucleus and other organelles like mitochondria and

chloroplasts. Depending on their answer to the Gram staining,

they can be divided in two main families: Gram negative

bacteria and Gram positive bacteria. Gram-positive bacteria

are those that are stained dark blue or violet by Gram staining.

This is in contrast to Gram-negative bacteria, which cannot

retain the crystal violet stain, instead taking up the counter-

stain (safranin or fuchsin) and appearing red or pink. The

different responsiveness of the bacteria to the Gram staining

test is due to the differences existing between their cell walls.

In particular, Gram-negative bacteria possess a multilayer

architecture of their cell membrane.1 The internal layer is

quite thin (B3 nm) and is constituted by peptidoglycan, a

polymer consisting of saccharide backbones cross-linked by

peptide side chains, forming a mesh-like local microstructure.

In contrast, the most external shell of these bacteria is formed

by a thicker wall (B7 nm) whose composition is different

among the two sides. The internal side is mainly characterized

by the presence of glycerol-based phospholipids, whereas the

external side is composed of lipopolysaccharides (LPS), large

molecules consisting of a glycolipid moiety and a saccharide

portion covalently linked.2

LPS are also called endotoxins because, once released from the

membrane, they play a key role in the pathogenesis of Gram-

negative infections due to their immunostimulatory properties.

They are indispensable for the bacterial growth and viability and

are responsible for the correct assembly of the membrane and its

structural integrity, being able to protect the bacterial cell from

physical and chemical attacks. From a chemical and bio-

synthetical viewpoint, they are composed of three parts:

—a glycolipid portion known as lipid A. This part is

composed of a disaccharide with multiple fatty acid tails,

embedded in the lipid bilayer while the rest of the LPS projects

toward the external environment;

—a core oligosaccharide portion, covalently attached to the

lipid A. In the core region the 3-deoxy-D-manno-octulosonic

acid residue (Kdo) is always present as well as, in most cases,

heptose residues;

—an external O-specific long polysaccharide chain, also

known as O-chain, that extends from the core oligosaccharide.

Although from a chemical point of view, the O-chain is

essentially similar to the core oligosaccharide, they are

aDepartment of Chemistry, University of Naples ‘‘Federico II’’,Naples, Italy

bCSGI (Consorzio per lo Sviluppo dei Sistemi a Grande Interfase),Florence, Italy

cDepartment of Organic Chemistry and Biochemistry, University ofNaples ‘‘Federico II’’, Naples, Italy

d Institut fur Festkorperforsching Forschungszentrum Julich and JulichCentre for Neutron Science, Garching bei Munchen, Germany

PAPER www.rsc.org/pccp | Physical Chemistry Chemical Physics

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http://dx.doi.org/10.1039/C0CP00066C

This journal is c the Owner Societies 2010 Phys. Chem. Chem. Phys., 2010, 12, 13574–13585 13575

biologically distinct. Indeed, the O-chain is a proper poly-

saccharide with a repeating oligosaccharide unit.

All three parts of LPS can vary among different Gram-

negative bacterial strains; the largest variability is located in

the O-chain portion (that can also be missing) which is more

exposed to selective pressures of the outer environment and to

modifications induced by external stimuli. O-chains are

characterized by high specificity within a species and makes

LPS immunogenic being the antigenic determinant recognized

by the host specific immune system; often even slight

modifications to the primary structure can be carried out to

avoid host detection. Usually, the presence or the absence of

the O-chain determines whether the LPS are considered

smooth or rough type, respectively, depending on the appearance

of the bacterial colonies on agar plates. Full length O-chains

make the LPS smooth type (S-LPS) whereas their absence

makes the LPS rough type (R-LPS, also known as LOS,

lipooligosaccharide).3

The lipid A portion of LPS is the most conserved part,

despite its composition may also vary. Indeed, the lipid A

component is recognized as the primary immunostimulator. It

acts as strong elicitor of the innate immune system by the

induction of inflammatory cytokines in mammalian cells.4

The recognition of bacterial lipid A allows the host to

unequivocally signal the presence of infection. The protein

toll-like receptor 4 (TLR-4) is the signal transducing receptor

of LPS; lipid A is the real effector required to activate TLR-4

signalling pathway in conjunction with a soluble co-receptor

protein lymphocyte antigen 96 (MD-2), which directly and

physically binds to LPS. The low and balanced concentrations

of these mediators and soluble immune response modulators

lead to a resulting inflammation that is one of the most

important and ubiquitous aspects of the immune host defence

against invading microorganisms. Beside these positive effects,

an uncontrolled and massive immune response, due to

the circulation of a large amount of endotoxins, leads to

symptoms of sepsis and septic shock.4

The LPS molecules present on the external surface of the

bacteria are stabilized by the electrostatic interactions existing

between the negative charges located in the lipid A part (1- and

40-phosphates) and in the adjacent Kdo moiety (carboxylate

groups), and the divalent cations present in the aqueous

environment (mainly Ca2+ and Mg2+). These interactions

make LPS layer semirigid with a highly ordered structure

and low permeability towards hydrophobic solutes, thus

explaining the high resistance of Gram-negative bacteria

towards hydrophobic host-released antimicrobial peptides

and negatively charged antibiotics.5

From the arguments summarized above, it appears evident

that many of the relevant aspects of Gram-negative bacteria

behavior can be ascribed to the peculiar physico-chemical

properties of the external layer of their cell wall, whose

architecture is in turn determined by the molecular structure

of the constituting LPS. Consequently, structural and

functional investigations on liposomes or membranes formed

by LPS molecules can greatly help to increase the knowledge

on the stability, low permeability and high resistance of the

outer membrane of Gram negative bacteria and also on their

biological activity. This information may, in turn, lead to new

strategies for the design and synthesis of new molecules

capable of penetrating into the cell wall and causing the

bacterium death. To this aim, several studies have been

performed using several techniques (fluorescence spectro-

scopy, light and electron microscopy, X-ray and neutron

scattering, differential scanning calorimetry, etc.), mainly on

LPS derived by Escherichia coli and Salmonella enterica.6–14

In this scenario, here we present a detailed investigation on

liposomes constituted of three different LPS: (i) R-LPS from

Burkholderia cenocepacia ET-12 type strain LMG 16656;

(ii) S-LPS from Burkholderia multivorans strain C1576;

(iii) R-LPS from Agrobacterium tumefaciens strain TT111.

The results are also compared with those obtained for a

R-LPS derived from S. enterica serotype minnesota strain

595 (Re mutant), presented in a previous work.15 Selection

of LPS is detailed in the Discussion section. Particularly, we

try to connect their self-aggregation behavior to the molecular

structure (shown in Fig. 1). In some samples, a diacylglycero-

phosphoethanolamine has been included in liposomes

formulation. Phosphatidylethanolamines represent, besides

LPS, the most abundant lipid component in the outer

membrane of Gram-negative bacteria.16

The investigation has been performed using an experimental

strategy which has been proved to be extremely informative on

liposome aqueous suspensions15 and combines dynamic light

scattering (DLS) to estimate liposome dimension, small angle

neutron scattering (SANS) to analyze the aggregate morphology

and to estimate the thickness of the lipid bilayer and electron

paramagnetic resonance (EPR) to investigate the dynamics of the

lipid hydrophobic tail in the bilayer. The report is organized as

follows: in the Results (section 3) the results obtained by different

techniques are presented separately; in the Discussion (section 4),

the findings are discussed and compared, in an attempt to achieve

general comprehension of the system behavior.

2. Materials and methods

2.1 Materials and sample preparation

LPS compounds have been extracted and purified as reported

elsewhere.17–19 Liposomes have been prepared by dissolving

an appropriate amount of the molecules in chloroform

(HPLC-grade, obtained from Sigma, St. Louis, MO, USA),

in order to have a final concentration of B1 mg ml�1.

The dissolution has been favored through a slight warming

(B40 1C) and a short sonication treatment (B5 min at 59 kHz).

Subsequently, appropriate amounts of the obtained solution

have been transferred to a round-bottom glass tube. A thin

film of the lipid has been obtained through evaporation of the

solvent with dry nitrogen gas and vacuum desiccation.

For preparing samples including 1,2-dioleyl-sn-glycero-3-

phosphoethanolamine (DOPE, obtained from Avanti Polar

Lipids, Birmingham, AL, USA), a suitable amount of phospho-

lipid dissolved in chloroform (concentration B10 mg ml�1)

has been mixed to the LPS solution in the round-bottom glass

tubes before desiccation. The DOPE/LPS molar ratio has been

set to 25/75, which is close to the LPS composition of the

external leaflet of the outer membrane of the Gram-negative

bacteria.20

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Fig. 1 Molecular structure of LPS from Burkholderia cenocepacia (A, mw = 3500 amu), Burkholderia multivorans (B, mw E 10000 amu),

Agrobacterium tumefaciens (C, mw = 3671 amu) and Salmonella minnesota (D, mw = 2400 amu).

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Samples to be analyzed by EPR also included 1% (w/w) of

spin-labeled phosphatidylcholine (1-palmitoyl-2-[n-(4,4-di-

methyloxazolidine-N-oxyl)]stearoyl-sn-glycero-3-phosphocholine,

n-PCSL, n = 5,14), purchased from Avanti Polar Lipids and

stored at �20 1C in ethanol solutions.

The samples have then been hydrated with different buffer

solutions prepared in D2O (isotopic enrichment >99.8%,

purchased from Sigma), at pH 7.4 (i.e. the physiological pH)

and at pH 9.1 (the pKa of the ammonium groups present onto

the polar head of DOPE molecules) and vortexed. Afterwards,

the obtained suspensions have been sonicated and repeatedly

extruded through polycarbonate membranes of 100 nm pore

size, at least 11 times. The final concentration of LPS is

0.5 mmol dm�3.

Buffer at pH 7.4 has been prepared by dissolving sodium

dihydrogenphosphate NaH2PO4 and sodium hydrogen-

phosphate Na2HPO4 in D2O at concentrations equal to 0.0773

and 0.123 mol dm�3, respectively, whereas buffer at pH 9.1 has

been prepared by dissolving sodium carbonate Na2CO3 and

sodium hydrogencarbonate NaHCO3 in D2O at concentrations

equal to 0.0132 and 0.187 mol dm�3, respectively. The

pH values has been then verified to be within 0.1 pH units

by means of a Radiometer pHM220 pH-meter, equipped with

a saturated calomel electrode and a glass electrode previously

calibrated with the IUPAC standard buffer solutions.21

2.2 DLS measurements

The light scattering setup was an ALV/CGS-3 based compact

goniometer system from ALV-GmbH (Langen, Germany).

The light source was constituted by a He–Ne laser operating

at 632.8 nm with a fixed output power of 22 mW.

In DLS, the intensity autocorrelation function g(2)(t) is

measured and related to the electric field autocorrelation

g(1)(t) by the Siegert relation. The parameter g(1)(t) can be

written as the Laplace transform of the distribution of the

relaxation rate G. Laplace transform was performed using the

RILT algorithm incorporated in the commercial ALV/CGS-3

software.22 From the relaxation rates, the translational

diffusion coefficient D may be obtained as23

D ¼ limq!0

Gq2

ð1Þ

where q = 4pn0/l sin(y/2) is the modulus of the scattering

vector, n0 is the refractive index of the solution, l is the

incident wavelength and y represents the scattering angle.

Thus D is obtained from the limit slope of G as a function

of q2, where G is measured at different scattering angles.

All measurements were performed at (25.00 � 0.05) 1C by

using a thermostat bath.

2.3 SANS measurements

SANS measurements were performed at 25 1C with the KWS2

instrument located at the Heinz Meier Leibnitz Source,

Garching Forschungszentrum (Germany). Neutrons with a

wavelength spread Dl/l r 0.2 were used. A two-dimensional

array detector at three different wavelengths (W)/collimation

(C)/sample-to-detector(D) distance combinations (W7AC8mD2m,

W7AC8mD8m and W19AC8mD8m), measured neutrons scattered

from the samples. These configurations allowed collecting data

in a range of the scattering vector modulus between 0.0019 and

0.179 A�1. The investigated samples were contained in a

closed quartz cell, in order to prevent the solvent evaporation,

and kept under measurements for a period such as to have

B2 million counts of neutrons. The obtained raw data were

then corrected for background and empty cell scattering.

Detector efficiency corrections, radial average and transformation

to absolute scattering cross sections dS/dO were made with a

secondary plexiglass standard.24,25

2.4 EPR measurements

EPR spectra of 5-PCSL or 14-PCSL in LPS samples were

recorded on a Elexys E-500 EPR spectrometer from Bruker

(Rheinstetten, Germany) operating in the X band. Capillaries

containing the samples were placed in a standard 4 mm quartz

sample tube. The temperature of the sample was regulated and

maintained constant during the measurement by blowing

thermostatted nitrogen gas through a quartz Dewar. The

samples were investigated in a temperature range spanning

from 5 to 45 1C. The instrumental settings were as follows:

sweep width, 120 G; resolution, 1024 points; modulation

frequency, 100 kHz; modulation amplitude, 1.0 G; time

constant, 20.5 ms; sweep time, 42 s; incident power, 5.0 mW.

Several scans, typically 16, were accumulated to improve the

signal-to-noise ratio.

3. Results

3.1 DLS measurements: aggregate dimension

The size of aggregates formed by LPS in buffer at pH 7.4, and

LPS-DOPE mixtures at pH 7.4 and 9 have been obtained

through DLS measurements, carried out in the angular range

30–1201. The relaxation time distribution, obtained by regularized

inverse Laplace transformation of the correlation functions,

are shown in Fig. 2–4. For pure LPS (solid lines), they are

monomodal for all the systems except for that constituted by

S-LPS from B. multivorans that showed a bimodal distribution,

indicating the presence of two well-distinct species in solution.

The experimental translational diffusion coefficients, D,

obtained from the relaxation rates as described in the

Experimental section, are reported in Table 1. In the same

table the literature data relative to S. minnesota are also

reported.15

In the approximation of very diluted solutions, D can be

directly related to the hydrodynamic radius of the aggregates,

RH, through the Stokes–Einstein equation, which holds for

infinitely diluted hard spheres diffusing in a continuous medium26

RH ¼kBT

6pZDð2Þ

where kB is the Boltzmann constant, T is the absolute

temperature, and Z is the viscosity of the medium (i.e. the solvent).

RH values of LPS aggregates are also reported in Table 1.

From inspection of the table, it is possible to infer the presence

of large diffusing aggregates with radii ranging between B100

and B180 nm, compatible with the presence of liposomial

aggregates. For the system composed of B. multivorans S-LPS,

smaller aggregates with RH E 25 nm are also present.

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The presence of DOPE in the liposome formulation does

not change the kind of aggregates present in each system at

both pH values investigated, see Fig. 2–4. In the case of S-LPS

derived from B. multivorans, DOPE alters the relative

populations of the two kinds of aggregates, in that the relative

volume fraction of larger aggregates increases.

3.2 SANS measurements: aggregate structural parameters

The structural parameters of the aggregates have been

obtained by applying the appropriate model to the experimental

scattering cross sections.27 Measurements have been performed

for all the systems in the absence and in the presence of DOPE,

at pH 7.4 and 9.1, see Fig. 5–7. In particular, the scattering

cross section pattern dS/dO for the aqueous dispersion of

R-LPS derived from B. cenocepacia and A. tumefaciens are

reported in Fig. 5 and 7, respectively (open circles). Qualitative

analysis of the cross sections allows revealing the presence of a

power law dS/dOp q�a in the small q region (qr 0.005 A�1),

with the exponent a > 2.28 Furthermore, in the intermediate

q region (q D 0.05 A�1) a shoulder is present. All these

features, consistent with the molecular architecture of the

molecules, are typical characteristics of the presence of multi-

lamellar vesicles/liposomes. Indeed, the value of the exponent

a depends on the average lamellarity of the vesicles29 whereas

the shoulder should arise from the interference occurring

among the concentric vesicular layers. Despite the presence

in literature of more complex models for multilamellar

vesicles,29 in order to reduce the number of fitting parameters,

we have used the simplest model30 available for structural

characterization of such aggregates. Accordingly, the scattering

function has been modeled considering the liposomes as a

collection of one-dimensional paracrystalline stack. The

theoretical expression of dS/dO is:

dSdOðqÞ ¼ 1

q2hjf ðqÞj2i 1þ hjf ðqÞj

2ihjf ðqÞj2i

ðSðqÞ � 1Þ !

þ dSdO

� �incoh

ð3Þ

where f(q) is the form factor of a bilayer, whereas S(q) is the

structure factor that takes into account the interferences

occurring among the bilayers belonging to a single stack.

Finally, (dS/dO)incoh represents the incoherent contribution

to the cross section measured, mainly due to the presence of

hydrogenated molecules. Both form and structure factors

depend in a complex way on the geometrical parameters of

the stack, namely the number of layers in the stack, N, the

mean layer thickness, d, and the distance between the centers

of two consecutive layers dl. Thus, optimized values of these

parameters have been obtained from data fitting. Actually, the

Fig. 2 Relaxation time distributions at 25 1C and y = 901 for the

following aqueous systems: LPS from Burkholderia cenocepacia at

pH 7.4 (continuous line); LPS from Burkholderia cenocepacia—DOPE

(3 : 1) at pH 7.4 (dashed line); LPS from Burkholderia cenocepacia—

DOPE (3 : 1) at pH 9 (dotted line).


following aqueous systems: LPS from Burkholderia multivorans at

pH 7.4 (continuous line); LPS from Burkholderia multivorans—DOPE

(3 : 1) at pH 7.4 (dashed line); LPS from Burkholderia multivorans—

DOPE (3 : 1) at pH 9 (dotted line).


following aqueous systems: LPS from Agrobacterium tumefaciens at

pH 7.4 (continuous line); LPS from Agrobacterium tumefaciens—

DOPE (3 : 1) at pH 7.4 (dashed line); LPS from Agrobacterium

tumefaciens—DOPE (3 : 1) at pH 9 (dotted line).

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number of layers N is generally only approximately determined;

it determines the upturn in scattering at low q values where the

total thickness of the stack is seen. Furthermore, because of the

low scattering contrast existing between the highly hydrated LPS

hydrophilic moieties and D2O bulk, the obtained d values

represent the thickness of the hydrophobic layer constituted of

the acyl chains rather than the whole thickness of LPS bilayer.

Both d and dl have been allowed to be polydisperse: the former

with a Schulz–Zimm distribution function related to the Zimm

polydispersity indexZ, the latter with a Gaussian distribution with

a standard deviation sdl. In particular, the ratio sdl/dl, named

Hosemann factor, has been estimated. The values of all the

parameters are collected in Table 2. It is worth pointing out that,

because of the limited instrument resolution, the polydispersity

parameters, Z and sdl/dl tend to be overestimated, since both the

resolution and the polydispersity smear out the oscillations

contained in the form factor. Inter-particle structure factor may

be approximated to the unity because of the low amount of LPS

(0.5 mmol dm�3, corresponding to a volume fraction of D0.07%

of solute volume fraction).

Different experimental evidences have been obtained for the

S-LPS derived from B. multivorans. In this case the scattering

profile obtained for the aqueous dispersion (open circles in

Fig. 6) is typical of two different size scattering objects:31,32 in

the region at high q (q > 0.006 A�1) the cross-section shows a

power law decrease, (dS/dO) p q�1, characteristic of

elongated cylindrical micelles, whose theoretical expression,

in the region where the power law holds, can be written as33

dSdOðqÞ ¼

fcylð1� fcylÞp2R2ðrc � r0Þ2

qexp � q2R2

4

� �

þ dSdO

� �incoh

ð4Þ

Table 1 Translational diffusion coefficients, D, and hydrodynamic radii, RH, obtained through DLS measurements at 25 1C. In the table, LPS areindicated by the name of the bacterium from which they are obtained

System 1012D=m2 s�1 RH=nm

Salmonella minnesotaa 2.7 � 0.9 80 � 20Burkholderia cenocepacia pH 7.4 1.66 � 0.15 146 � 13Burkholderia cenocepacia—DOPE pH 7.4 1.39 � 0.06 175 � 7Burkholderia cenocepacia—DOPE pH 9.1 2.14 � 0.05 113 � 3Burkholderia multivorans pH 7.4 9.59 � 0.15 25.3 � 0.8

1.78 � 0.11 136 � 8Burkholderia multivorans—DOPE pH 7.4 8.75 � 0.18 27.8 � 0.6

1.33 � 0.06 183 � 8Burkholderia multivorans—DOPE pH 9.1 8.8 � 0.2 27.6 � 0.6

1.32 � 0.07 184 � 10Agrobacterium tumefaciens pH 7.4 1.64 � 0.03 148 � 3Agrobacterium tumefaciens—DOPE pH 7.4 1.86 � 0.02 130 � 2Agrobacterium tumefaciens—DOPE pH 9.1 1.64 � 0.04 148 � 4

a Data from ref. 15.

Fig. 5 Scattering cross sections at 25 1C for the following aqueous

systems: LPS from Burkholderia cenocepacia at pH 7.4 (open circles);

LPS from Burkholderia cenocepacia—DOPE (3 : 1) at pH 7.4

(full triangles); LPS from Burkholderia cenocepacia—DOPE (3 : 1) at

pH 9 (open squares). For a better comparison, data have been multiplied

by a scale factor, as indicated. Fitting curves to the experimental data

through the models reported in the text are also shown.


systems: LPS from Burkholderia multivorans at pH 7.4 (open circles);

LPS from Burkholderia multivorans—DOPE (3 : 1) at pH 7.4

(full triangles); LPS from Burkholderia multivorans—DOPE (3 : 1) at



through the models reported in the text have are also shown.

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where R is the radius of the base, fcyl the cylinder volume

fraction, and rc � r0 the scattering length density difference

between the cylinders and the solvent. In the low q region

(q o 0.006 A�1) the trend is similar to what was found for the

B. cenocepacia and A. tumefaciens, denoting the presence of

multilamellar liposomes. In this case, scattering cross sections

have been analyzed assuming each kind of aggregate to scatter

independently from the other and expressing (dS/dO) as the

sum of the eqn (3) and (4), taking into account for the relative

number density of the objects, strictly connected to the volume

fractions fcyl and flip. This procedure allows the estimation of

the micelle radius R that results to be (4.2 � 0.2) nm, while the

liposome structural parameters are reported in Table 2.

Although the lack of the Guinier regime for the micelles does

not allow obtaining a value of the length of cylindrical

micelles, l, nonetheless an estimation for such value can be

obtained from the knowledge of the DLS hydrodynamic

radius RH that for a cylinder is given by34

RH ¼ffiffiffiffiffiffiffiffiffiffiffi3

4R2l

3

rð1:0304þ 0:0193xþ 0:06229x2

þ 0:00476x3 þ 0:00166x4 þ 2:66� 10�6x7Þð5Þ

where R is the cylinder radius and x= ln[l/(2R)]. From eqn (5)

the length of the micelles formed by B. multivorans S-LPS has

been estimated to be (68 � 2) nm.

In Table 2 we also report the structural parameters obtained

for S. minnesota R-LPS.15 In this case only unilamellar

liposomes were detected.

In the presence of DOPE in aggregate formulation, at both

investigated pH values, the scattering profile is quite similar

for all three LPS considered in the present work. In all cases,

dS/dO shows a power law decrease with a o 2 at low q and a

shoulder at higher q (q D 0.02 A�1 for both B. cenocepacia

R-LPS and B. multivorans S-LPS and q D 0.07 A�1 for

A. tumefaciens R-LPS). Consequently the experimental data

were fitted modeling the systems as constituted by multi-

lamellar vesicles, using the model developed by Kotarchyk

and Ritzau.30 The fitting parameters are reported in Table 2.

Interestingly, in the presence of DOPE, also in the case of

S-LPS derived from B. multivorans no evidence of smaller

aggregate was detected.

3.3 EPR measurements: bilayer structuring

EPR spectroscopy, by using spin-labeled lipids has been

proved to give substantial information on the acyl chains

structuring in the lipid bilayers.35–38 In the present study, the

samples investigated were phosphatidylcholine spin-labeled on

the 5 or 14 C-atom of the sn-2 chain (5-PCSL and 14-PCSL,

respectively) incorporated in LPS liposomes. Incorporation,

and molecular dispersion, of the spin-label into the LPS

bilayer are highlighted by the absence of any evidence of

spin-exchange in the registered spectra, as would be expected

in the case of spin-label self-aggregation. 5-PCSL bears

the radical label close to the molecule head group, and

consequently allows monitoring the behavior of the region


systems: LPS from Agrobacterium tumefaciens at pH 7.4 (open circles);

LPS from Agrobacterium tumefaciens—DOPE (3 : 1) at pH 7.4

(full triangles); LPS from Agrobacterium tumefaciens—DOPE (3 : 1) at



through the models reported in the text are also shown.

Table 2 Structural parameters at 25 1C for the liposomes formed by LPS, obtained through the fitting of the model described in the text to SANSdata. The table reports the number of lamellae, N, the average lamellar thickness, d, the Zimm index of the thickness distribution function, Z, themean distance between two consecutive lamellae, dl, and the Hosemann factor, sdl/dl. Finally, for samples containing both cylindrical micelles andliposomes, the relative volume fractions of both kinds of aggregates fcyl and flip are also reported. In the table, LPS are indicated by the name ofthe bacterium from which they are obtained

System N d=nm Z dl=nm sdl=dl fcyl flip

Salmonella minnesotaa 1 4.3 � 0.2Burkholderia cenocepacia pH 7.4 4 � 1 2.6 � 0.2 10.5 � 0.1 17.0 � 0.3 0.59 � 0.01Burkholderia cenocepacia—DOPE pH 7.4 3 � 1 2.4 � 0.6 10.3 � 0.1 26.4 � 0.4 0.55 � 0.01Burkholderia cenocepacia—DOPE pH 9.1 4 � 1 2.8 � 0.2 10.3 � 0.1 23.4 � 0.4 0.81 � 0.02Burkholderia multivorans pH 7.4b 2 � 1 4.9 � 0.2 10.4 � 0.1 30.0 � 0.5 0.60 � 0.01 0.0018 � 0.0002 0.0049 � 0.0005Burkholderia multivorans—DOPE pH 7.4 4 � 1 5.0 � 0.1 10.3 � 0.1 32.0 � 0.5 0.60 � 0.01 0.0019 + 0.0002 0.0048 � 0.0005Burkholderia multivorans—DOPE pH 9.1 5 � 1 4.2 � 0.1 10.5 � 0.1 27.8 � 0.5 0.62 � 0.01 0.0020 � 0.0002 0.0047 � 0.0005Agrobacterium tumefaciens pH 7.4 3 � 1 4.7 � 0.4 10.3 � 0.1 13.7 � 0.5 0.60 � 0.01Agrobacterium tumefaciens—DOPE pH 7.4 5 � 1 4.8 � 0.4 10.3 � 0.1 11.5 � 0.4 0.60 � 0.01Agrobacterium tumefaciens—DOPE pH 9.1 4 � 1 4.5 � 0.2 10.3 � 0.1 16.9 � 0.3 0.60 � 0.01

a Data from ref. 15. b In this sample cylindrical micelles were also present, with radius R = (4.2 � 0.2) nm and length (68 � 2) nm, see text.

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of the membrane inner core closer to the polar external layers.

In contrast, 14-PCSL bears the radical label close to the

terminal methyl group of the acyl chain, thus allowing

monitoring the behavior of the more internal region of the

membrane hydrophobic core.

In all the systems analyzed in the present work, the 5-PCSL

spectrum presents a clearly defined axial anisotropy, see Fig. 8,

thus indicating that, in all cases, the mobility of the label in the

region of the bilayer just below the hydrophilic external

surface is strongly reduced. In contrast, all the 14-PCSL

spectra show an almost isotropic three-line signal, indicative

of a rather free motion of the radical label.

In an attempt to quantitatively analyze the spectra, we

determined the order parameter, S. This parameter is related

to the angular amplitudes of motion of the label, which in turn

reflects the motion of the acyl chain segment to which the label

is bound. In particular, S was calculated according to the

relation

S ¼ðTk � T?ÞðTzz � TxxÞ

aN

a0Nð6Þ

where TJ and T> are two phenomenological hyperfine

splitting parameters which can be experimentally determined

for each spin-labeled phospholipid as shown in Fig. 8 (note

that 2T0? ¼ 2T? � 1:6, see ref. 39). Txx and Tzz are the

principal elements of the real hyperfine splitting tensor in the

spin Hamiltonian of the spin-label, which can be measured

from the corresponding single-crystal EPR spectrum and are

reported in the literature (Txx = 6.1 G and Tzz = 32.4 G,

ref. 39). aN and a0N are the isotropic hyperfine coupling

constants for the spin-label in crystal state and in the

membrane, respectively, given by:

aN =1

3(Tzz+2Txx) (7)

a0N ¼

1

3ðTk þ 2T?Þ ð8Þ

The isotropic hyperfine coupling constant is an index of the

micropolarity experienced by the nitroxide; in particular, it

increases with the environmental polarity. The aN=a0N ratio in

eqn (6) corrects the order parameter for polarity differences

between the crystal state and the membrane.

The S and a0N values obtained from the spectra registered at

25 1C are collected in Table 3. In the case of S. minnesota

R-LPS we report the value determined from the 5-PCSL

spectrum reported in literature.15 By comparing the results

obtained for 5-PCSL and 14-PCSL it is possible to observe

that both parameters decrease with increasing the depth at

which the label is inserted in the bilayer, indicating that both

the acyl chains ordering and the environmental polarity

decrease. Concerning variations due to the peculiar LPS, it is

to be observed that the values obtained for 5-PCSL show that

liposomes formed by R-LPS from S. minnesota present values

of order parameter substantially lower than those observed in

the case of B. cenocepacia and A. tumefaciens R-LPS. For

S-LPS from B. multivorans an intermediate value is obtained.

It is interesting to observe that ordering of acyl chains in the

LPS bilayer is higher than that observed for typical phospho-

lipids (S E 0.6 for 5-PCSL39).

The lower S values obtained for 14-PCSL reflect the higher

fluidity of the more internal region of the bilayer hydrophobic

core. Among the various LPS, the same trend found for

5-PCSL is observed. For both spin-labels, Table 3 shows that

the a0N values do not depend on the considered LPS.

In the case of 5-PCSL, we also studied the S and a0N

variation with temperature, see Fig. 9. In all cases, a continuous

decrease was registered, thus indicating that the dynamics of

the lipid chain significantly increases with temperature. In the

case of R-LPS from S. minnesota, perusal of the figure reveals

a very broad sigmoidal trend, with the inflection centered

Fig. 8 EPR spectra of 5-PCSL and 14-PCSL in liposomes formed by

LPS from Burkholderia cenocepacia (A and A0 for 5- and 14-PCSL,

respectively),Burkholderia multivorans (B, B0),Agrobacterium tumefaciens

(C, C0) at pH 7.4 and 25 1C. The magnetic field scan range is 90 G.

Table 3 Microstructural parameters at 25 1C for the liposomesformed by LPS, through the quantitative analysis of EPR datadescribed in the text. The table reports the order parameter, S, andthe isotropic hyperfine coupling constant, a

0N . In the table, LPS are

indicated by the name of the bacterium from which they are obtained

System S a0N=G

5-PCSLSalmonella minnesotaa 0.69 � 0.01 15.4 � 0.1Burkholderia cenocepacia pH 7.4 0.77 � 0.02 15.6 � 0.2Burkholderia multivorans pH 7.4 0.73 � 0.01 15.5 � 0.2Agrobacterium tumefaciens pH 7.4 0.76 � 0.02 15.6 � 0.214-PCSLBurkholderia cenocepacia pH 7.4 0.24 � 0.02 13.6 � 0.2Burkholderia multivorans pH 7.4 0.20 � 0.03 13.5 � 0.2Agrobacterium tumefaciens pH 7.4 0.27 � 0.02 13.6 � 0.2

a Data computed from the spectrum reported in ref. 15.

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between 30 and 35 1C; this evidence is related to a transition

from gel- to fluid-phase of the LPS acyl chain arrangement in

the bilayer. For all the other LPS considered, only a slight

slope change is observed in the same temperature range.

Even in this case, no significant a0N variation was observed

(mean value 15.6 � 0.3 G).

4. Discussion

The focus of the present work relies on the relationship

between the LPS molecular structure and the morphology

and behavior of the aggregates they form in aqueous medium.

Indeed, the molecular structure determines the local self-

organization of the molecules, which in turn influences the

aggregate architecture and dynamics. For this reason it is

worthy to analyse in details the main analogies/differences

among the molecular structures of the LPS used, shown in

Fig. 1, before discussing the experimental results.

Two of the LPS considered in the present work, specifically

those derived from B. cenocepacia and A. tumefaciens, lack the

O-chain and therefore are R-LPS. In contrast, B. multivorans

LPS presents an extended O-chain (i.e., it is a S-LPS), whose

primary structure consists of two O-polysaccharide chains

present in different amounts and made up of repeating units

both containing deoxy sugars. In this case, Fig. 1 does not

show the entire S-LPS molecule, but only the lipid A and the

repeating units structures. B. multivorans S-LPS presents the

same lipid A of B. cenocepacia R-LPS, and a similar core

oligosaccharide. Thus, the comparison between the results

obtained for B. cenocepacia R-LPS and B. multivorans

S-LPS allows to discriminate the effect of the O-chain on the

molecules self-aggregation.

The R-LPS derived from A. tumefaciens presents a peculiar

lipid A, because of the presence of an unusually long chain

fatty acid (C28 : 0 (27-OH)), not stoichiometrically esterified

by a 3-hydroxy-butyroyl residue at its hydroxy group.40 Thus,

the analysis of the results obtained for A. tumefaciens R-LPS

highlights the extent to which LPS self-aggregation is

influenced by the lipid A primary structure.

Last, we compare the results obtained for these three LPS

with those already published by us relative to the R-LPS

derived from S. enterica serotype minnesota strain 595

(Re mutant).15 The R-LPS from the Re mutants represents

the deep-rough chemotype with the shortest saccharidic chain

length formed by the lipid A and Kdo as the sole constituent of

the core. Thus, in the present work, the comparison between

the properties of S. minnesota R-LPS aggregates with those

presented by the other LPS gives an indications of the

relevance of the glycolipid portion with respect to the

saccharidic component in determining the molecule self-

aggregation.

However, before starting to discuss the experimental results,

it is to be stressed that all the comparisons presented below

have to be handled with caution, since a single-variable

analysis is not feasible, the LPS molecules differentiating for

more than one feature.

The data collected in the present work indicate that all the

considered LPS form liposomes, whose dimension is coarsely

regulated by pore dimension of the membrane used for the

extrusion procedure. These aggregates, similar to phospholipid

liposomes, are likely to be metastable. However, their

morphological features are constant over long periods

(weeks, at least). The main driving force of LPS self-aggregation

is the hydrophobic interaction among the lipid A acyl chains.

This interaction forces the molecules to self-organize in a

bilayered structure with the acyl chains forming an inner

hydrophobic domain and the saccharidic portion protruding

in the external aqueous medium. The bilayer thickness, d, can

be evaluated by SANS measurements. As discussed in the

Results section, the d values represent the thickness of the

hydrophobic layer constituted by the acyl chains rather than

the whole thickness of LPS bilayer. B. cenocepacia presents the

lower d value with respect to the other LPS (d = 2.6 nm). The

linear extension of the lipophilic part of a single molecule of

lipid A is approximately 1.9 nm, so that a bilayer composed of

two opposing leaflets of LPS molecules with completely

extended acyl chains perpendicular to the disaccharidic back-

bone is expected to have a thickness of at leastB3.8 nm. Thus,

our experimental evidence suggests either a certain degree of

interdigitation between the acyl chains of the two opposing

lipid A leaflets, or a tilting of the acyl chains with respect

to the disaccharidic backbone. EPR measurements allow to

discriminate between these two possibilities; in fact, inter-

digitation reduces the mobility of the more internal segments

of the acyl chains, so that the spectrum of the spin-probe

14-PCSL is expected to present a slower component.41 The

14-PCSL spectrum in liposomes formed by B. cenocepacia

R-LPS presents an almost isotropic lineshape indicating a

relatively fast motion of the chain segments, an evidence of

no interdigitation to occur. Consequently, we conclude that

the low d value obtained by SANS is likely to be due to a

tilting of the acyl chains relative to the disaccharide units

delimiting the bilayer inner hydrophobic domain. A similar

chain arrangement in LPS bilayers was suggested by the

analysis of FTIR and AFM experiments.42,43

Interesting is the comparison of the self-association

behavior of B. cenocepacia R-LPS with the A. tumefaciens

and S. minnesota ones. In the last two cases, the higher bilayer

Fig. 9 Order parameter, S, of 5-PCSL in liposomes formed by LPS

from Burkholderia cenocepacia (full squares), Burkholderia multivorans

(open circles), Agrobacterium tumefaciens (open squares) and

Salmonella minnesota (full circles), at pH 7.4 as a function of the

temperature.

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thickness could be related to a different arrangement of the

acyl chains, which probably tend to be perpendicular to

the disaccharide backbone. This change could be induced

by the higher number of acyl chains per disaccharide unit

(6 for S. minnesota R-LPS vs. 5 for B. cenocepacia R-LPS) or

to the presence of the unusually longer fatty acid, in the case of

A. tumefaciens R-LPS. For S. minnesota R-LPS, the different

orientation significantly reduces the acyl chains ordering, as

shown by the S value obtained for 5-PCSL by EPRmeasurements.

In the case of A. tumefaciens R-LPS, the relatively high

ordering of the acyl chains in both 5- and 14- position could

support the hypothesis that the acyl chain of the (C28:0 (27-OH))

fatty acid passes through both leaflets constituting the

bilayer,40 with a consequent mobility reduction. Actually,

the presence of the long acyl chain could be an evolutive

strategy for stabilizing liposomes formed by A. tumefaciens

R-LPS, which are more negatively charged than those derived

from B. cenocepacia and B. multivorans LPS, whose phosphate

groups are partially substituted by a 4-deoxy-4-amino-arabinose

moiety, see Fig. 1.

Even more stimulating is the comparison between the

d values obtained for liposomes formed by B. multivorans

S-LPS and B. cenocepaciaR-LPS, which present the same lipid

A and a similar saccharidic core, and differ only in the

O-chain, which is absent in the latter case. As discussed above,

SANS measurements are almost insensitive to the saccharidic

portions of the bilayer, and consequently one could expect

similar SANS results for the two LPS. However, this is

absolutely not confirmed by the experimental results, showing

a d value much higher for liposomes formed by B. multivorans

S-LPS. This evidence has to be related to an indirect effect of

the O-chain. In consideration of the bulkiness of the saccharidic

portion of B. multivorans S-LPS, strong steric repulsion occurs

among the hydrophilic moieties of adjacent molecules in the

bilayer, thus disturbing the molecules packing. This distortion

propagates to the acyl chains, affecting their arrangement in

the hydrophobic inner layer, reducing the tightness of their

packing and their tilting with respect the bilayer surface,

and finally resulting in a d increase. This interpretation is

supported by the reduction of the S parameter derived from

EPR. Thus, the comparison between the data collected for

B. cenocepacia R-LPS and B. multivorans S-LPS indicates

that the self-aggregation of the glycolipid portions of LPS

molecules is strongly affected by the saccharidic portion, in a

complex interplay of hydrophobic, steric and electrostatic

interactions.

In this context, here we comment on the peculiarity of a

S-LPS (i.e., the LPS derived from B. multivorans) with respect

to R-LPS (all the others considered LPS), in that only in the

first case, according to the DLS measurements, liposomes

coexist with elongated micelles. Coexistence of micelles and

vesicles is well documented in the recent literature. Interestingly,

it is often found in aqueous mixtures of amphiphiles presenting

an extended hydrophilic moiety, such as a poly(ethylene oxide

chain).44–46 Conformation of the PEG coil is considered to be

decisive in establishing the morphology of the supramolecular

aggregates. We propose the polysaccharidic chain of S-LPS to

play a similar role. For steric reasons, a ‘‘globular’’ coil would

favor formation of spherical micelles, while a more extended

chain conformation is required in bilayered structures. Thus our

evidence has to be ascribed to the bulkiness of the hydrophilic

portion of the B. multivorans S-LPS molecule, which leads to an

increased curvature of the aggregate surface, finally resulting in

the formation of rodlike micellar aggregates.

Interestingly, the 5-PCSL EPR spectrum in aggregates

formed by S-LPS derived from B. multivorans presents the

same line shape observed for the other LPS, with no super-

position of signals deriving from micellar aggregates. This

evidence, already observed for other lipids,47 suggests that

the dynamic state of the lipid acyl chains is preserved, despite

the change in morphology.

Coming back to a general discussion on liposomes formed

by LPS molecules, local ordering of the acyl chains is expected

to be affected by temperature. In the present work this aspect

was analyzed by EPR. With increasing temperature, the

fluidity of the bilayer increases, as highlighted by the

S decrease. In the case of the R-LPS deriving from S. minnesota,

the S values decrease according to a broad sigmoidal trend,

with an inflection at 30–35 1C. This trend could be related to a

transition of the acyl chains from a gel to a liquid crystalline

phase (a - b), which has been largely reported in the

literature.5 This transition is much less steep than that

observed for typical phospholipids.13,47 Furthermore, our data

indicate that it becomes even smoother in the case of LPS

derived from B. cenocepacia, B. multivorans and A. tumefaciens,

for which only a slight slope change is observed. The reason

why in the case of LPS the transition is so smooth, could be the

polydispersity of acyl chain length, and their different

positioning with respect to the saccharidic headgroup.

For all the considered LPS, we have observed the tendency

of the liposomes to re-arrange forming multilayered

structures. This is unequivocally shown by SANS data, even

though, because of the quality of the data and of limitations of

the adopted fitting model, the number of concentric lamellae

can be only qualitatively estimated. The extrusion technique

we used in the present work is considered a reliable method to

produce unilamellar vesicles,48 even though in a limited number

of cases it has also been found to lead to formation of

multilamellar aggregates. This could depend on lipid

composition49,50 or on external conditions, such as solvent

composition.51 Concerning LPS, we tentatively propose liposome

multilamellarity to be related to the favorable interaction

among saccharidic surfaces of concentric bilayers, probably

connected to the formation of an extended network of

H-bonds. Interestingly, a similar interpretation has been

recently proposed to explain why glycolipids tend to form

multi- and not unilamellar structures.52

Last, we comment on the effect of inclusion of DOPE in

liposomes formulation. Generally speaking, the effects are not

dramatic. The only relevant DOPE effect was registered in the

case of S-LPS derived from B. multivorans. In this case, the

addition of DOPE leads to an increase of the vesicles

population (see Fig. 3), as put in evidence by DLS results.

Furthermore, the reduction of the micelles population is such

that they are not detected by SANS measurements. These

experimental evidences indicate that the bilayer is stabilized by

the insertion of DOPE, which causes a reduction of the steric

repulsion among the LPS headgroups.

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In conclusion, the results presented in this work have high-

lighted the relationship between the molecular structure of

some LPS molecules and the structure of their aggregates.

Interestingly, the relationship is often not obvious, in that the

aggregate architecture and dynamics are the final result of a

complex interplay of all the possible interactions (hydrophobic,

electrostatic, steric) set up among the LPS molecules. In

biological context, our results suggest that the rich biodiversity

of LPS molecular structure could be fundamental to finely

tuning the structure and functional properties of the outer

membrane of Gram negative bacteria and, consequently, their

biological behavior.

Acknowledgements

The authors thank CSGI (Consorzio Interuniversitario per lo

sviluppo dei Sistemi a Grande Interfase) and MIUR (PRIN

2007) for financial support. B. multivorans cells were kindly

furnished by Dr Paola Cescutti (Universita di Trieste),

B. cenocepacia cells were kindly furnished by Dr Anthony

De-Soyza (University of Newcastle) and R-LPS from

A. tumefaciens was a kind gift from Dr Cristina De Castro

(Universita di Napoli). Forschungszentrum Julich is

acknowledged for provision of beam time. SANS experiments

were supported by the European Commission, NMI3 contract

RII3-CT-2003-505925. The authors thank Prof. Lucia

Costantino for her helpful comments. Finally, we thank the

referees whose comments helped improve the manuscript.

References

1 N. Ruiz, D. Kahne and J. Silhavy Thomas, Advances inunderstanding bacterial outer-membrane biogenesis, Nat. Rev.Microbiol., 2006, 4(1), 57–66.

2 A. Silipo, C. De Castro, R. Lanzetta, M. Parrilli and A. Molinaro,in Prokaryotic Cell Wall Compounds: Structure and Biochemistry,ed. H. Konig, H. Claus and A. Varma, Springer, Heidelberg, 2010.

3 O. Holst and A. Molinaro, in Microbial Glycobiology,ed. A. P. Moran, Elsevier, London, 2009.

4 U. Seydel, A. J. Ulmer, S. Uhlig and E. T. Rietschel, in MemnrnStructure in Disease and Drug Therapy, ed. G. Zimmer, MarcelDekker Inc., New York, 1999.

5 K. Brandenburg and U. Seydel, A comment on the preparation ofliposomes from and on the beta .tautm. alpha acyl chain meltingbehavior of rough mutant lipopolysaccharide, Biochim. Biophys.Acta, Biomembr., 1991, 1069(1), 1–4.

6 K. Nixdorff, J. Gmeiner and H. H. Martin, Interaction oflipopolysaccharide with detergents and its possible role in thedetergent resistance of the outer membrane of Gram-negativebacteria, Biochim. Biophys. Acta, Biomembr., 1978, 510(1), 87–98.

7 T. Gutsmann, A. B. Schromm, M. H. J. Koch, S. Kusumoto,K. Fukase, M. Oikawa, U. Seydel and K. Brandenburg,Lipopolysaccharide-binding protein-mediated interaction oflipid A from different origin with phospholipid membranes,Phys. Chem. Chem. Phys., 2000, 2(20), 4521–4528.

8 K. Brandenburg, M. Matsuura, H. Heine, M. Muller, M. Kiso,H. Ishida, M. H. J. Koch and U. Seydel, Biophysical characterizationof triacyl monosaccharide lipid A partial structures in relation tobioactivity, Biophys. J., 2002, 83(1), 322–333.

9 H. Labischinski, E. Vorgel, W. Uebach, R. P. May andH. Bradaczek, Architecture of bacterial lipid A in solution.A neutron small-angle scattering study, Eur. J. Biochem., 1990,190(2), 359–63.

10 E. Urban, A. Bota and B. Kocsis, Non-bilayer formation in theDPPE-DPPG vesicle system induced by deep rough mutant ofSalmonella minnesota R595 lipopolysaccharide, Colloids Surf., B,2006, 48(2), 106–11.

11 K. Nomura, T. Inaba, K. Morigaki, K. Brandenburg, U. Seydeland S. Kusumoto, Interaction of lipopolysaccharide and phospho-lipid in mixed membranes: solid-state 31P-NMR spectroscopic andmicroscopic investigations, Biophys. J., 2008, 95(3), 1226–1238.

12 N. C. Santos, A. C. Silva, M. A. R. B. Castanho, J. Martins-Silvaand C. Saldanha, Evaluation of lipopolysaccharide aggregation bylight scattering spectroscopy, ChemBioChem, 2003, 4(1), 96–100.

13 E. Urban, A. Bota, B. Kocsis and K. Lohner, Distortion of thelamellar arrangement of phospholipids by deep rough mutantlipopolysaccharide from Salmonella minnesota, J. Therm. Anal.Calorim., 2005, 82(2), 463–469.

14 M. F. Henning, S. Sanchez and L. Bakas, Visualization andanalysis of lipopolysaccharide distribution in binary phospholipidbilayers, Biochem. Biophys. Res. Commun., 2009, 383(1), 22–26.

15 G. D’Errico, A. Silipo, G. Mangiapia, A. Molinaro, L. Paduanoand R. Lanzetta, Mesoscopic and microstructural characterizationof liposomes formed by the lipooligosaccharide from Salmonellaminnesota strain 595 (Re mutant), Phys. Chem. Chem. Phys., 2009,11(13), 2314–2322.

16 R. F. Epand, P. B. Savage and R. M. Epand, Bacterial lipidcomposition and the antimicrobial efficacy of cationic steroidcompounds (Ceragenins), Biochim. Biophys. Acta, Biomembr.,2007, 1768(10), 2500–2509.

17 T. Ierano, A. Silipo, P. Cescutti, M. R. Leone, R. Rizzo,R. Lanzetta, M. Parrilli and A. Molinaro, Structural study andconformational behavior of the two different lipopolysaccharideO-antigens produced by the cystic fibrosis pathogen Burkholderiamultivorans, Chem.–Eur. J., 2009, 15(29), 7156–7166, S7156/1–S7156/4.

18 A. Silipo, A. Molinaro, T. Ierano, A. De Soyza, L. Sturiale,D. Garozzo, C. Aldridge, P. A. Corris, C. M. A. Khan,R. Lanzetta and M. Parrilli, The complete structure and pro-inflammatory activity of the lipooligosaccharide of the highlyepidemic and virulent gram-negative bacterium Burkholderiacenocepacia ET-12 (strain J2315), Chem.–Eur. J., 2007, 13(12),3501–3511, S3401/1–23501/9.

19 C. De Castro, A. Carannante, R. Lanzetta, V. Liparoti,A. Molinaro and M. Parrilli, Core oligosaccharide structure fromthe highly phytopathogenic Agrobacterium tumefaciens TT111and conformational analysis of the putative rhamnan epitope,Glycobiology, 2006, 16(12), 1272–1280.

20 M. Ishinaga, R. Kanamoto and M. Kito, Distribution of phospho-lipid molecular species in outer and cytoplasmic membranes ofEscherichia coli, Journal of Biochemistry, 1979, 86(1), 161–5.

21 A. K. Covington, F. S. Bates and R. A. Durst, Definition ofpH scales, standard reference values, measurement of pH andrelated terminology, Pure Appl. Chem., 1985, 57(3), 531–42.

22 R. Peters, ALV-5000/E/EPP% ALV-60X0 for WINDOWS-95/NT4.0 Software, 3.0.1.12, 2003.

23 G. A. Brehm and V. A. Bloomfield, Analysis of polydispersity inpolymer solutions by inelastic laser light scattering, Macro-molecules, 1975, 8(5), 663–5.

24 G. D. Wignall and F. S. Bates, Absolute calibration of small-angleneutron scattering data, J. Appl. Crystallogr., 1987, 20(1), 28–40.

25 T. P. Russell, J. S. Lin, S. Spooner and G. D. Wignall, Inter-calibration of small-angle X-ray and neutron scattering data,J. Appl. Crystallogr., 1988, 21(6), 629–38.

26 H. J. V. Tyrrell and K. R. Harris, Diffusion in Liquids:A Theoretical and Experimental Study, Butterworths, London, 1984.

27 M. Kotlarchyk and S. H. Chen, Analysis of small angle neutronscattering spectra from polydisperse interacting colloids, J. Chem.Phys., 1983, 79(5), 2461–9.

28 M. Vaccaro, R. Del Litto, G. Mangiapia, A. M. Carnerup,G. D’Errico, F. Ruffo and L. Paduano, Lipid based nanovectorscontaining ruthenium complexes: a potential route in cancertherapy, Chem. Commun., 2009, (11), 1404–1406.

29 H. Frielinghaus, Small-angle scattering model for multilamellarvesicles, Physical Review E: Statistical, Nonlinear, and Soft MatterPhysics, 2007, 76(5–1), 051603/1–051603/8.

30 M. Kotlarchyk and S. M. Ritzau, Paracrystal model of the high-temperature lamellar phase of a ternary microemulsion system,J. Appl. Crystallogr., 1991, 24(5), 753–8.

31 M. Vaccaro, G. Mangiapia, L. Paduano, E. Gianolio, A.Accardo, D. Tesauro and G. Morelli, Structural and relaxometriccharacterization of peptide aggregates containing gadolinium

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complexes as potential selective contrast agents in MRI,ChemPhysChem, 2007, 8(17), 2526–2538.

32 M. Vaccaro, A. Accardo, D. Tesauro, G. Mangiapia, D. Loef,K. Schillen, O. Soederman, G. Morelli and L. Paduano,Supramolecular Aggregates of Amphiphilic GadoliniumComplexes as Blood Pool MRI/MRA Contrast Agents: Physico-chemical Characterization, Langmuir, 2006, 22(15), 6635–6643.

33 A. Radulescu, R. T. Mathers, G. W. Coates, D. Richter andL. J. Fetters, A SANS Study of the Self-Assembly in Solution ofSyndiotactic Polypropylene Homopolymers, Syndiotactic Poly-propylene-block-poly(ethylene-co-propylene) Diblock Copolymers,and an Alternating Atactic-Isotactic Multisegment Polypropylene,Macromolecules, 2004, 37(18), 6962–6971.

34 S. Hansen, Translational friction coefficients for cylinders ofarbitrary axial ratios estimated by Monte Carlo simulation,J. Chem. Phys., 2004, 121(18), 9111–9115.

35 D. Marsh, Structural and thermodynamic determinants of chain-melting transition temperatures for phospholipid and glycolipidsmembranes, Biochim. Biophys. Acta, Biomembr., 2010, 1798(1),40–51.

36 A. Lange, D. Marsh, K. H. Wassmer, P. Meier and G. Kothe,Electron spin resonance study of phospholipid membranesemploying a comprehensive line-shape model, Biochemistry,1985, 24(16), 4383–92.

37 M. Moser, D. Marsh, P. Meier, K. H. Wassmer and G. Kothe,Chain configuration and flexibility gradient in phospholipidmembranes. Comparison between spin-label electron spin resonanceand deuteron nuclear magnetic resonance, and identification ofnew conformations, Biophys. J., 1989, 55(1), 111–23.

38 W. L. Hubbell and H. M. McConnell, Molecular motion in spin-labeled phospholipids and membranes, J. Am. Chem. Soc., 1971,93(2), 314–26.

39 L. M. Gordon and R. D. Sauerheber, Studies on spin-labeled egglecithin dispersions, Biochim. Biophys. Acta, Biomembr., 1977,466(1), 34–43.

40 A. Silipo, C. de Castro, R. Lanzetta, A. Molinaro and M. Parrilli,Full structural characterization of the lipid A components fromthe Agrobacterium tumefaciens strain C58 lipopolysaccharidefraction, Glycobiology, 2004, 14(9), 805–815.

41 I. Plasencia, F. Baumgart, D. Andreu, D. Marsh and J. Perez-Gil,Effect of acylation on the interaction of the N-Terminal segment of

pulmonary surfactant protein SP-C with phospholipid membranes,Biochim. Biophys. Acta, Biomembr., 2008, 1778(5), 1274–1282.

42 U. Seydel, M. Oikawa, K. Fukase, S. Kusumoto andK. Brandenburg, Intrinsic conformation of lipid A is responsiblefor agonistic and antagonistic activity, Eur. J. Biochem., 2000,267(10), 3032–3039.

43 S. Roes, U. Seydel and T. Gutsmann, Probing the Properties ofLipopolysaccharide Monolayers and Their Interaction with theAntimicrobial Peptide Polymyxin B by Atomic Force Microscopy,Langmuir, 2005, 21(15), 6970–6978.

44 F. Li, S. Prevost, R. Schweins, A. T. M. Marcelis, F. A. M.Leermakers, M. A. Cohen Stuart and E. J. R. Sudhoelter,Small monodisperse unilamellar vesicles from binary copolymermixtures, Soft Matter, 2009, 5(21), 4169–4172.

45 C. Leal, S. Roegnvaldsson, S. Fossheim, E. A. Nilssen andD. Topgaard, Dynamic and structural aspects of PEGylatedliposomes monitored by NMR, J. Colloid Interface Sci., 2008,325(2), 485–493.

46 M. Vaccaro, G. Mangiapia, A. Radulescu, K. Schillen,G. D’Errico, G. Morelli and L. Paduano, Colloidal particlescomposed of amphiphilic molecules binding gadolinium complexesand peptides as tumor-specific contrast agents in MRI: physico-chemical characterization, Soft Matter, 2009, 5(13), 2504–2512.

47 G. D’Errico, A. M. D’Ursi and D. Marsh, Interaction of a PeptideDerived from Glycoprotein gp36 of Feline ImmunodeficiencyVirus and Its Lipoylated Analogue with Phospholipid Membranes,Biochemistry, 2008, 47(19), 5317–5327.

48 B. Mui, L. Chow and M. J. Hope, Extrusion technique togenerate liposomes of defined size, Methods Enzymol., 2003,367(Liposomes, Part A), 3–14.

49 P. Fromherz, Lipid-vesicle structure: size control by edge-activeagents, Chem. Phys. Lett., 1983, 94(3), 259–66.

50 N. E. Gabriel and M. F. Roberts, Spontaneous formation of stableunilamellar vesicles, Biochemistry, 1984, 23(18), 4011–15.

51 M. A. Kiselev, P. Lesieur, A. M. Kisselev, D. Lombardo,M. Killany and S. Lesieur, Sucrose solutions as prospectivemedium to study the vesicle structure: SAXS and SANS study,J. Alloys Compd., 2001, 328(1–2), 71–76.

52 J. Howe, M. Von Minden, T. Gutsmann, M. H. J. Koch, M. Wulf,S. Gerber, G. Milkereit, V. Vill and K. Brandenburg, Structuralpreferences of dioleoyl glycolipids with mono- and disaccharidehead groups, Chem. Phys. Lipids, 2007, 149(1–2), 52–58.

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Characterization of liposomes formed by lipopolysaccharides from Burkholderia cenocepacia,...

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