TAMING THE GRIFFIN
Jobst Liebau
Taming the Griffin
Membrane interactions of peripheral and monotopic glycosyltransferases anddynamics of bacterial and plant lipids in bicelles
Jobst Liebau
©Jobst Liebau, Stockholm University 2017 ISBN print 978-91-7649-978-8ISBN PDF 978-91-7649-979-5 Cover art by Gefion Liebau: A griffin guarding a lipid membrane inside an NMR spectrometer. Printed in Sweden by Universitetsservice US-AB, Stockholm 2017Distributor: Department of Biochemistry and Biophysics, Stockholm University
Contents
List of Publications I
List of Additional Publications III
Abbreviations V
List of Figures VII
1 Introduction 11.1 Where does the journey lead us? . . . . . . . . . . . . . . . . 1
2 Lipid membranes in vivo and in vitro 52.1 General features of biological membranes . . . . . . . . . . . 6
2.2 Lipid dynamics . . . . . . . . . . . . . . . . . . . . . . . . . 7
2.3 Biological membranes of plants and bacteria . . . . . . . . . . 9
2.3.1 E. coli inner membranes . . . . . . . . . . . . . . . . 9
2.3.2 Lipopolysaccharides . . . . . . . . . . . . . . . . . . 12
2.3.3 Chloroplast membranes . . . . . . . . . . . . . . . . 12
2.4 Membrane mimetics for biophysical applications . . . . . . . 13
2.4.1 Large unilamellar vesicles . . . . . . . . . . . . . . . 13
2.4.2 Micelles and bicelles . . . . . . . . . . . . . . . . . . 14
3 Glycosyltransferases 173.1 Membrane interaction of GT-B glycosyltransferases . . . . . . 18
3.2 The glycosyltransferase WaaG . . . . . . . . . . . . . . . . . 20
4 Methods 214.1 Circular dichroism . . . . . . . . . . . . . . . . . . . . . . . 22
4.2 Fluorescence spectroscopy . . . . . . . . . . . . . . . . . . . 22
4.2.1 Fluorescence of aromatic amino acids . . . . . . . . . 22
4.2.2 Quenching . . . . . . . . . . . . . . . . . . . . . . . 23
4.2.3 The parallax method . . . . . . . . . . . . . . . . . . 23
4.2.4 Anisotropy . . . . . . . . . . . . . . . . . . . . . . . 24
4.2.5 Stopped-flow fluorescence . . . . . . . . . . . . . . . 25
4.2.6 Measurements of the dissociation constant KD . . . . . 25
4.3 Nuclear magnetic resonance . . . . . . . . . . . . . . . . . . 26
4.3.1 Translational diffusion . . . . . . . . . . . . . . . . . 27
4.3.2 Structure determination . . . . . . . . . . . . . . . . . 28
4.3.3 Dynamics . . . . . . . . . . . . . . . . . . . . . . . . 29
4.3.4 Distance measurements using paramagnetic relaxation
enhancements . . . . . . . . . . . . . . . . . . . . . . 31
5 Results and Discussion 335.1 Mimicking plant and bacterial membranes . . . . . . . . . . . 33
5.1.1 Size of bicelles . . . . . . . . . . . . . . . . . . . . . 34
5.1.2 Lipid composition of bicelles made from E. coli phos-
pholipids . . . . . . . . . . . . . . . . . . . . . . . . 34
5.1.3 Rapid dynamics in bicelles . . . . . . . . . . . . . . . 35
5.1.4 Immersion depths of carbon atoms in membranes . . . 36
5.1.5 Bicelles as a plant and bacterial membrane mimetic . . 38
5.2 Monotopic and peripheral glycosyltransferases . . . . . . . . 39
5.2.1 Anchoring of WaaG to membranes . . . . . . . . . . . 39
5.2.2 Membrane interaction of full-length WaaG . . . . . . 40
6 Conclusions 43
Populärvetenskaplig Sammanfattning 45
Populärwissenschaftliche Zusammenfassung 49
Popular science summary 53
Acknowledgements 57
References 59
List of Publications
The following publications and manuscripts are included in this thesis and are
referred to by their roman numerals in the text. Reprints of the published
articles were made with permission from the publishers.
I New membrane mimetics with galactolipids: Lipid properties infast-tumbling bicellesWeihua Ye, Jobst Liebau, and Lena Mäler, The Journal of PhysicalChemistry B, 117(4), 1044−1050 (2013).
II Characterization of bicelles constructed from native Escherichia colilipidsJobst Liebau, Pontus Pettersson, Philipp Zuber, Candan Ariöz, and Lena
Mäler, Biochimica et Biophysica Acta – Biomembranes, 1858(9),
2097−2105 (2016).
III Immersion depths of lipid carbons in bicelles measured byparamagnetic relaxation enhancementJobst Liebau and Lena Mäler, The Journal of Physical Chemistry B,
121(32), 7660−7670 (2017).
IV Membrane interaction of the glycosyltransferase WaaGJobst Liebau, Pontus Pettersson, Scarlett Szpryngiel, and Lena Mäler,
Biophysical Journal, 109(3), 552−563 (2015).
V The glycosyltransferase WaaG: a peripheral membrane protein?Jobst Liebau, Biao Fu, Christian Brown, and Lena Mäler, submittedmanuscript.
List of Additional Publications
i Anionic lipid binding to the foreign protein MGS provides a tightcoupling between phospholipid synthesis and protein overexpressionin Escherichia coliCandan Ariöz, Weihua Ye, Amin Bakali, Changrong Ge, Jobst Liebau,
Hans-Jörg Götzke, Andreas Barth, Åke Wieslander, and Lena Mäler,
Biochemistry, 52(33), 5533−5544 (2013).
ii Effect of lipid bilayer properties on the photocycle of greenproteorhodopsinLjubica Lindholm, Candan Ariöz, Michael Jawurek, Jobst Liebau, Lena
Mäler, Åke Wieslander, Christoph von Ballmoos, and Andreas Barth,
Biochimica et Biophysica Acta – Bioenergetics, 1847(8), 698−708
(2015).
iii Characterization of fast-tumbling isotropic bicelles by PFG diffusionNMRJobst Liebau, Weihua Ye, and Lena Mäler, Magnetic Resonance inChemistry, 55(5), 395−404 (2017).
iv Substrate-mimicking domain of nucleotide-exchange factorFes1/HspBP1 ensures efficient release of persistent substrates fromHsp70Naveen Kumar Chandappa Gowda, Jayasankar Mohanakrishnan Kaimal,
Roman Kityk, Chammiran Daniel, Jobst Liebau, Marie Öhman, Matthias
P. Mayer, and Claes Andréasson, submitted manuscript.
Abbreviations
CD circular dichroism
CL cardiolipin
CMC critical micelle concentration
DGDG digalactosyl diacylglycerol
DHPC 1,2-dihexanoyl-sn-glycero-3-phosphatidylcholine
DMPC 1,2-dimyristoyl-sn-glycero-3-phosphatidylcholine
DMPE 1,2-dimyristoyl-sn-glycero-3-phosphatidylethanolamine
DPC dodecylphosphocholine
EPR electron paramagnetic resonance
GT glycosyltransferase
LPS lipopolysaccharide
LUV large unilamellar vesicle
MD molecular dynamics
MGDG monogalactosyl diacylglycerol
NOE nuclear Overhauser enhancement
PE phosphatidylethanolamine
PG phosphatidylglycerol
PRE paramagnetic relaxation enhancement
SANS small-angle neutron scattering
SQDG sulfoquinovosyl diacylglycerol
List of Figures
2.1 Representatives of different classes of membrane-associated
proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
2.2 Dynamics and their approximate correlation times in lipids and
membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
2.3 Lipids found in E. coli inner membranes . . . . . . . . . . . . 10
2.4 Glycolipids found in chloroplast membranes . . . . . . . . . . 10
2.5 Schematic structure of lipopolysaccharides . . . . . . . . . . . 11
2.6 Approximate diameters of LUVs, micelles, and bicelles . . . . 14
2.7 Idealized depiction of a small isotropic bicelle with q = 0.5 . . 15
3.1 GT-A, GT-B, and GT-C fold of GTs . . . . . . . . . . . . . . 18
3.2 Model of the membrane interaction of membrane-associated
GT-B GTs . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19
4.1 Pulse sequence for measuring translational diffusion . . . . . . 28
4.2 Illustration of dynamic parameters in lipids . . . . . . . . . . 32
5.1 Membrane interaction of the glycosyltransferase WaaG via its
anchor MIR-WaaG . . . . . . . . . . . . . . . . . . . . . . . 41
6.1 WaaG anchoring to a lipid membrane . . . . . . . . . . . . . 47
1. Introduction
We’re just about to walk off the map.—George Mallory, 1921 British Mt. Everest reconnaissance expedition
Venturing out into uncharted territory, never knowing which obstacle needs to
be negotiated next, or whether this is even going to be possible—this is the
challenge that adventurers and scientists face alike. While George Mallory
was climbing Mt. Everest with an obscure map in his left hand and an ice pick
in his right, life science very often starts with a pipette in one’s right hand and
a test tube in one’s left. The scientist’s endeavor is less dangerous, but in no
way less exciting or challenging than the mountaineer’s. Both the adventurer
and the scientist push to extend the limits of knowledge about the world in an
effort to remove the blank spots from their respective maps.
1.1 Where does the journey lead us?
This work is an endeavor to explore the smallest realms of life. The uncharted
territory consists of proteins and lipids and their mutual interactions. The ice
picks are biophysical methods, and instead of obscure maps, models or ideas
of proteins and lipids that scientists have developed guide the way.
Membranes are highly complex aggregates of lipids and proteins. Fre-
quently, lipids are considered to be the ‘matrix’ for membrane proteins. This is
evident in, for example, the simple depiction often found in text books of lipid
membranes as two parallel lines indicating the membrane surface, onto which
a complicated protein structure is drawn. However, such depictions neglect
the crucial functional importance of lipids and protein-lipid interactions for
many biological processes. The importance of membranes goes beyond mere
structural function, and an understanding of lipid properties in membranes and
their interaction with proteins is crucial for a comprehensive understanding of
membrane-related processes.
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Lipid membranes have been extensively studied with biophysical tools. To
do so, native membranes very often need to be simplified or modified to ren-
der them amenable to experimentation. Membrane mimetics can be as simple
as an organic solvent, or as complex as a natural membrane extract, depend-
ing on the question that is being asked and the method used. The first focus
of this work is on the characterization by solution-state NMR of membrane
mimetics that mimic natural membranes more faithfully than those previously
employed. In particular, dynamics and localization of bacterial and plant lipids
inside membranes are investigated.
If lipids are important, so are proteins that synthesize lipids. A defec-
tive membrane reduces or abolishes the viability of organisms. It has been
shown, for instance, that defects in the structure of lipopolysaccharides (LPS),
lipids containing long polyglycan chains that are located in the outer leaflet of
the outer membrane in Gram-negative bacteria, destabilize the outer bacterial
membrane and make the bacterium more susceptible to antibiotics. This vul-
nerability can be exploited when searching for novel antibiotic targets. Multi-
resistant bacteria have become an imminent threat to society and it has recently
been estimated that by 2050 as many patients will die from infections caused
by bacteria resistant to all known antibiotics as will die of cancer [142]. More-
over, many forms of clinical treatment rely on the availability of functional
antibiotics. Even if the above prediction is not realized, antibiotic resistance
is already a threat today. Meanwhile, fewer and fewer new antibiotics are ap-
proved each year [187].
Proteins involved in the synthesis of bacterial lipids are thus promising
antibiotic targets. One such protein is the glycosyltransferase (GT) WaaG, a
membrane-associated protein that is involved in the synthesis of LPS. WaaG is
not only of interest because it is a promising antibiotic target but also because
of its peculiar function at the membrane: WaaG catalyzes the formation of a
glycosidic bond between a glucose sugar and a lipid acceptor. To do so, the
hydrophilic nucleotide sugar has to be brought into close proximity with a lipid
acceptor. At the same time, WaaG itself seems to be a hybrid between a soluble
and a membrane protein. Like the metaphor in the title of this thesis suggests,
WaaG is a microscopic analogy of a griffin. A griffin is a hybrid between lion
and eagle. WaaG appears to have properties that make it a hybrid creature of
its own kind. It can exist in both a soluble and a membrane-bound form. The
second focus of this thesis is thus on the investigation of the membrane interac-
tions of WaaG, and whether the observations made are in agreement with the
model of the membrane interaction of membrane-associated GTs previously
put forward.
The questions outlined above are addressed by a set of biophysical meth-
ods, including circular dichroism, fluorescence methods, and solution-state
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NMR, with a focus on the latter. These biophysical methods put different
demands on the systems studied, but what they all have in common is that
both the membrane and the protein need to be simplified or modified for ob-
servations to be possible. Simplifications or modifications include the use of
membrane mimetic systems like vesicles or bicelles, the study of isolated seg-
ments of WaaG, and modifications of WaaG necessary to introduce reporter
probes.
The thesis is structured as follows: properties of plant and bacterial lipid
membranes and of membrane systems that can mimic such lipid bilayers are
discussed in chapter 2. A brief overview of GTs in general and membrane-
associated GTs in particular is given in chapter 3. This overview includes
background information on the GT WaaG. The biophysical methods used to
study both the membrane interaction of WaaG and lipid properties are outlined
in chapter 4. The two questions that are at the center of this thesis, how lipids
localize and move in ‘realistic’ membrane systems, and how WaaG interacts
with such or similar membrane systems, are discussed in chapter 5, and a per-
spective on possible future research avenues is outlined in chapter 6. The work
is concluded by a popular science summary.
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2. Lipid membranes in vivo and
in vitro
God made the bulk; surfaces were invented by the devil.—Wolfgang Pauli
Biological membranes are highly complex aggregates consisting mainly of
lipids and proteins. Membranes compartmentalize cells, forming a semi-
permeable, protective barrier between the cell or compartment and the sur-
rounding environment, creating a confined space in which proteins and other
molecules exist at high concentrations and in which most processes required
to sustain life occur. Since membranes are the walls and gates of a cell, many
of those essential biological processes occur in or at membranes. Therefore,
apart from their protective and barrier functions, membranes also have other
functional roles, which comprise among others regulation and organization
of proteins, maintenance of charge and concentration gradients, information
transduction, and supply of lipid substrates for proteins [48; 125].
Biological membranes have to be simplified in order to make them
amenable to biophysical study. Membrane mimetics are of reduced complexity
compared to biological membranes and address method-specific limitations.
For instance, in solution-state NMR ‘large’ systems cannot be studied easily
and therefore membrane mimetics need to be ‘small’ (see also section 4.3).
In the following sections, general features of biological membranes are
outlined, dynamics that lipids undergo in membranes are discussed, and the
compositions of plant and bacterial membranes are described. At the end of
this chapter, a brief overview of membrane mimetics relevant to this work is
given.
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2.1 General features of biological membranes
Despite the large variation in composition and the specialized functions of
some membranes and their components, some general properties of mem-
branes can be identified.
Membranes are formed from lipids which are amphiphile molecules con-
sisting of a hydrophilic headgroup and hydrophobic acyl chains (see Fig. 2.3
and 2.4). Driven by the hydrophobic effect, lipids spontaneously phase-separate
from aqueous solvent [125]. Depending on the lipid, the phase formed can be
lamellar, hexagonal, or cubic. The amphiphilic shape hypothesis provides a
means of predicting the phase properties of lipids [65]: if the headgroup’s
cross-sectional area is larger or smaller than the acyl chains’, lipids are of
‘conical’ shape and form a hexagonal or inverted hexagonal phase. Hexag-
onal phases are observed for lysolipids, which are lipids with only one acyl
chain. Inverted hexagonal phases are formed by lipids with small headgroups,
such as 1,2-dimyristoyl-sn-glycero-3-phosphatidylethanolamine (DMPE). If
the headgroup’s and the acyl chains’ cross sectional areas are similar in size,
the lipid is ‘cylindrical’ and prone to form bilayers. This is the case for 1,2-
dimyristoyl-sn-glycero-3-phosphatidylcholine (DMPC) or 1,2-dimyristoyl-sn-
glycero-3-phosphatidylglycerol (DMPG) [49]. A lamellar phase is character-
ized by a lipid bilayer, in which the polar headgroups of the lipids face the
aqueous environment and the hydrophobic acyl chains point towards the cen-
ter of the bilayer. The lipid composition of membranes needs to be such that
the membrane bilayer is stable, but at the same time some curvature stress is
required to provide the membrane with sufficient elasticity to enable processes
such as cell division, membrane fusion, and endo- or exocytosis. This means
that membranes have to consist of lamellar as well as non-lamellar lipids [64].
As described in the next section, biological membranes are in a liquid-
crystalline state providing them with the relevant flexibility to perform their
functions [164]. However, growing evidence suggests that the lateral organiza-
tion of lipids is not homogeneous. In eukaryotic membranes, this is manifested
by lipid rafts, typically enriched in sphingolipids and cholesterol. Rafts allow
for lateral sorting of membrane-associated proteins which means that certain
membrane proteins, mostly involved in signaling and trafficking, are enriched
in these rafts. Therefore, an important biological function is attributed to them
[53; 163]. In prokaryotes, a heterogeneous distribution of lipids has also been
observed, in particular clusters of anionic lipids [7; 20]. For instance, cell poles
in Escherichia coli are enriched in cardiolipin (CL) [134].
Membrane proteins are associated with or embedded into lipid bilayers.
Proteins can be categorized by their mode of membrane interaction; represen-
tatives of some important groups are shown in Fig. 2.1. Peripheral proteins
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In order to maintain a liquid-crystalline phase, organisms adjust the mem-
brane composition to environmental conditions. It has, for instance, been
shown that E. coli alters its acyl chain composition in response to changes
in growth temperature such that the gel-to-liquid phase transition temperature
of the lipid mixture is about 10 ◦C below the growth temperature and non-
lamellar phases are observed approximately 20 ◦C above it [83; 135].
In the liquid-crystalline phase, lipids undergo a broad range of motions on
a wide range of timescales (see Fig. 2.2) [52; 56]. As explained in section
4.3.3, dynamics can be described by an amplitude of motion, represented by
an order parameter, and by a reorientation time of the motion, represented by
a correlation time [69; 119; 120; 196; 197]. Bond oscillations and gauche-
trans isomerizations have characteristic correlation times on the order of pi-
coseconds. These motions are particularly rapid and unrestricted in lipid acyl
chains. Consequently, the hydrophobic region of bilayers has liquid-like prop-
erties [186]. Lateral and rotational diffusion times are on the order of nanosec-
onds [52; 118; 186]. These diffusion processes give rise to the fluidity of the
membrane [125; 186]. While lateral dynamics are rapid, transversal flip-flop
motions, where a lipid flips from one leaflet of the bilayer to the other, have
very long correlation times of hours or days, such that an asymmetric lipid
distribution between the leaflets of a bilayer can be maintained [71; 133].
This is for instance the case in plasma membranes of eukaryotes, or in the
outer membrane of Gram-negative bacteria [49; 184]. Lipid transport across
the membrane is facilitated by a protein class called flippases [213]. Finally,
membranes undergo undulations: thermally agitated, collective lipid motions
that occur on a broad range of timescales between microseconds and seconds.
Undulations relate to the elasticity of the membrane and are thus crucial for
processes that involve shape modifications of the membrane, e.g. endo- and
exocytosis [56].
In lipids, the glycerol backbone displays the highest rigidity, and gauche-
trans isomerizations are restricted in particular for the g1–g2 bond (see Fig. 2.3
for carbon atom nomenclature in lipids). Consequently, the glycerol backbone
undergoes only relatively slow dynamics, such as lipid rotational diffusion and
wobble [18; 52]. Phospholipid headgroups are mainly oriented parallel to the
membrane surface, however, the motional degrees of freedom about the P–O
bonds of the phosphate moiety and about the headgroup carbon-carbon bonds
are higher than for the glycerol carbons, such that other orientations of the
headgroup with respect to the membrane are also possible, albeit less likely
than the parallel orientation [160; 165; 212]. The highest flexibility is ob-
served for the terminal carbon atoms of the acyl chain, in particular the methyl
carbon [165]. Moreover, the probability distribution of the location of the
terminal carbons inside the membrane is broad since their motion is nearly
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2.3.2 Lipopolysaccharides
The composition of the outer leaflet of the outer membrane of Gram-negative
bacteria differs from the other leaflets in that its main lipid component is
lipopolysaccharides (LPS). The outer membrane serves as a permeability bar-
rier for toxic compounds such as antibiotics, while nutrients and ions enter the
periplasmic space through pore proteins located inside the outer membrane
[141]. Moreover, LPS is required for the structural integrity of the outer mem-
brane [210]. For instance, ΔwaaG mutants can only produce a truncated ver-
sion of LPS which moreover lacks phosphorylations of the LPS core (see also
section 3.2). Those mutants are highly susceptible to hydrophobic antibiotics,
and are immobile due to their lack of flagella and pili [105; 145].
A schematic structure of LPS is shown in Fig. 2.5. LPS consists of a
hydrophobic moiety, called lipid A, the recognition of which by the toll-like
receptor 4 leads to the activation of the human immune system. A polysac-
charide connects to lipid A. This polysaccharide is subdivided into inner and
outer core and O-antigen. The inner core is highly conserved within a family,
whereas the outer core shows more variation. The O-antigen is a chain consist-
ing of repeating polyglycan units that show a large structural diversity between
different strains of the same species. LPS is assembled up to the outer core at
the inner leaflet of the inner membrane by gene products encoded by three
operons in the chromosomal waa region. The gmhD and waaA operons en-
code proteins involved in the inner core assembly, and the gene products of the
waaQ operon, among them WaaG, are responsible for the outer core assembly
and for core phosphorylations. Most GTs involved in the LPS core synthesis
are predicted, or have been shown to be peripheral membrane proteins located
at the inner leaflet of the inner membrane, and it has been suggested that they
form a coordinated complex. Following the core assembly, the deep-rough, i.e.
nascent, LPS is transferred to the outer leaflet of the inner membrane where the
O-antigen is attached. LPS is then transported across the periplasmic space and
to the outer leaflet of the outer membrane [72; 193].
2.3.3 Chloroplast membranes
Chloroplast membranes consist of phospholipids (~40 %) and glycolipids
(~60 %, see Fig. 2.4). More than 50 % of the chloroplast lipids are the galac-
tolipids monogalactosyl diacylglycerol (MGDG) and digalactosyl diacylglyc-
erol (DGDG) which contain one and two galactose moieties in the headgroup,
respectively [90]. MGDG and DGDG are thus believed to be the most abun-
dant lipids in nature [62]. Notably, the thylakoid membranes in chloroplasts
consist to 80 % of MGDG and DGDG, with smaller amounts of sulfoquinovo-
syl diacylglycerols (SQDG) and PG [90]. MGDG is a non-bilayer forming
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lipid, whereas DGDG and SQDG are bilayer-forming lipids [47]. This means
that, similar to bacterial membranes, also chloroplast membranes are under
substantial curvature stress which is required for correct function of membrane
proteins and for membrane integrity [201]. Moreover, due to their high abun-
dance in thylakoid membranes, it is believed that galactolipids play a crucial
role in optimal function of photosynthesis [74]. Under stress conditions, in
particular when phosphate is scarce, synthesis of DGDG and SQDG is up-
regulated and the ratio of glycolipids in chloroplast membranes increases to
~80 %. Moreover, under phosphate deprivation, DGDG also replaces phos-
pholipids in extra-plastidial membranes, where it is not found otherwise. In
this way, phosphate is made available for other essential biological processes
[74; 188].
2.4 Membrane mimetics for biophysical applications
A broad range of membrane mimicking systems have been used for biophys-
ical investigations. They range from in vivo membranes to organic solvents.
The degree of complexity-reduction required depends on the phenomenon to
be investigated and on the method used. Within the scope of this work, relevant
membrane mimetics comprise large unilamellar vesicles (LUVs), micelles, and
bicelles. A size comparison is shown in Fig. 2.6.
2.4.1 Large unilamellar vesicles
LUVs are spherical membrane bilayers that have a typical diameter of 100 nm.
Among the mimicking systems used here, they emulate the size and (with im-
portant exceptions) the low curvature of natural membranes best. The mem-
brane composition can be controlled and a broad range of lipid mixtures can be
used to produce LUVs. Moreover, the size and lamellarity can be controlled,
such that it is also possible to make small, giant, and multilamellar vesicles. It
is also possible to produce vesicles with heterogeneous lipid distributions both
laterally, with asymmetric leaflet compositions, and transversal, with lipid do-
main formation [71]. Finally, LUVs provide a closed compartment which al-
lows for maintaining concentration or charge gradients across the membrane
[71; 127; 137]. However, LUVs cannot be used in NMR studies since the lipid
concentration is too low and the aggregates are too big. In solid-state NMR
applications, multilamellar vesicles have been used to study lipid or protein
properties [45]. However, for the purpose of solution-state NMR substantially
smaller membrane mimetics are required.
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bate [8; 124; 183]. In 31P NMR studies of q = 0.5 bicelles, DHPC (detergent)
and DMPC (lipid) show distinct peaks despite identical headgroups. From that
it is concluded that detergent and lipid molecules experience different chem-
ical environments and that they sequester into different regions of the bicelle
[61]. Additional experimental evidence suggests that detergents are located in
a micelle-like environment and are thus found in the bicelle rim whereas lipids
are located in a central bilayer. However, it is unclear how strongly detergent
and lipids are separated [8; 61; 124]. Finally, SANS data are consistent with
the notion that small isotropic bicelles are oblate objects [124].
Since bicelles contain a lipid bilayer, they are better mimetics of natu-
ral membranes than micelles and are still amenable to solution-state NMR
experiments. For some proteins, it has been shown that bicelles are better
mimetics than micelles maintaining the structure and function of these pro-
teins [17; 40; 41]. However, there are also reports that protein structures can
be distorted in bicelles [54]. It can be speculated that this is due to interactions
between the protein and detergents in the bicelle rim. Thus, a frequent concern
when using bicelles as membrane mimetics is whether or not interactions of
the protein with the detergent molecules are significant.
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3. Glycosyltransferases
Whether or not we find what we are seekingIs idle, biologically speaking.—Edna St. Vincent Millay
Glycosyltransferases (GTs) form a vast group of proteins that catalyze the for-
mation of glycosidic bonds between sugar moieties from an activated donor
molecule and a broad range of acceptor molecules [21; 102]. As of Septem-
ber 2017, about 350 000 GTs in 104 families are listed in the carbohydrate-
active enzymes database (www.cazy.org) [25; 38]. Families are defined by
sequence-similarity, and it is expected that all members of the same family
adopt the same three-dimensional fold. Despite the large number of GTs,
only three major folds have been observed, called GT-A, GT-B, and GT-C
fold [22; 34; 121; 122]. Only a handful of GTs that do not adopt any of these
folds have been described in the literature [102].
A crucial structural feature found in both the GT-A and the GT-B fold is the
Rossmann-like fold which is a structural motif that binds nucleotides. Thus,
all nucleotide sugar-dependent GTs adopt either a GT-A or a GT-B fold [179].
The Rossmann fold is a β -strand/α-helix/β -strand motif [70]. In GTs, the
motif is extended to several alternating β -strand and α-helices, giving rise to
a tertiary structure in which the β -strands form a β -sheet that is wrapped by
α-helices. Additionally, insertions of structural elements into the alternating
β/α/β motif are frequently observed [174].
Fig. 3.1 shows representatives of the three major folds observed in GTs.
GT-A GTs consist of two Rossmann-like domains that are tightly connected
to each other. The N-terminal domain binds the nucleotide sugar while the
C-terminal domain binds the acceptor molecule. The function of most GT-A
GTs is dependent on a divalent metal ion which is bound by a Asp-X-Asp mo-
tif. However, examples of GT-A GTs that do not contain the Asp-X-Asp motif
exist. In contrast, GT-B GTs consist of two Rossmann-like domains that are
connected by a flexible linker. The active site is located in the cleft between
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3.2 The glycosyltransferase WaaG
The glycosyltransferase WaaG is involved in the synthesis of the core of LPS
molecules (see section 2.3.2 and Fig. 2.5), which are the main constituents of
the outer leaflet of the outer membrane in Gram-negative bacteria [131; 145].
WaaG has been identified as a potential antibiotic target, since WaaG-deficient
cell lines are immobile and hypersensitive to antibiotics [105]. This is because
cells with a deletion of the gene encoding WaaG (ΔwaaG cells) cannot produce
full-length LPS (see Fig. 2.5). Instead, they contain a truncated version, called
deep-rough LPS. Apart from lacking the outer core and O-antigen, ΔwaaGdeep-rough LPS also lacks phosphate moieties that are crucial for membrane
integrity. Therefore, the protective LPS leaflet is severely compromised in
ΔwaaG cells [210]. Moreover, ΔwaaG cells do not contain flagella and pili
and are thus immobile.
WaaG catalyzes the transfer of a glucose moiety from UDP-glucose to the
nascent chain of LPS. The stereochemistry of the sugar is maintained during
the reaction, i.e. WaaG is classified as a retaining GT [131]. WaaG belongs
to the GT4 family of GTs, which may be the most ancient family of retaining
GTs since it is the only family of retaining GTs found in primitive archaea
[38; 131]. WaaG has been classified as a monotopic GT [2]. This classifica-
tion is based on the subcellular localization of WaaG at the membrane which is
manifested by the need for detergent during purification. However, the crystal
structure of WaaG was obtained from samples that did not contain detergent
[131].
WaaG shows strong structural similarities to other membrane-associated
GT-B GTs, like MurG [75], WaaC [63], and WaaA [158]. It is therefore hy-
pothesized that the mode of membrane interaction of WaaG is in agreement
with the model outlined in section 3.1 [2].
��
4. Methods
The absence of evidence is not the evidence of absence.—Anon.
The size of lipids and proteins is on the order of nanometers. At the same
time, molecular processes may be very rapid—some processes, like gauche-
trans isomerizations, occur on the timescale of picoseconds, other processes,
like chemical exchange, are slower but still of biological relevance. These
properties of lipids and proteins determine the requirements that a biophysical
method must meet to yield insights into the features to be studied. The choice
of the method also depends on the question asked. At times, atomic resolution
is required to get an understanding of a property of interest; other times, molec-
ular resolution is sufficient. For some processes, fast or slow dynamics may be
of interest, in other instances a static picture provides sufficient information.
Each method has advantages, disadvantages, and limitations in terms of time
and spatial resolution and with respect to sample requirements. Thus, to ob-
tain a comprehensive answer to a biological question, ideally a set of methods
should be used. Three methods were employed in the framework of this thesis:
far-UV circular dichroism reveals the secondary structure of proteins and has
relatively low requirements on the sample. Fluorescence methods can be em-
ployed to measure nanosecond dynamics and nanometer distances; however,
they require the presence of a fluorophore. Finally, with solution-state NMR
a broad range of dynamics can be investigated and protein structures can be
determined. However, NMR puts very high requirements on the sample. For
instance, high concentrations are required, the molecule investigated must not
be too large, and it often has to be isotope-labeled.
In the following the methods relevant for this thesis are briefly discussed.
More comprehensive background information on theory and application of the
methods may be found in the references.
��
4.1 Circular dichroism
Far-UV circular dichroism (CD) of the peptide bond provides information on
the secondary structure content of a peptide or protein. CD signals arise due to
the differential absorption of left- and right-handed circularly polarized light
by a chromophore that either contains a chiral center, is covalently bound to a
chiral center, or is located in an asymmetric environment.
In proteins, the far-UV CD spectrum stems mainly from the π → π∗ tran-
sition at 190 nm and from the n→ π∗ transition at 220 nm of the peptide bond,
with minor contributions from aromatic amino acids. Secondary structure el-
ements give rise to different asymmetric environments of the peptide bond
which in turn leads to CD spectra that are characteristic for different types of
protein secondary structure. From a comparison with CD spectra of proteins
of known structure, the secondary structure of a protein can be inferred.
Quantum mechanically, CD is described by the rotational strength R:
Δε ∝ R = ℑ{⟨
Φ f∣∣m |Φi〉〈Φi|μμμ
∣∣Φ f⟩} �= 0 (4.1)
where Δε is the CD signal and mmm and μμμ are the magnetic and electric dipole
moment operators. A CD signal is observed if R is non-zero. This is the case
when mmm and μμμ are non-zero, i.e. when the molecule is a chromophore, and
when they are non-perpendicular as is the case for a chiral chromophore or for
an achiral chromophore in an asymmetric environment [88; 93; 94; 152; 203].
4.2 Fluorescence spectroscopy
Fluorescence spectroscopy is a versatile method to obtain structural and dy-
namic information for a fluorophore. Among others, it can be employed to
determine the rotational correlation time of a fluorophore, to assess whether a
fluorophore localizes in a lipophilic or hydrophilic environment, how deep it
is immersed into a lipid bilayer, and how strongly it interacts with a bilayer
[103]. Techniques to obtain such information are discussed in the following.
4.2.1 Fluorescence of aromatic amino acids
Upon absorption of a photon, an electron in a fluorophore is excited into a
higher electronic state. Within picoseconds it relaxes to the lowest vibrational
level of the first excited electronic state. Fluorescence occurs when the elec-
tron relaxes back into the electronic ground state upon emission of a photon.
The wavelength of the emitted photon is longer than the wavelength of the
absorbed photon which gives rise to the characteristic Stokes’ shift of fluores-
cence excitation and emission spectra. Fluorescence is only one pathway to
��
reach the electronic ground state. Other, non-radiative pathways exist and de-
crease the fluorescence lifetime and intensity [103].
Aromatic amino acids are fluorescent. However, because of their low quan-
tum yield, Phe residues are typically not observable if Tyr or Trp residues are
present [58]. The fluorescence lifetime of Tyr and Trp residues is on the order
of nanoseconds, i.e. molecular dynamics on that timescale can be observed
if the underlying processes affect fluorescence properties of the fluorophores
[103; 154]. Moreover, Trp and Tyr residues are enriched at membrane inter-
faces which is often exploited in fluorescence studies of protein-membrane
interactions [95].
4.2.2 Quenching
Quenching is non-radiative relaxation of an electron in an excited electronic
state to the electronic ground state. Quenching thus competes with radiative
processes, i.e. fluorescence. The relation between loss in intensity and con-
centration of a molecule that quenches fluorescence is described by the Stern-
Volmer equation [172]:
I0
I= 1+KSV [Q] (4.2)
where I is the fluorescence intensity at quencher concentration [Q], I0 is the
fluorescence intensity in the absence of quencher, and KSV is the Stern-Volmer
quenching constant.
This relation holds for collisional quenching, in which fluorescence is
quenched by collisions between the quencher and the fluorophore, and for
static quenching, where quencher and fluorophore form a non-fluorescent com-
plex. If both processes occur, the relation between [Q] and I0/I is not linear
[50].
A number of molecules quench fluorescence of aromatic amino acids
through collisions, among them the water-soluble acrylamide and amphiphilic
nitroxide-labeled lipids. Fluorescence quenching can thus be used to monitor
fluorophore accessibility to aqueous or lipid environments [101].
4.2.3 The parallax method
In the parallax method, the quenching efficiency, and thus accessibility of
nitroxide-labeled quenchers to a fluorophore, is exploited to obtain an immer-
sion depth estimate of the fluorophore inside a lipid bilayer, assuming that the
positions of the quenchers inside the bilayer are known [1; 28; 29]. The method
relies on using two lipids to which a quencher (a nitroxide moiety) is attached,
one with a shallow immersion depth of the quencher (1) and one with a deeper
��
immersion depth of the quencher (2). The distance of the fluorophore from the
bilayer center can be estimated by [28]:
z = Lc1 +
(−L2
21− ln(I1/I2)πC
2L21
)(4.3)
where Lc1 is the distance of the shallow quencher from the bilayer center, L21
is the distance between shallow and deep quencher, C is the quencher concen-
tration in the membrane plane, and I1 and I2 are the fluorescence intensities in
the presence of the shallow and deep quencher, respectively.
In Eq. 4.3 the immersion depth of the fluorophore is ‘triangulated’ from
the quenching efficiencies of the two quenchers, assuming that the quencher
positions are known. The fluorophore needs to be located between the deep
and the shallow quencher to obtain meaningful results and it must be located
within the quenching radius of the quenchers. It has been shown that quencher
or fluorophore dynamics do not affect the immersion depth estimate signifi-
cantly [1].
In order to measure fluorophore immersion depths inside lipid bilayers,
nitroxide-labeled lipids are commonly employed. Nitroxide moieties can be
located in the headgroup and at various positions of the acyl chain and there-
fore immersion depths can be analyzed for the entire width of the membrane.
The positions of the quenchers are known from molecular dynamics (MD)
simulations and various experimental methods, like fluorescence, NMR and
electron paramagnetic resonance (EPR) [101].
4.2.4 Anisotropy
The probability of absorption of plane polarized light by a fluorophore de-
pends on the orientation of the fluorophore’s absorption dipole moment with
respect to the plane of the electric field vector of the electromagnetic wave.
The absorption probability is zero if both are perpendicular to each other and
it is maximum if the electric field vector and absorption dipole moment are
parallel. Following absorption of plane polarized light, the fluorophore rotates
during the fluorescence lifetime and finally emits a photon preferentially along
its emission dipole moment. This leads to a depolarization of the polarized
excitation light. Three effects need to be distinguished: 1. Since absorption
and emission are statistical effects, depolarization always occurs when large
numbers of randomly oriented fluorophores are observed. In the absence of
other depolarization effects, the maximum anisotropy is 0.4. 2. Depolarization
occurs when absorption and emission dipole moments are not parallel. This
gives rise to the fundamental anisotropy r0 of a fluorophore which is smaller
than 0.4. 3. Depolarization occurs because of the rotational reorientation of the
��
fluorophore during its fluorescence lifetime leading to an observed anisotropy
which is lower than the fundamental anisotropy. As a consequence, anisotropy
measurements are sensitive to reorientational dynamics on the timescale of the
fluorescence lifetime of the fluorophore. Furthermore, resonance energy trans-
fer also leads to a decrease in anisotropy [103]. For instance, the fundamental
anisotropy of Tyr residues is approximately 0.3 [104]; however, it is decreased
to 0.19 if resonance energy transfer occurs between neighboring Tyr residues
[66].
The anisotropy r is defined as [103]:
r =I‖ − I⊥
I‖+2I⊥(4.4)
where I‖ and I⊥ are the emitted fluorescence intensities when the emission
polarizer is oriented parallel or perpendicular with respect to the excitation
polarizer.
4.2.5 Stopped-flow fluorescence
In stopped-flow fluorescence experiments, two solutions are rapidly mixed and
the change in fluorescence of a reporter fluorophore is monitored. For sim-
ple binding processes, where one protein molecule binds to a patch of n lipid
molecules, a monoexponential change in fluorescence intensity is expected
with a characteristic rate constant kobs. kobs depends linearly on the lipid con-
centration [L] [139]:
kobs = kd + ka[L] (4.5)
where kd and ka are the dissociation and association rate constants of the bind-
ing. The dissociation constant KD is obtained as the ratio of kd and ka. Stopped-
flow fluorescence spectroscopy thus provides a means to quantify membrane
binding of proteins.
4.2.6 Measurements of the dissociation constant KD
The equilibrium dissociation constant KD between two molecules can be de-
termined by any experimental parameter that distinguishes the bound from the
unbound population. Examples for such parameters are the observed rate con-
stant kobs or the anisotropy, discussed in the previous sections. Moreover, for
Tyr and Trp residues the quantum yield and thus the fluorescence intensity in-
creases when they are moved from a polar to a non-polar environment [58].
The observed intensity is a weighted average between the fluorescence emis-
sion of fluorophores in the bound and unbound populations and can thus be
��
used to obtain KD. Assuming a single binding site, KD is obtained from:
I− I0
I0= Imax
[L]KD +[L]
(4.6)
where I0 is the fluorescence intensity of the unbound molecule, I is the fluores-
cence intensity at lipid concentration [L], and Imax is the fluorescence intensity
when all protein is bound [29; 108].
4.3 Nuclear magnetic resonance
Some atomic nuclei can absorb electromagnetic radiation when placed inside
an external magnetic field. This phenomenon is called nuclear magnetic res-
onance. Only a cursory overview of some aspects of NMR theory relevant to
this thesis can be given. More comprehensive information on basic NMR prin-
ciples, theory, and applications can be found in textbooks by e.g. Keeler [92],
Cavanagh et al. [26] and Kowalewski and Mäler [97].
Protons and neutrons possess an intrinsic quantum property called ‘spin’.
The spin property propagates to nuclei, which are composed of protons and
neutrons. The spin property is described by the spin quantum number S, which
can assume integer and half-integer values. Nuclei with S = 0, e.g. 12C, are not
NMR-active. For other nuclei, like 1H, 15N, 13C, and 31P, S = 1⁄2. Properties
of nuclei with higher spin quantum numbers, e.g. 2H or 14N, are not discussed
here.
Spin 1⁄2 nuclei have two degenerate energy levels, a ‘spin-up’ or α-state and
a ‘spin-down’ or β -state. If placed in an external magnetic field, the degener-
acy of the spin states is lifted and the separation of energy levels is proportional
to the magnetic field strength. The proportionality constant is called the gyro-
magnetic ratio, and it is nucleus-specific. At typical magnetic field strengths
of 9 – 20 T the separation of the energy levels for a proton is on the order of
several 100 MHz, i.e. radiofrequencies can be absorbed by the nuclei.
Radiofrequency pulses can be used to disturb the equilibrium spin state
thus creating non-equilibrium states that provide insights into structural and
dynamic properties of the molecule under investigation. NMR experiments
can provide structural information at atomic resolution and dynamic informa-
tion on a broad range of timescales. However, there are some limitations to the
investigation of biological samples by NMR. The energy difference between
the α- and β -state is low compared to thermal energy, therefore the states are
almost equally populated and thus NMR is a low-sensitivity method. There-
fore, micro- to millimolar sample concentrations are typically required. Fur-
thermore, isotope-labeling is required for many NMR applications, since the
natural abundance of the spin 1⁄2 isotopes 13C and 15N in e.g. organic molecules
��
such as proteins and lipids is 1 % and 0.4 %, respectively. Finally, ‘large’ bio-
logical molecules are a challenging to investigate by NMR. The large number
of atoms that give rise to observable signals leads to spectral crowding and
uninterpretable spectra. Even more importantly, large molecules exhibit broad
peaks due to their slow tumbling, which gives rise to high transverse relaxation
rates. This effect aggravates spectral crowding and may render peaks indistin-
guishable from noise. From a theoretical point of view, the latter problem is
the most serious one. However, experimental methods in combination with
site-specific labeling schemes can alleviate the adverse effects of line broaden-
ing.
In the following, NMR experiments that are relevant for this work are dis-
cussed. First, it is discussed how molecule size and binding can be analyzed
by translational diffusion experiments, then a brief overview of methods to
determine the three-dimensional structure of small peptides is given. Finally,
NMR relaxation methods for determining fast internal motions of molecules
and nanometer distances are presented.
4.3.1 Translational diffusion
Measurements of the translational diffusion coefficient Dt are carried out to
obtain information on the size and association of molecules [116].
The translational diffusion coefficient describes the stochastic thermal mo-
tion of molecules in solution for a system at equilibrium. The diffusion coeffi-
cient depends on the temperature of the system, its viscosity, and the morphol-
ogy of the diffusing particle. For a sphere, the translational diffusion coeffi-
cient is given by the Stokes-Einstein-Sutherland equation [51; 173]:
Dt =kBT
6πηrH(4.7)
where kB is the Boltzmann constant, T is the temperature, η is the viscosity,
and rH is the radius of the sphere, which is also called the hydrodynamic ra-
dius. Diffusion measurements can thus be used to determine particle sizes.
However, rH represents actual dimensions only for a spherical particle. De-
viations between the hydrodynamic radius and actual dimension arise due to
non-spherical shapes [5; 12; 31], due to obstruction of translational diffusion
by particles in solution [9; 89; 166], and because of hydration layers [42]. For
a non-spherical particle, rH differs from the particle’s actual dimensions, since
Dt depends on the morphology of the particle. Yet for particles that are, to a
good approximation, spherical (like micelles, small isotropic bicelles, or glob-
ular proteins) diffusion experiments can provide a good estimate of their size.
Moreover, association of molecules can be studied by diffusion experiments,
��
provide structural constraints for atoms in the molecule are performed. A suf-
ficient number of structural constraints limits the possible molecular confor-
mations to a small set of structures [26; 35; 207].
The most important structural constraints are obtained from nuclear Over-
hauser enhancements (NOEs), an effect that is based on through-space dipole-
dipole interactions between nuclei. If the energy levels of one nucleus are satu-
rated, magnetization is transferred through space via dipole-dipole interactions
to nearby nuclei. The distance dependence of the NOEs is r−6, and typically
distances up to 6 Å between nuclei can be observed. A quantitative interpre-
tation of NOEs is usually not reliable but a sufficiently large set of qualitative
spatial constraints is often sufficient for structure determination [35; 84].
The secondary structure element in which an amino acid is located af-
fects the magnetic environment and thus the chemical shifts of, e.g. Hα , Cα ,
and Cβ atoms. Secondary chemical shifts, i.e. deviations from the random
coil chemical shift for these nuclei in a specific amino acid, are characteristic
for α-helical or β -sheet environments so that the secondary structure element
in which an amino acid is located can be predicted from secondary chemical
shifts. This in turn constrains the rotamer space of the amino acids’ backbone,
giving rise to dihedral angle constraints [11; 37; 161; 202].
Other methods to obtain even more structural constraints from NMR ex-
periments exist. Examples are spatial constraints obtained from paramagnetic
relaxation enhancements (PREs) as described in section 4.3.4, orientational
constraints obtained from residual dipolar couplings [30], and hydrogen bond
constraints [26].
To obtain an NMR structure, all constraints need to be fulfilled simulta-
neously. Based on the amino acid sequence, polypeptide chains are generated
in silico, and the spatial arrangement of the amino acids is altered in an it-
erative way to fulfill the constraints. The quality of the obtained structures,
i.e. their ability to fulfill all constraints simultaneously, is evaluated by an en-
ergy function. Given that there are no major constraint violations and that the
three-dimensional structure is physically allowed, a small set of lowest energy
conformers represents the NMR structure [67; 68; 159; 159].
4.3.3 Dynamics
One particular strength of NMR is that dynamics on a broad range of timescales
can be probed. Timescales relevant for lipid motions range from very rapid
motions, like gauche-trans isomerizations in lipids, that occur on a picosecond
timescale, to very slow motions like spontaneous lipid flip-flops, that typically
happen on timescales on the order of hours (see section 2.2 and Fig. 2.2). Im-
portantly, dynamic events like protein folding, domain motions, and backbone
��
dynamics can be captured by NMR methods [81]. Here, the discussion is lim-
ited to NMR experiments that are employed to measure rapid lipid dynamics
that occur on a pico- to nanosecond timescale. Descriptions of methods that
are used to assess other types of motions are beyond the scope of this thesis.
In NMR, nano- to picosecond dynamics are investigated by measuring re-
laxation rates of non-equilibrium spin states [80; 97]. Relaxation of such states
is mediated by anisotropic interactions of spins which are in turn modulated
by molecular dynamics occurring at appropriate frequencies. For spin 1⁄2 nu-
clei the most important relaxation mechanisms are dipole-dipole interactions
between two spins, chemical shift anisotropy, which is due to the anisotropic
electronic environment at the location of the investigated spin, and paramag-
netic relaxation, which occurs for instance as an additional dipole-dipole relax-
ation mechanism between an unpaired electron and the investigated spin (see
section 4.3.4). Molecular dynamics give rise to random and local disturbances
of the magnetic field that occur at the frequency of the molecular motions.
These disturbances return non-equilibrium spin states to equilibrium and act
thus analogously to the intentional generation of non-equilibrium spin states
by radiofrequency pulses [92].
NMR relaxation parameters are linked to molecular dynamics via the spec-
tral density function J(ω). J(ω) is a probability function that describes the
distribution of motional frequencies ω in a molecule. At the same time, re-
laxation parameters can be expressed in terms of the spectral density function.
For systems with coupled spins, such as 13C–1H bonds in lipids or proteins,
the dominant relaxation mechanism is mediated through dipole-dipole interac-
tions. Then, the longitudinal relaxation rate R1 and the NOE are given by:
R1 =d2
CH10
[J(ωH −ωC)+3J(ωC)+6J(ωH +ωC)] (4.9)
NOE = 1+γH
γC
6J(ωH +ωC)− J(ωH −ωC)
J(ωH −ωC)+3J(ωC)+6J(ωH +ωC)(4.10)
where dCH =−μ0hγCγH/(8π2r3CH) and ωC and ωH are the Larmor frequencies
of 1H and 13C, respectively, μ0 is the vacuum permeability, h is Planck’s con-
stant, γH and γC are the gyromagnetic ratios of 1H and 13C, respectively, and
rCH is the 13C–1H bond length [80; 97].
Even though relaxation is driven by molecular motions, interpretation of
measured relaxation rates in terms of molecular motions is not straightforward
as long as the functional form of J(ω) is unknown. In its simplest form J(ω) is
expressed in terms of a single correlation time τc of a molecular motion, which
describes the rate of that motion:
J(ω) =τc
1+ω2τ2c
(4.11)
��
As discussed in the next section, this form of the spectral density function
can be used in paramagnetic systems to describe the reorientation of the vector
between the nucleus under investigation and a free electron.
The model-free approach allows for a more comprehensive description of
fast molecular motions. The motions are characterized in terms of an order
parameter S that describes the amplitude of motion, and by a correlation time
τ that describes the rate of motion. This approach was originally designed to
describe internal motions in proteins [119; 120] and detergents [69; 196; 197]
but it has been extended to characterize the motions that lipids undergo in
membranes [5; 52]. The extended spectral density function is given by:
J(ω) =
(S2
loc−S2locS2
lip
)τlip
1+ω2τ2lip
+
(1−S2
loc
)τT
1+ω2τ2T
(4.12)
where τT = τlipτloc/(τlip + τloc) and S2loc and τloc are interpreted as the am-
plitude and rate of motion of 13C–1H bond vectors, and S2lip and τlip refer to
the motions of the entire lipid. Both local and lipid motions occur on pico-
to nanosecond timescales and thus mediate spin relaxation. One central as-
sumption in the derivation of Eq. 4.12 is that local and lipid motions occur on
different timescales and are therefore independent of each other [119; 120].
An illustration of a possible interpretation of such motions in lipids is given
in Fig. 4.2. The rate of motion describes the reorientation time of the motional
vector, which is the 13C–1H bond vector for the local motion and the long axis
of the lipid for the lipid motion [97], and the amplitude of motion is described
by the aperture of a cone, in which the motional vector reorients.
4.3.4 Distance measurements using paramagnetic relaxation enhance-ments
In the presence of non-covalently bound unpaired electrons, paramagnetic re-
laxation enhancement (PRE) is a dipole-dipole relaxation mechanism that oc-
curs in addition to nuclear dipole-dipole relaxation when a free electron is
located in proximity of the observed nucleus. PREs are observed for both the
transverse and the longitudinal relaxation rates [36; 143]. The paramagnetic
relaxation enhancement Γ1 of the longitudinal relaxation rate R1 is described
by the Solomon-Bloembergen equation and is at sufficiently high magnetic
field given by [15; 16; 167; 168]:
Γ1 =2
5
( μ0
4π
)2
γ2Cg2
e μ2BS(S+1)J(ω)< r−6 > (4.13)
where μ0 is the magnetic permeability of vacuum, γC is the gyromagnetic ra-
tio of carbon, ge is the electron g-factor, μB is the Bohr magneton, and S is the
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5. Results and Discussion
EYPHKA! num = Δ+Δ+Δ.—Carl F. Gauß
Membrane-associated GT-B GTs have the peculiar property that they catalyze
the formation of a glycosidic bond between a hydrophilic sugar donor sub-
strate and a lipid acceptor molecule. Moreover, peripheral GT-B GTs can ex-
change between a hydrophilic and a membrane environment. The GT WaaG is
a representative of this class of proteins. Since deletion of the gene encoding
WaaG has deleterious effects on the outer membrane of Gram-negative bacte-
ria, WaaG is considered to be a potential antibiotic target [2; 3; 105].
To study membrane interactions of proteins, the system under investiga-
tion needs to be amenable to the methods employed. Solution-state NMR in
particular puts high demands on the sample, one central limitation being that
larger objects give rise to broader linewidths leading to spectral overlap and
eventually to line broadening into noise. Therefore, membrane mimetics need
to be employed, which are amenable to NMR investigations, and which, at the
same time, faithfully mimic natural membranes.
Two central questions are addressed in the following: 1. Can faithful mem-
brane mimetics be produced that are useable in solution-state NMR studies and
what are the characteristics of such systems? 2. How do membrane-associated
GT-B GTs interact with membranes?
5.1 Mimicking plant and bacterial membranes
Membrane mimetics are necessarily compromises between the constraints of
the experimental method and the requirement for faithful emulation of physi-
ological conditions. It is expected that membrane interactions of proteins are
more native-like the closer a lipid bilayer mimics physiological membranes.
For solution-state NMR, small isotropic bicelles constitute such a compromise.
��
Bicelles are small enough to be amenable to solution-state NMR experiments,
but at the same time, the lipid composition can be modified [195].
In Papers I and II, bicelles that mimicked plant and bacterial membranes
were produced and their characteristics analyzed. These characteristics are:
1. size of the bicelle, 2. lipid composition, and 3. pico- to nanosecond lipid
dynamics. Furthermore, in Paper III a method was developed to determine the
immersion depths of lipid carbon atoms in membranes.
5.1.1 Size of bicelles
The bicelle size can be determined from the translational diffusion coefficient
of lipids which have a negligible monomer solubility in water. As discussed
in section 4.3.1 the hydrodynamic radius obtained from translational diffu-
sion measurements reflects the radius that a sphere with the observed diffusion
coefficient would have, i.e. diffusion measurements do not report on the mor-
phology of the investigated particle and dimensions are approximately accurate
only if it is roughly spherical in shape.
In Paper I, the diffusion coefficients of bicelles enriched in the galactolipids
MGDG and DGDG were determined. It was found that enrichment in both
MGDG and DGDG leads to an increase in bicelle size when compared to bi-
celles that only contain DMPC as the lipid component. The difference is at-
tributed to the bulkier headgroup size and the acyl chain composition (see Fig.
2.4). The dominant species of MGDG and DGDG contained 18:3-16:3 acyl
chains while DMPC acyl chains are saturated and 14 carbons long. It was also
observed that DGDG containing bicelles were on average smaller than MGDG
containing bicelles despite the bulkier headgroup of DGDG. The most signif-
icant difference between DGDG and MGDG is that the former is a bilayer
forming lipid while the latter is not [47]. Instead, MGDG forms an inverted
hexagonal phase. It is therefore speculated that the difference in hydrodynamic
radius actually reflects morphologic differences. Due to the bilayer forming
propensities of both DMPC and DGDG a lipid mixture of these two lipids
may still to a good approximation conform to the ideal bicelle model (see Fig.
2.7). On the other hand, the lipid bilayer in bicelles that contain MGDG might
be under considerable stress, leading to morphological changes of the bicelle
which result in smaller diffusion coefficients of MGDG containing bicelles
compared to DGDG containing bicelles.
5.1.2 Lipid composition of bicelles made from E. coli phospholipids
In Paper II, E. coli inner membranes were mimicked using E. coli phospho-
lipid extract as the lipid component in bicelles. The phospholipid composition
was analyzed by 31P-NMR, in which distinct peaks for all three phospholipids
��
that are found in E. coli (PE, PG, and CL) were observed. When cultures were
grown at 30 ◦C, a phospholipid composition was obtained that corresponds to
previously published data [200]. PE lipids are most abundant (~80 %) and the
amount was unchanged when cells were grown at 20 ◦C. In contrast, the com-
position of anionic lipids changed at lower growth temperature. This effect is,
however, not attributed to the difference in growth temperature but rather to the
difference in growth phase. Cells were harvested after 24 hours, at which point
cultures grown at 20 ◦C are in the exponential phase, whereas cultures grown
at 30 ◦C are in the stationary phase. In the stationary phase, CL is formed
at the expense of PG [73]. The observed increase in CL, which is accom-
panied by a decrease in PG in cultures grown at 30 ◦C compared to cultures
grown at 20 ◦C, is therefore a growth phase effect. At growth temperatures
above 30 ◦C, the phospholipid composition changed dramatically, an effect
that can be attributed to the temperature-dependent expression of the phos-
phatidylserine synthase PssA, an essential enzyme for the synthesis of PE. At
higher temperatures, the expression of PssA is impaired leading to a decrease
in PE content in the membranes. This effect is specific for the AD93 E. colistrain used in this study [44; 91].
The acyl chain composition of the E. coli phospholipid extract was mon-
itored by 13C-NMR and it was found that the amount of unsaturated lipids
increases at lower growth temperature. E. coli bacteria regulate membrane flu-
idity via acyl chain modifications and thus maintain the membrane in a liquid-
crystalline phase [43; 110]. An increase in amount of unsaturations decreases
the phase transition temperature, i.e. the observed change in amount of un-
saturations is a temperature effect. Finally, PE can be depleted altogether in
membranes by using a strain that cannot express PssA [44].
5.1.3 Rapid dynamics in bicelles
In Papers I and II, lipid dynamics that occur on a pico- to nanosecond timescale
were analyzed for bicelles containing galactolipids, and bicelles consisting of
E. coli phospholipid extract, respectively. As described in section 4.3.3, the
model-free approach allows for obtaining an amplitude of motion, given by
the squared order parameter S2, and a rate of motion, given by the correlation
time τ , for 1H–13C bonds in lipids (S2loc and τloc) and for the entire lipid (S2
lipand τlip).
While S2loc parameters for MGDG and DGDG headgroup carbons were on
the same order as for glycerol carbons, S2loc parameters were somewhat lower
for headgroup carbons of PE and PG in E. coli bicelles. At the same time,
correlation times for galactolipid headgroup carbons were about one order of
magnitude larger than for headgroup carbons of E. coli phospholipids. Thus,
��
the rigidity of 1H–13C bonds in the headgroup depends on the size and struc-
ture of the headgroup. Larger headgroups show an increased rigidity and the
same is true for the rigid ring structure of galactose headgroups. The glycerol
region of the lipid is rigid for all lipid types and correlation times are several
hundred picoseconds. It should be noted that the dynamic parameters are not
entirely reliable for the glycerol and the galactolipid headgroup carbons since
the assumption of timescale separation (see section 4.3.3) between local and
lipid motions is violated. In consequence, the order parameters and correlation
times are over- and underestimated, respectively [119; 120].
The rigidity of the 1H–13C bonds in the upper acyl chain is comparable to
the carbons in smaller headgroups like PE or PG. The rigidity decreases down
the acyl chain to almost unrestricted motions of the terminal methyl carbons.
This trend is broken by acyl chain modifications. While a slight local increase
in rigidity is observed for unsaturated carbon atoms, S2loc is increased signifi-
cantly for carbons in cyclopropane rings.
The model-free approach can also provide order parameters and correla-
tion times that report on lipid dynamics. While these parameters were pre-
sumed and fixed in Paper I, they were treated as variables in Paper II. For lipids
in E. coli bicelles, S2lip was determined to be about 0.5 and the lipid correlation
time was found to be 800 ps. The rigidity is thus on the upper end of what
has been reported for other lipids [109], while the correlation time is com-
parable to correlation times found by EPR for lipids in the liquid-crystalline
phase [24; 130]. It can therefore be concluded that the bicelle membrane is in
a liquid-crystalline phase similar to the in vivo situation.
5.1.4 Immersion depths of carbon atoms in membranes
Immersion depths of molecules in membranes are an important parameter
for the understanding of biological processes that occur in or at membranes.
PREs can be employed to measure immersion depths since they are distance-
dependent, as described in section 4.3.4. They are particularly suited for mem-
brane immersion depth measurements since the distance over which PREs are
effective (up to 30 Å) [82] is on the same order as the thickness of a typical
membrane, which is about 40 Å [98–100; 146]. In Paper III a method was
developed to measure the immersion depths of lipid carbon atoms in bicelles
using PREs of the longitudinal relaxation rate R1.
Bicelles were doped with four different paramagnetically labeled lipids
that contained paramagnetic moieties in the headgroup and at various posi-
tions in the acyl chain, and PREs were measured for lipid carbon atoms. The
carbon atom PREs reflect the localization with respect to the paramagnetic
moieties inside the membrane and thus a qualitative estimate of the immersion
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depths of the carbon atoms can be obtained [33; 128]. For example, when the
paramagnetic label was located at the far end of the acyl chain, the observed
PREs were strongest for the terminal acyl chain carbon atoms and weakest for
headgroup carbon atoms.
Carbon immersion depths were quantified employing immersion depths
and immersion depth distributions of the paramagnetic moieties, which are
known from MD simulations [101]. Experimental PREs were measured in the
presence of four different paramagnetically labeled lipids, which provided four
distance constraints for the immersion depth of each carbon atom. Theoretical
PREs were computed for a simple bicelle model where each lipid was rep-
resented by a cylinder and the detergent molecules in the rim were neglected.
Carbon atoms were placed at random immersion depths inside the center of the
lipid cylinder and one lipid was randomly chosen to carry the paramagnetic
label, located at a position obtained from the known immersion depths and
immersion depth distributions. From this model, theoretical PREs were cal-
culated using Eq. 4.13 by summing up all contributions from identical carbon
atoms. The theoretical PREs depend only on the carbon immersion depths1,
i.e. from a least square fit of the theoretical to the experimental PREs, the
carbon immersion depths were obtained. The fitting algorithm did not provide
accurate immersion depths for only one simulated bicelle, since the measured
PREs were ensemble averages, i.e., the conformational space of the possible
locations of the paramagnetic label had to be sampled. Therefore, the fitting
procedure was repeated for hundreds of bicelles. The theoretical PREs and fit-
ted carbon immersion depths were the means of the in silico bicelle ensemble.
Using this approach it was found that the headgroups in zwitterionic and E.coli bicelles are approximately 20 Å away from the bilayer center, the glycerol
carbons are roughly located at 16 Å from the bilayer center, and the first acyl
chain carbons are right below. The terminal acyl chain carbons are closest to
the bilayer center and acyl chain modifications (unsaturations and cyclopropa-
nations) in E. coli bicelles are about 8 Å away from the bilayer center.
Somewhat crude assumptions were made in the calculation of theoretical
PREs. For instance, a simple form of the spectral density function (see Eq.
4.13) was assumed, the bicelle model consisted of cylindrical lipids, and de-
tergents were neglected altogether. Nonetheless, the results are in good agree-
ment with results obtained by other methods [77; 99; 138; 144; 165].
The method presented in Paper III provides a means to quantitatively de-
termine immersion depths in membranes with an accuracy of 3 – 5 Å, and can
be extended to immersion depth measurements of membrane proteins or other
membrane-associated molecules.
1Eq. 4.13 has the form Γ1 =Cr−6 where C is a constant. Since not all parameters that constitute
C were known, C was treated as an additional, global variable in the fitting procedure.
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5.1.5 Bicelles as a plant and bacterial membrane mimetic
Solution-state NMR, in particular, puts high demands on samples of biological
molecules, as the investigated molecules must not be too big. Since natural
membranes are large aggregates of proteins and lipids, membrane mimetics
that are sufficiently small are required to conduct NMR experiments.
Bicelles maintain a lipid bilayer which is dissolved in aqueous solution by
detergents located in the rim of the bicelles. In this work, it was shown that
the lipid composition can be adjusted in such a way that it faithfully mimics
plant or bacterial membranes. A major limitation for the production of bicelles
is that the lipid mixture must be prone to form bilayers. It was, for instance,
not possible to produce bicelles from synthetic lipids with the same phospho-
lipid composition as E. coli inner membranes. This indicates that a delicate
equilibrium between bilayer and non-bilayer forming lipids exists in natural
membranes. However, by choosing strain, growth temperature, and time point
of cell harvest, the final lipid composition of bicelles can be tuned to some
degree.
Properties of the bicelle, the bicelle lipids, and individual carbons of the
lipids can be investigated in some detail using NMR methods. Analysis of the
dynamics showed that the lipids are in a liquid-crystalline phase, and that lipid
dynamics display a large degree of heterogeneity along the membrane normal,
from very restricted local motions of the glycerol backbone to almost unre-
stricted motions at the terminus of the acyl chains. Moreover, from measure-
ments of carbon immersion depths, lipids could be localized inside the mem-
brane. As expected, their headgroup and glycerol carbons are located at or near
the surface of the membrane, and the acyl chains point towards the center of the
membrane. At the same time, the uncertainties in the immersion depths esti-
mates were high, especially for unsaturated and methyl carbons. This indicates
a broad distribution of these carbons inside the membrane. Taken together the
experimental evidence suggests that the lipids in bicelles behave similarly to
what can be expected for lipids in physiological membranes.
There are, however, drawbacks when using bicelles for protein-lipid in-
teraction studies. Firstly and most importantly, often, it cannot be ruled out
that the protein under investigation interacts with the detergent molecules in
the rim. Such interactions may, similar to interactions with detergent micelles,
distort the structure and impair the function of the protein. Secondly, the mor-
phology of the lipid bilayer may be affected when non-bilayer forming lipids
are introduced into the lipid mixture. Finally, it is still unclear to what degree
and under which conditions detergents and lipids segregate in bicelles into a
detergent rim and a lipid bilayer region.
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5.2 Monotopic and peripheral glycosyltransferases
Based on studies of monotopic and peripheral GTs a model of their mem-
brane interaction has been proposed (see Fig. 3.2). In this model, the N-
terminal domain is anchored to the membrane while the C-terminal domain
binds less strongly to the membrane, undergoing an unspecified motion that
is required to accommodate the donor and acceptor molecules at the active
site in the cleft between the two domains [2; 57]. However, in membrane-
associated GTs this motion has not yet been observed. Moreover, it is not
known whether membrane-associated GTs bind certain lipids specifically and
membrane affinities are largely unknown.
Here, the applicability of this model to the GT WaaG was tested and the
open questions outlined above were addressed.
5.2.1 Anchoring of WaaG to membranes
To study the anchoring mechanism of WaaG to membranes, a potential mem-
brane interacting region of WaaG (MIR-WaaG) was identified and studied in
Paper IV. The selection of the segment was based on three properties: 1. it is
exposed in the crystal structure, 2. it contains positively charged residues, and
3. it contains Tyr residues that have previously been shown to be important for
membrane interaction [95].
It was found by fluorescence anisotropy and quenching studies that MIR-
WaaG interacts with vesicles enriched in anionic charge but does not bind
to zwitterionic vesicles. As evidenced by CD spectra, MIR-WaaG adopts an
α-helical structure in anionic vesicles, as expected from the crystal structure
[131], whereas it is random coil in buffer and in zwitterionic vesicles. More-
over, employing the parallax method (see section 4.2.3), the average immer-
sion depth of the Tyr residues could be determined to be ~16 Å. They are thus
located at a similar membrane immersion depth as the lipid glycerol backbone
[99].
To get a more detailed understanding of the structure of MIR-WaaG and
the localization of its residues within a membrane, the NMR structure was
determined in dodecylphosphocholine (DPC) micelles. The NMR structure
of MIR-WaaG agrees remarkably well with the equivalent segment in the
crystal structure. The localization of individual residues was determined
qualitatively by investigating the peak intensity reduction upon addition of
the water-soluble paramagnetic molecule Gd3+ diethylenetriamine pentaacetic
acid-bismethylamide (Gd(DTPA-BMA)) [10; 143; 211]. The intensity reduc-
tion is stronger for solvent exposed residues than for residues located inside the
DPC micelle. It was found that a central part of MIR-WaaG is located inside
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the micelle while its N- and C-terminus are either solvent exposed or at the mi-
celle surface. These data were supported by hydrogen exchange experiments
[76; 107] that showed NOE cross-peaks between Tyr residues and water pro-
tons and hydrogen exchange between water and guanidinium protons in Arg
residues. From this information, the localization of MIR-WaaG in micelles
could be derived (see Fig. 5.1a). Based on the combined data, a model of the
membrane interaction of WaaG was proposed (see Fig. 5.1b), in which the N-
terminal domain anchors WaaG to the membrane via electrostatic interactions
between MIR-WaaG and anionic lipids. The interactions are potentially en-
hanced by Tyr residues located at the membrane’s interface and hydrophobic
residues pointing towards the center of the membrane.
5.2.2 Membrane interaction of full-length WaaG
WaaG has been suggested to be a monotopic membrane protein [2], and, as
discussed above, the N-terminal domain anchoring segment MIR-WaaG inter-
acts with lipids in a charge-dependent way. However, no information on the
membrane affinity of full-length WaaG was available. Therefore, these affini-
ties were determined in Paper V. Additionally, quantitative membrane-binding
studies could potentially reveal lipid-specificity of the binding as well as do-
main dynamics.
The interactions between full-length WaaG and LUVs of varying lipid
compositions were investigated by stopped-flow fluorescence. To be able to
use fluorescence spectroscopy, Trp residues were introduced into exposed re-
gions of either the N- or the C-terminal domain in positions likely to be affected
by membrane binding of WaaG. Upon interaction with lipids, the environment
of the Trp residues changed leading to a change in fluorescence intensity that
could be observed in real time. The change in fluorescence intensity is charac-
terized by an observed rate constant kobs, which depends on the lipid concen-
tration. From measurements of kobs at various lipid concentrations, association
and dissociation rate constants were obtained [139].
As expected from previous studies [136], WaaG does not interact with
zwitterionic vesicles, but it does interact with vesicles enriched with either
of the two anionic lipid species found in E. coli inner membranes, PG and CL.
However, WaaG does not preferentially bind either of these lipids; however,
the affinity is determined by the surface charge density of the vesicles. WaaG’s
affinity for vesicles made of E. coli phospholipid extract with 20 % anionically
charged lipids [114; 200], is lower than for vesicles containing 40 % synthetic
anionic lipids. Moreover, no difference in affinity was observed between WaaG
mutants that contained a Trp-mutation in either the N- or C-terminal domain.
In principle, different affinities of the flexibly linked domains, and thus mo-
��
tions between the domains or with respect to the membrane, could be observed
by this assay as long as the Trp environment is affected by the motions. This
would be the case if the C-terminal domain performs an ‘up-and-down’ move-
ment while the N-terminal domain is permanently anchored. However, equal
affinities, as observed here, do not necessarily mean that the two domains do
not move with respect to each other. For instance, it has been hypothesized
that the C-terminal domain performs a rolling motion [2; 57], which might not
change the environment of the Trp residue. Moreover, the C-terminal domain
might only move in the presence of the natural nascent LPS substrate.
The membrane affinities of very few membrane-associated GTs have thus
far been reported. Among them are the monotopic GT MGS which binds mem-
branes with nM – pM affinity [110] and the peripheral GT PimA which binds
membranes with μM affinity [153]. It is found for WaaG that its affinity for
anionic membranes is on the order of μM, i.e. WaaG seems to be a peripheral
membrane protein rather than a monotopic one, as previously suggested. It
is, however, unclear whether this observation is also true in vivo, since con-
vincing data on the in vivo concentration of WaaG have not been reported
[157; 169; 192].
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6. Conclusions
The travails of the mountains lie behind us.Before us lie the travails of the plains.—Berthold Brecht
One of the aims of this work was to increase the understanding of the func-
tion of a peculiar class of proteins—membrane-associated GT-B GTs. The
interest in membrane-associated GTs has two aspects. Firstly, membrane-
associated GTs operate at the membrane surface and their function is to bring
two molecules with very different physical properties, a hydrophilic donor
sugar and a lipid acceptor molecule, into proximity so that a glycosidic bond
can be formed. At the same time, many of these GTs themselves seem to be
peripheral membrane proteins, i.e. hybrids between soluble and permanently
membrane-bound proteins. Secondly, some peripheral or monotopic GTs are
involved in the synthesis of glycolipids like LPS. Impairment of their func-
tion can be lethal to bacteria and these GTs are therefore promising antibiotic
targets. This demonstrates the relevance of another focus of this work that re-
volves around another peculiar class of biological molecules—lipids. The im-
portance of a properly ‘tuned’ and functioning membrane to a living organism
cannot be underestimated. Defects in the lipid composition of the membrane,
in the structure of the lipids, or of the lipid dynamics can lead to severe mal-
function of membrane-related biochemical processes.
The research described in this thesis shed light on the mode of membrane
interaction of a representative of the class of membrane-associated GT-B GTs,
WaaG. Electrostatic interactions play a key role in binding the N-terminal do-
main of WaaG to membranes. WaaG does not discriminate between anionic
lipids, but it senses the surface charge density of the membrane. The affinity
for membranes has been reported for very few GTs and here we find that the
membrane affinity of WaaG is rather weak; it may thus be categorized as a
peripheral membrane protein.
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Lipids undergo a broad range of motions inside membranes. Rapid lipid
motions were probed here by NMR relaxation in bicelles that faithfully mim-
icked the membrane composition of bacterial and plant membranes. The glyc-
erol backbone is rather rigid and therefore mostly subjected to lipid wobbling
and rotational reorientation. Motions in the distal lipid regions, in particular at
the acyl chain terminus, are much more rapid and dominated by trans-gauche
isomerizations and bond reorientations. Lipid headgroup flexibility depends
on the size of the headgroup. Small headgroups undergo more rapid and less
restricted motions than larger headgroups. Furthermore, by employing PREs
an ‘atomic ruler’ was developed that allows the measurement of the immersion
depth of lipid carbon atoms in membranes.
The studies of lipid properties in realistic membrane mimetics and the in-
vestigations of the membrane interactions of WaaG were largely conducted
separately. However, a synthesis of the approaches would be desirable to get a
better understanding of WaaG’s membrane interaction in more realistic mem-
branes. In particular, it would be desirable to apply NMR techniques, which
are particularly suited to understand dynamic processes, to systems containing
WaaG and natural E. coli lipids. However, the resulting aggregates are very
large, even if small isotropic bicelles are employed so that less conventional
NMR techniques are required to conduct such investigations. One approach
would be to site-specifically label WaaG, and investigate properties of the la-
bel or the effects the label has on lipids in the membrane. Such studies could
lead to a more profound understanding of how peripheral GT-B GTs interact
with lipids and their substrates. On a larger scale, it is not understood if and
how different GTs act together at the membrane surface to synthesize LPS.
Moreover, some GT-B GTs seem to undergo large conformational changes
upon membrane and substrate binding, making this class of proteins a worth-
while target for further studies. Finally, multi-resistant bacteria have become
an imminent threat to society. Therefore, it is absolutely crucial to identify new
targets for antibiotics, to understand their function, and to impair it. WaaG and
other LPS-synthesizing proteins are such targets, and it is therefore worthwhile
to continue the effort to understand their mode of action at the membrane sur-
face.
As mentioned in the introduction, the scientist’s endeavor is not unlike the
mountaineer’s venture. At this point of the thesis a mountain range has been
negotiated and before us lie the plains that promise rest. Or do they? In the
distance, new undiscovered mountain ranges loom—and the grasslands are un-
charted territory. Some blank spots on the map of knowledge may have been
filled in during this endeavor. Yet, the mountaineer pushes on, and the scientist
asks new questions.
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Populärvetenskaplig Sammanfattning
Som när en Grip i mörkret genom öcknarMed vingadt lopp kring fält och sanka dalarFörföljer Arimaspen, som försåtligtHar hans förtrodda skatt från honom röfvat.—John Milton, Det förlorade paradiset
Man kan tänka sig proteiner som små, biologiska maskiner – de minsta maski-
nerna som finns, endast en miljondel av en millimeter i storlek. Tillsammans
med fetter och kolhydrater är de livets viktigaste byggstenar. Dessa små par-
tiklar kallas också för molekyler. Proteiner är mycket komplicerade molekyler,
som är sammansatta av tusentals atomer på ett komplicerat sätt till en välord-
nad struktur som katalyserar, det vill säga genomför, kemiska reaktioner med
en oerhörd precision. Ett protein kan katalysera en kemisk reaktion, som an-
nars skulle ta dagar eller veckor, på bråkdelen av en sekund. Något som är även
mer förvånande är att proteiner kontrollerar alla processer som sker i en cell,
så att livet blir möjligt. Utan proteiner finns det inget liv.
Men för att liv ska vara möjligt behövs det mer än ett par proteiner. En an-
nan viktigt sorts molekyler är lipider, som är en särskild grupp av fetter. Lipider
är den viktigaste beståndsdelen i membraner, som man kan föreställa sig som
husväggar. Så som husväggar skyddar människor från väder, omger membra-
ner celler och skyddar dem från deras omgivning. Membraner skapar en inne-
och en utemiljö. På cellens insida finns det livsviktiga proteiner och andra mo-
lekyler. Från utsidan får cellen näring, t. ex. kolhydrater, men där finns också
giftiga molekyler, som cellen måste skydda sig mot.
Membraner är uppbyggda av lipider. Lipiderna måste framställas och här
kommer proteiner in i bilden. Speciella proteiner finns för att framställa olika
lipider som är nödvändiga för att bygga upp ett membran. Ett av dessa prote-
iner har studerats i denna avhandling. Det kallas för WaaG och kommer ifrån
bakterien Escherichia coli. Vi undersökte hur proteinet utför sitt arbete i bak-
terien, det vill säga i en organism som består av en enda cell.
WaaG katalyserar ett enda, men mycket viktigt steg, i tillverkningen av en
speciell sorts lipider i bakterien. Detta är ett skäl till, varför vi vill förstå hur
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proteinet fungerar. Som bekant finns bakterier som orsakar sjukdomar och mot
dessa finns antibiotika som dödar bakterien. Ett sätt att döda bakterierna är att
förstöra proteinerna som är involverade i uppbyggandet av membranen, det vill
säja t. ex. WaaG. Bakterierna kan inte leva med ett skadat eller förstört mem-
bran. För att kunna inhibera ett protein som t. ex. WaaG måste vi dock förstå
hur det fungerar. Men det finns ytterligare ett skäl till, varför vi är intresserade
av WaaG: Proteinet är inte bara inblandat i uppbyggandet av cellens membran,
det har egenskapen som gör att det liknar ett mytiskt fabeldjur: WaaG är en
grip.
Vem är gripen?
De antika grekerna berättar historier om en varelse, som har ett lejons kropp
och svans, och en örns huvud och vingar: en grip. Gripar är luftens och landets
kreatur. Tack vare den särskilda kombinationen av örn- och lejonegenskaper,
känner de sig hemma i båda omgivningarna. WaaG är en faktiskt existerande,
mikroskopisk analogi till en grip – ett kreatur av två världar. Tack vare dess
särskilda kombination av egenskaper känner proteinet sig hemma i två omgiv-
ningar som inte kunde vara mer olika varandra: vatten och lipider, som är en
undergrupp av fetter.1 Vatten och fett går inte att blanda. Det vet alla som har
någonsin försökt att diska en infettad stekpanna med vatten.
Hur kan ett protein trivas i båda omgivningarna? Detta är svårt att före-
ställa sig och därför undersökte vi WaaG noggrannare och hittade ett område
i proteinet, som kan docka som ett ankare till membranen (se figur 6.1). Detta
ankare orsakar inga problem när WaaG befinner sig i vatten, men så fort det
finns en lipidmembran i närheten dockar ankaret i WaaG till membranet.
Var lever gripen?
I den antika litteraturen finns det olika teorier om var gripar lever. Några för-
fattare tror att de vistas i fjärran länder i norden, andra misstänker att de bor
i fjärran östern. De flesta författare är dock överens om att griparna attraheras
starkt av guld. De påstås faktiskt vara väktare av otroliga skatter.
WaaG attraheras i sin egen del av en skatt: av vissa lipidmolekyler. Dessa
lipider måste vara negativt laddade, så att ankaret, som är positivt laddat, bin-
der till dessa lipider och på detta sätt till membranen. Ankaret fungerar inte
utan negativt laddade lipider.
1Ordet “lipid” härstammar från det grekiska ordet för fett: λ íπoς .
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Existerar gripen på riktigt?
WaaG är alltså ett hybridprotein, som gillar både vatten och lipider, men det
måste befinna sig vid ett lipidmembran för att genomföra sitt jobb, vilket är
att bygga nya lipider. WaaG ankrar till negativt laddade lipider i membranen
genom ett positivt laddad ankare. Och medan detta händer, dansar lipiderna sin
dans och nya framställas om och om igen.
Alla dessa observationer och resultat bygger på undersökningar som gjor-
des i ett provrör. Men händer allt det här även i levande bakterier? Vi kan med
ganska stor säkerhet säga att några av iakttagelserna även försiggår i levande
celler. Lipider rör sig i alla celler. WaaG binder elektrostatiskt till membraner.
Men binder det jämnt och under alla förhållanden? Rör sig WaaG också? Det
vet vi inte än. Vi förstår nu bättre hur WaaG och lipiderna fungerar tillsam-
mans, men varje svar leder till nya frågor. Vetenskap handlar om att utvidga
kunskap och att få ett större underlag för en bättre förståelse av världen om-
kring oss. Med denna avhandling har kunskapen om en grupp proteiner som
beter sig mycket egendomligt utvidgats. Dessutom förstår vi bättre, hur lipi-
der rör sig och hur de interagerar med proteiner. Detta betyder att ännu en vit
fläck på kunskapens karta, som nämndes inledningsvis i denna avhandling, kan
suddas ut.
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Populärwissenschaftliche
Zusammenfassung
Wie wenn ein Greif mit schnellbeschwingtem LaufBergauf, bergab, durch Wüste, Wald und MoorDen Arimasp, der ihm, ein schlauer Dieb,Sein streng bewachtes Gold entwandt, verfolgt.—John Milton, Das verlorene Paradies
Proteine sind Eiweiße und man kann sie sich wie kleine, biologische Maschi-
nen vorstellen – die kleinsten Maschinen, die es gibt, nur ein Millionstel ei-
nes Millimeters klein. Zusammen mit Fetten und Kohlenhydraten sind sie die
wichtigsten Bausteine des Lebens. Diese winzigen Teilchen nennt man auch
Moleküle. Proteine sind sehr komplizierte Moleküle, die sich auf eine kom-
plizierte Art und Weise aus Tausenden von Atomen zu einer Struktur zusam-
mensetzen, die mit großer Präzision chemische Reaktionen katalysieren, d.h.
durchführen, kann. Ein Protein kann eine chemische Reaktion, die ansonsten
Tage oder Wochen dauern würde, in Bruchteilen einer Sekunde katalysieren.
Mehr noch: Proteine kontrollieren alle Prozesse, die in einer Zelle vor sich ge-
hen, sodass Leben möglich wird. Ohne Proteine gäbe es kein Leben.
Allerdings wird zum Leben noch mehr benötigt als nur Proteine. Ein weite-
re wichtige Klasse von Molekülen sind Lipide, die eine besondere Gruppe von
Fetten sind. Lipide sind der wichtigste Bestandteil von Membranen, die man
sich wie Hauswände vorstellen kann. Genauso wie Hauswände Menschen vor
dem Wetter schützen, umgeben Membranen Zellen und schützen sie vor ihrer
Umgebung. Membranen schaffen ein „Innen“ und ein „Außen“. Im Inneren
der Zelle befinden sich lebenswichtige Proteine und andere Moleküle. Von au-
ßerhalb bezieht die Zelle Nahrung, z.B. Kohlenhydrate, aber dort befinden sich
auch giftige Moleküle, vor denen sich die Zelle schützen muss.
Membranen bestehen aus Lipiden. Lipide müssen gebaut werden und an
dieser Stelle kommen Proteine ins Spiel. Es gibt spezielle Proteine, die Lipide
herstellen. Eines dieser Proteine wurde hier untersucht. Es wird WaaG genannt
und findet sich in dem Bakterium Escherichia coli. Wir haben untersucht, wie
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das Protein seine Arbeit in Bakterien, also Lebewesen, die aus nur einer einzi-
gen Zelle bestehen, verrichtet.
WaaG katalysiert einen einzigen, aber äußerst wichtigen Schritt in der
Produktion von bestimmten Lipiden in Bakterien. Das ist ein Grund, weswe-
gen wir verstehen wollen, wie es funktioniert. Manche Bakterien verursachen
ernsthafte Krankheiten. Antibiotika heilen solche Krankheiten, indem sie die
Bakterien töten. Eine Möglichkeit, dies zu tun, ist, die Proteine zu zerstören,
die mit dem Aufbau von Membranen zu tun haben, also zum Beispiel WaaG.
Mit einer beschädigten oder zerstörten Membran können Bakterien nicht le-
ben. Bevor wir die Funktion von WaaG unterbinden können, müssen wir je-
doch verstehen, wie es funktioniert. Es gibt aber noch einen weiteren Grund,
aus dem wir an WaaG interessiert sind: Es ist nicht nur in den Aufbau der
Zellmembran involviert, es hat gleichzeitig Eigenschaften, die es einem my-
thischen Fabelwesen ähneln lassen: Es ist ein Greif.
Wer ist der Greif?
Die alten Griechen erzählen Geschichten von einem Wesen, das den Körper
und Schwanz eines Löwen hat und den Kopf und die Flügel eines Adlers: der
Greif. Greife sind Kreaturen der Luft und des Landes. Dank einer speziellen
Kombination von Adler- und Löweneigenschaften, fühlen sie sich in beiden
Umgebungen zu Hause. WaaG ist eine tatsächlich existierende, mikroskopi-
sche Analogie des Greifs – eine Kreatur zweier Welten. Dank einer speziellen
Kombination von Eigenschaften fühlt es sich in zwei Umgebungen zu Hause,
die unterschiedlicher nicht sein könnten: Wasser und Lipide, die eine Unter-
gruppe der Fette sind.1 Wasser und Fett kann man nicht mischen. Das weiß
jeder, der schon einmal versucht hat, eine fettige Pfanne mit Wasser zu säu-
bern.
Wie kann sich ein Protein in beiden Umgebungen wohlfühlen? Dies ist
schwer vorstellbar und deshalb haben wir uns WaaG genauer angeschaut und
einen Bereich gefunden, der wie ein Anker an die Membran andocken kann
(siehe Abbildung 6.2). Dieser Anker verursacht keine Schwierigkeiten, wenn
WaaG sich in Wasser befindet, aber sobald eine Lipidmembran in der Nähe ist,
dockt er WaaG an die Membran an.
Wo lebt der Greif?
In der antiken Literatur gibt es verschiedene Theorien dazu, wo Greife leben.
Manche Autoren vermuten sie in fernen Ländern im Norden, andere spekulie-
1Das Wort „Lipid“ stammt von dem griechischen Wort für Fett: λ íπoς .
��
Einige der Atome sitzen sehr tief innerhalb der Membran und kommen nur
sehr selten an ihre Oberfläche. Andere befinden sich fast immer an der Ober-
fläche der Membran. Um das genauer zu messen, haben wir eine Methode ent-
wickelt, mit der man die Eintauchtiefe von Lipidatomen in die Membran mes-
sen kann. Die Methode kann man sich wie ein „atomares Lineal“ für sehr klei-
ne Abstände vorstellen – als würde man den Abstand zweier Millimeterstriche
auf einem Lineal noch 10 Millionen Mal unterteilen. Doch noch eindrucks-
voller ist, dass wir einen Tanz der Lipide, also ihre Bewegungen, beobachten
konnten. Einige Atome in den Lipiden bewegen sich sehr schnell, andere lang-
samer. Und so muss es sein. Die Membran ist keine steinharte Wand, sondern
weich und beweglich. Moleküle müssen sich bewegen, ansonsten können die
chemischen Reaktionen, die zu Leben führen, nicht vonstattengehen.
Gibt es den Greif tatsächlich?
WaaG ist also ein Zwitterprotein, das Wasser genauso wie Lipide mag, aber
es muss sich an der Lipidmembran befinden, um seine Arbeit, den Bau neu-
er Lipide, zu verrichten. WaaG bindet sich an negativ geladene Lipide in der
Membran mittels eines positiv geladenen Ankers. Und während dies geschieht,
tanzen die Lipide ihren Tanz und neue werden wieder und wieder hergestellt.
All diese Beobachtungen und Ergebnisse beruhen auf Untersuchungen, die
in einem Reagenzglas gemacht wurden. Aber geschieht all das auch in leben-
den Bakterien? Wir können mit großer Sicherheit sagen, dass einige der Beob-
achtungen auch in lebenden Zellen vor sich gehen. Lipide bewegen sich in al-
len Zellen. WaaG bindet elektrostatisch an Membranen. Aber bindet es immer
und unter allen Umständen? Bewegt es sich ebenfalls? Das wissen wir noch
nicht. Wir verstehen nun besser, wie WaaG und die Lipide gemeinsam funk-
tionieren, aber jede Antwort führt zu neuen Fragen. In der Wissenschaft geht
es darum Wissen zu erweitern und Grundlagen für ein besseres Verständnis der
Welt um uns herum zu legen. Mit dieser Arbeit wurde das Wissen über eine
Gruppe von Proteinen, die sich sehr merkwürdig verhält, erweitert. Außerdem
verstehen wir besser, wie sich Lipide bewegen und wie sie mit Proteinen inter-
agieren. Das bedeutet, dass wir einen weiteren weißen Fleck auf der Karte des
Wissens, von der auf den ersten Seiten dieser Arbeit die Rede war, beseitigen
können.
��
Popular science summary
As when a gryphon through the wildernessWith winged course, o’er hill or moory dale,Pursues the Arimaspian, who by stealthHad from his wakeful custody purloinedThe guarded gold.—John Milton, Paradise Lost
Proteins can be imagined as tiny, biological machines—the tiniest machines
that exist, just one millionth of a millimeter long. Together with fats and carbo-
hydrates they are the most important building blocks for life. These small parti-
cles are also called molecules. Proteins are immensely complicated molecules
that are made up of thousands of atoms in a complicated way that gives rise to
an ordered structure that catalyzes, i.e. performs, chemical reactions with ex-
tremely high precision. A protein can catalyze a chemical reaction in fractions
of a second that would otherwise take days or weeks to happen. Yet, what is
most astonishing is that proteins control all processes that happen in cells, so
that they may live. Without proteins, there can be no life.
However, more is needed for life than just a bunch of proteins. Another
important class of molecules are lipids, which are a specific group of fats.
Lipids are the most important constituents of membranes that can be thought
of as the walls of a house. Just like the walls protect people from the malev-
olence of the weather, membranes surround cells and protect them from the
malevolence of their environment. Membranes create an inside and an outside
environment. On the inside of a cell, vital proteins and other molecules are
found. On the outside of the cell there is nutrition, e.g. carbohydrates, but also
toxic molecules, from which the cell needs to protect itself.
Membranes consist of lipids. Lipids need to be manufactured by the cell
and that is where proteins enter the stage. Special proteins exist that manufac-
ture various types of lipids that are needed to build up a membrane. One of
these proteins was studied in this thesis. It is called WaaG and it is found in
the bacterium Escherichia coli. We investigated how WaaG functions inside
bacteria, which are organisms that consist of only one cell.
��
WaaG catalyzes one single but very important step in the construction of a
particular kind of lipid found in bacteria. This is one reason why we want to
understand how it works. Some bacteria cause serious diseases and such dis-
eases can be cured by killing the bacteria with antibiotics. One way of killing
them is by destroying proteins like WaaG that are involved in constructing
the bacterial membrane. Bacteria cannot survive with a damaged or destroyed
membrane. However, before we can impair the function of WaaG, we need to
understand how it works. But there is another reason why we are interested in
WaaG: Not only is it involved in the construction of the membrane, it also has
properties which makes it resemble a mythical creature: It is a griffin.
Who or what is the Griffin?
The ancient Greeks tell stories of a mythical creature that has the body and tail
of a lion, and the head and wings of an eagle: the griffin. Griffins are creatures
of the air and creatures of the land. Thanks to their peculiar combination of
eagle and lion characteristics, they are at home in both environments. WaaG is
a real-life, microscopic analogy to a griffin—a creature of two worlds. Thanks
to its peculiar combination of properties, this protein is at home in two envi-
ronments that couldn’t differ more: water, and lipids, which are very similar to
oil.1 Oil and water don’t mix. Everyone who tried to clean a greasy pan with
water knows that.
How can a molecule like both environments? This is hard to imagine and
this is why we had a closer look at WaaG. It turned out that it contains a re-
gion which can dock like an anchor to the membrane (see figure 6.3). Having
this anchor doesn’t cause WaaG any trouble in water, but once there is a lipid
membrane, it anchors WaaG to it.
Where does the Griffin live?
In ancient literature, several theories were put forward as to where griffins
might live. Some authors suspect that they reside in distant lands in the north;
others speculate they are found far to the east. Most of them agree, however,
that griffins are strongly attracted by gold. They are the guardians of unimag-
inable treasures.
WaaG is also attracted to its own kind of treasure: certain types of lipid
molecules. These lipids need to be negatively charged, so that the anchor,
which is positively charged, binds to the lipids and thus to the membrane. If
there are no negatively charged lipids, the anchor doesn’t work.
1The word ‘lipid’ originates from the Greek word for fat: λ íπoς .
��
Is the Griffin real?
So, WaaG is a hybrid protein that likes water and lipids alike but needs to be
at a lipid membrane to do its job, which is to build more lipids. It binds to
negatively charged lipids in the membrane using its positively charged anchor.
And, all the while, the lipids dance their dance and new ones are manufactured
over and over again.
However, all of our studies were done in a test tube. Is this all also happen-
ing in a real living cell? Some of the observations are most likely true for living
cells. Lipids move in all cells. WaaG binds electrostatically to membranes—
but does it always bind? Does it also move? This we don’t know yet. We have
gained a better understanding of how WaaG and the lipids work together, but
every answer gives rise to many new questions. Science is about extending
knowledge and laying foundations for a better understanding of the world that
surrounds us. We now have a better understanding about a very strangely be-
haved group of proteins. We also have a toolkit to look at the molecular dance
of lipids and proteins, and how they interact with each other. This means that
we can fill in another blank spot on the map of knowledge mentioned on the
first pages of this book.
��
Acknowledgements
It takes two to tango.—Pearl Bailey
Even though I wrote large parts of this book sitting by myself on a balcony
with a glimpse of Täby’s central park to my left and a forest of succulents to
the right, this thesis is not and cannot be the product of a solitary endeavor. On
the contrary, many people have contributed to it in one way or another. I’m
very grateful to all of them for sharing their knowledge and experience with
me, for discussions and advice, for criticism and support.
I’m grateful to Lena Mäler for supervising my work, for giving me freedom
to try out my ideas, and for coming up with new thoughts when I thought I
reached a dead end, for guidance to wrap up a story and for forcing me to
present it to others.
I like to thank Andreas Barth who undertook the task of scrutinizing my work
during all these years.
Special thanks go out to present, former, and adopted group members, in alpha-
betical order: Ane Metola, Axel Abelein, Biao Fu, Christian Brown, JoanPatrick, Johannes Björnerås, Montse Tersa, Pontus Pettersson, ScarlettSzpryngiel, Sofia Unnerståle, and Weihua Ye. I highly appreciate your ad-
vice that helps me in solving simple and complicated problems, I thank you for
climbing real and imaginary mountains with me and for laying science aside,
too.
I’m thankful for collaborations with Candan Ariöz, Claudio Muheim, andSusan Black and her group in Aberdeen who let me in on the secrets of protein
and lipid purification.
��
Creating a pleasant atmosphere during work, lunch, and fika, when traveling
to courses and conferences, and helping out with the small things in life is
extremely important. Thank you for doing this, Anna Wahlström, AstridGräslund, Cecilia Wallin, Cesare Baronio, Chenge Li, Christoph Loderer,Eloy Vallina, Erik Sjölund, Fan Yang, Fatemeh Madani, Göran Eriks-son, Huabing Wang, Ida Lindström, Inna Grinberg, Jakob Dogan, JensDanielsson, Johan Nordholm, Jüri Jarvet, Maurizio Baldassarre, NadjaEremina, Nicklas Österlund, Sarah Leeb, Sebastian Wärmländer, SeongilChoi, Syed Haq, Therese Grönlund, and Xin Mu.
Without technical and administrative support, nothing would work. Thank you
to: Britt-Marie Olsson, Haidi Astlind, Matthew Bennett, and TorbjörnAstlind.
Zum Schluss möchte ich meiner Familie danken. Uta und Till, Gefion undUrs, Nico und Johannes, Christian, Albrecht und Meng: Ihr habt mich bei
meinen Entscheidungen unterstützt und beraten, auch wenn das oft zur Folge
hatte, dass ich in Schweden war und nicht an unseren Familientreffen teil-
nehmen konnte. Ihr habt mir den Weg gewiesen und euch immer wieder auf
den Weg gemacht. Danke!
Having written this, I get the impression that Pearl Bailey was not quite right:
Sometimes, it takes more than two to tango.
��
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