solubilization and crystallization of membrane proteins · development of lipopeptide detergents...
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DEVELOPMENT OF LIPOPEPTIDE DETERGENTS FOR THE
SOLUBILIZATION AND CRYSTALLIZATION OF MEMBRANE PROTEINS
Clare-Louise McGregor
A thesis submitted in conformity with the requirements for the degree of Master of
Science, Graduate Department of Medical Biophysics, University of Toronto
O Copyright by Clare-Louise McGregor (2000)
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DEVELOPMENT OF LIPOPEPTIDE DETERGENTS FOR THE
SOLUBILIZATION AND CRYSTALLIZATION OF MEMBRANE PROTEINS
Master of Science, 2000
Clare-Louise McGregor, Department of Medical Biophysics, University of Toronto
Integral membrane proteins (IMPs) are involved in numerous cellular fûnctions.
Knowledge of their 3D structure is crucial to understanding their mechanism of action.
X-ray crystallography is the most powemil technique used to solve the 3D structure of
IMPs to atomic resolution. However, IMPs represent Iess than 1% of the protein data
bank structures. The bottleneck in obtaining these structures is the inabiiity to generate
well-ordered crystals. IMPs have large hydrophobic domains that are solubilized with
detergents which do not favor crystal formation.
This thesis presents a new class of detergents, Lipopeptide detergents (LPDs), designed to
improve the crystallization properties of IMPs. Their design, synthesis and purification
are preseoted. Their secondary structure, pH stability, micelle size and ability to
solubilize lipid bilayers are characterized. Finally, this thesis presents evidence
demonstrating that LPDs are superior to a traditional detergent, octylglucoside, in
maint;iining the solubility and stability of a mode1 IMP, bactenorhodopsin.
TABLE OF CONTENTS:
- . ....................................................................................... Abstract LI
List of Tables ................................................................................. vi
................................................................................ List of Figures vi ... ................................................................................. Abbreviations v u
CHAPTER 1 : INTRODUCTION
......................... 1.1 Membranes and Membrane Proteins .. ............. 1
1.2 The Problems in Membrane Protein Crystailization
.......................................................... A) Hydrophobic Domains 5
........................................................................ B) Detergents 6
..................... C) Micelles - A Consequence of the Hydrophobie Effect 8
....................................... D) Membrane Protein Crystai Formation 12
................................................. E) Protein-Detergent Complexes 14
1.3 Success in Achieving High-Resolution Structural Idormation of Membrane Proteins
........................... 1 -4 Novel Approaches to Crystallizing Membrane Proteins 20
............... A) Antibody-mediated Crystallization of Membrane Proteins 20
....................................................... B) Internai Fusion Proteins 21
...................................................................... D) Peptitergents 22
............................................................. E) Lipidic Cubic Phases 23
.................................................................................. 1.5 Objective 24
CHAPTER 2: EXPERIMENTAL DESIGN
....................................................... 2.1 Lipopeptide Detergent Design 25
............................................................. 2.2 Peptide Synthesis Strategy -28
............................................................... 2.3 Physical Characterization 31
........................................................... 2.4 Solubilization of Liposomes 36
.................................................. 2.5 Bacteriorhodopsin Stability Triais 36
iii
2.6 Crystallization Trials ................................................................... 39
CHAPTER 3: MATERIALS AND METHODS
..................................... ............................ 3.1 Peptide Synthesis .. 40
3 -2 LPD Purification ....................................................................... 42
3.3 Electrospray Ionization Mass Spectrometry ........................................ 42
3.4 LPD Concentration Determination .................................................. 43
3 -5 Circular Dichroism ...................................................................... -43
3 -6 Micelle Size Determination
A) Gel Filtration Analysis ...................................... .. . . . . . . . . . 44
B) Sedimentation Equilibrium Anaiysis ........................................ 44
3 -7 Liposome Solubilization
A) Liposome Preparation ......................... .. .. .... .............. 46
B) Phosphate Determination .......................... .... ..................... 46
........................................................... C) Light Scattering 4 7
3.8 Purple Membrane Isolation and Bacteriorhodopsin Purification
A) Purple Membrane Isolation .................................................... 47
B) Bactenorhodopsin Purification .............................................. 49
3.9 Bacteriorhodopsin Stabilization
A) Detergent Exchange ............................................................ -49
................................................................. B) Detergent Assay 50
C) Solubility and Stability Analysis of Bactenorhodopsin .................... 51
D) Sedirnentation Equilibrium Analysis of the LPD Solubilized BR ......... 51
3.1 O Crystailization Trials .................................................................. -52
CHAPTER 4: RESULTS
4.1 LPD Synthesis and Purification ........................................................ 53
4.2 Characterization of the LPD Series
.............................................. A) Hydrophobicity and Solubility 53
B) Secondary Structure .......................................................... 59
................................................................... C) Micelle Size 63
D) Liposome Solubilization .................................................... 70
4.3 Bactenorhodopsin Purifkation and Stability Trials ......................... .... 74
................................................................. 4.4 Cryçtaliization Trials 86
CHAPTER 5: DISCUSSION AND FUTURE WORK
5.1 Characterization of LPDs ................................ ,.,. ....... 90
5.2 LPD Solubilization of Phosphoiipid Bilayers ..................... .. ............. ... 95
5.3 BR Stability and Crystaljization in Association with LPDs ...................... 98
................................................. 5.4 Future Work ..................... .., 101
............................................................... REFERENCES ........... ... IO4
List of Tables:
.... Table 1.1 : Crystallized membrane proteins with high-resolution a-helical structures 18
.... Table 1.2. Crystaliized membrane proteins with high-resolution p-barre1 structures 19
Table 3.1 : Rotor speeds used for sedimentation equiiibrium analysis ..................... 45
Table 4.1 : Cornparison of LPD calculated and observed LPD molecular weights ........ 56
Table 4.2: Comparison of gel filtration and sedimentation equilibrium ultraceneifugation
determination of micelle size .................................................................... 66
List of Figures:
Figure 1.1 : Fluid mosaic mode1 of a biological membrane ................................... 2
Figure 1.2: Examples of commonly used detergents in membrane protein solubilization
and purification ..................................................................................... 7
Figure 1.3 : Micelle formation .................................................................... 9
Figure 1.4: Comparison of phases. molecular shapes and packing parameters of a
............... traditional detergent and a phospholipid ..................................... .. 11
..................... Figure 1.5. Membrane protein c r y d types ......................... .. 13
Figure 1.6. Phase diagram of an example detergent CsEs .................................... 16
Figure 2.1 : LPD design ........................................................................... 26
Figure 2.2. LPD synthesis flow chart ..................................................... 30
Figure 2.3. LPD modelled using RasMol Version 2.6. ....................................... 33
Figure 2.4. Cornparison of molecular shapes of monomers and micelles .................. 34
Figure 2.5: A) Proposed self-assembly of LPD rnonorners B) Comparison o f the
stabilization of an IMP by LPDs and traditional detergents ................................. 35
Figure 2.6. Ribbon diagram of bacteriorhodopsin ............................................ 37
Figure 4.1 : Cornparison of LPD-12 elution profles fiom C4 RP-HPLC .................. 54
Figure 4.2. ESI-MS spectnim for LPD C-12 following Cq RP-HPLC .................... 55
Figure 4.3. Cd RP-HPLC elution of the LPD series .......................................... 58
Figure 4.4. CD wavelength scan of the LPD series .......................................... 60
Figure 4.5. pH dependence on secondary structure .......................................... 61
Figure 4.6. Concentration dependence of LPDs on secondary structure .................. 62
Figure 4.7. Elution profile of LPDs on Superdex 75 HR.30 SEC coIumn ................ 64
Figure 4.8: Estimate of LPD micelle size using calibration molecular weight standards
................................... ,. .............................................................. 65
Figure 4.9. Sedimentation equilibrium ultracentrifugation of LPD- 1 2 ................ -68
Figure 4.10: Calculation of the apparent molecular weights of LPD-12 and LPD-20 with
respect to changing partial specific volume ...................... ,. .................. 69
Figure 4.1 1 : Cornparison of concentration titration of 0.1 rnM PC liposomes ....... 72
Figure 4.12. Summary of 0.1 mM PC Liposome solubilization ........................... 73
Figure 4.13 : PM isolation and BR purification ........................................... 75
Figure 4.14. OG remaining in retentate following the exchange wash steps .. ....... 76
Figure 4.15: Coomassie stained 10-20% SDS polyacryIamide tricine gel determinhg the
minimm LPD -12 concentration required to fùlly solubilize BR ....................... 77
Figure 4.16: Time course monitoring the stabiIity of LPD-12 solubilized BR to determine
........ the minimum LPD- 12 concentration required to maintain the solubility of BR 80
... Figure 4.17. Time course monitoring the stability of 0.5 mM LPD solubilized BR - 8 1
Figure 4.18: Spectra monitoring the stability of BR solubiiized over a 30 day period .. 82
Figure 4.19: Comparison of LPD- 18 spectra monitoring the stability of BR solubilized
.......................................................................... over a 32 day period 83
Figure 4.20 : Sedimentation equilibrium ultracentrifûgation of LPD- 1 2 solubilized BR
......................................................................... ......................... .. 84
Figure 4.21 : Prelimùiary modelling of an LPD solubilized BR trimer ................. 85
Figure 4.22. LPD-20 solubilized BR crystailization trials ................................ 88
Figure 4.23 : Phase sepration examples of LPD-20 solubilized BR .................... 89
vii
ABBREVIATIONS:
Boc - t-butoxycarbonyl
BR - bacteriorhodopsin
CD - circular dichroism
CFTR - cystic fibrosis trammembrane conductance regulator
cmc - critical micelle concentration
DCM - dichloromethane
DDM - dodecyimaltoside
DIPEA - diisopropylethylamine
DLS - dynamic light scattering
DMF - dimethylformamide
ES1 - electrospray ionkation
Fmoc - 9-fluorenylmethoxycarbonyl
HATU - O-(7-azabenzotriaz01-I-y1)- 1 1 ¶3¶3-te- hexafluorophosphate
HEPES - N-2-hydroxyethylpiperazine-N'-2-ethanesul acid
HPLC - high-performance liquid chromatography
IMP - integral membrane protein
LDAO - laury ldimethy lamine oxide
LPD - lipopeptide detergent
MBHA - 4-methy lbenzhy dry lamioe
MDR - multidmg resistant
MO - 1 -monooleoyl-rac-glycerol
MP - 1 -monopalmitoley 1-rac-glycerol
N - aggregation number
NMR - nuclear magnetic resonance
OD - optical density
OG - octylglucoside
PC - phosphatidylcholine
P-gp - P-glycoprotein
PLB - phospholipid bilayer
viii
PM - purple membrane
RP-HPLC - reverse phase HPLC
SDS - sodium dodecyl sulfate
SEC - size exclusion chromatography
SGM - standard growth media
S N - signal to noise ratio
SOS - sum of squares
TFA - trifluoroacetic acid
TFMSA - trifluorometbane sulfonic acid
UV - ultraviolet
CHAPTER 1: INTRODUCTION
1.1 Membranes and Membrane Proteins
Biological membranes provide both structural and functional roles within a ce11 or
organelle. The traditionai ikid mosaic mode1 of a biological membrane depicts proteins
embedded within a phospholipid bilayer. This bilayer is composed of phospholipids
oriented such that the hydrophilic heads face the aqueous environment and the
hydrophobic tails form the large interior of the cellular membrane (Figure 1.1) (Lodish et
al., 1995). The bilayer serves as a permeability barrier within prokaryotic and eukaryotic
cells and also compartmentalizes the organelles within eukaryotic cells. The proteins
embedded within this bilayer, however, MfiU functional roles for the ce11 such as solute
transport, signal transduction and cell-celi recognition. There are two types of proteins
associated with biological membranes: integral and peripheral membrane proteins.
Integral membrane proteins (IMPs) are those proteins that embed within the phospholipid
bilayer. Monotopic IMPs embed only on one side of the bilayer, whereas bitopic and
polytopic IMPs extend the entire width of the bilayer, crossing the bilayer once or several
times, respectively (Tsukihara et al., 2000). Peripheral membrane proteins, on the other
hand, are only extrinsically associated with the membrane through protein-protein
interactions or protein-lipid interactions (Lodish et al., 1995). This thesis focuses on
integral membrane proteins. More specifically, this thesis describes the development of a
novel detergent designed to facilitate the structural analysis of integral membrane
proteins.
- PLB hydrophobic core
v peripheral proteins integral proteins
Figure 1.1: Fluid mosaic mode1 of a biological membrane (Adapted nom Lodish et al.,
1995). Integral membrane proteins are embedded within the hydrophobic core of the
phospholipid bilayer (PLB) whereas peripherai membrane proteins associate via protein
andor Lipid interactions.
IMPs are predicted to account for 20-30% of the products encoded in eubacterial, archaen
and eukaryotic genomes (WaiIin et al., 1998). If an IMP is overexpressed, deleted or
mutated, diseases could mise as a result of the disniption of normal cellular functioning.
A number of extensively investigated IMPs are relevant to human health. For exampie, P-
glycoprotein (P-gp l) is a membrane protein which is overexpressed in tumor drug
resistant phenotypes (Roepe, 2000). Although a "drug pump" model was proposed in
1973 to explain its mode of action in conferring multtidnig resistance, little progress has
been made in confirming or disputing this model due to a lack of structural data. Cystic
fibrosis transmembrane conductance regulator (CFTR) and human erythrocyte anion
exchanger 1 (band 3) are two other examples of membrane proteins in which mutations
result in diseased phenotypes. The deletion of phenylalanine 508 in CFTR, for instance,
results in a processing defect which fails to target this protein to epitheiial membranes.
Symptoms, therefore, arise as a result of the disruption of the flow of salt and water
across these epithelial membranes and patients present with cystic fibrosis (Hwang et al.,
1999). While this disease affects the pancreas, intestines, sweat ducts and reproductive
tracts, its most severe disfùnction arises in the lung (McCarty, 2000). Band 3, on the
other hand, is an abundant chlonde / bicarbonate exchanger found in erythrocytes. This
protein is invoived in membrane stability, erythropoiesis and acid-base regdation of the
blood (Peters et al., 1996 and Wang et al., 1994). Its mutated form results in the
reduction in the integrity of the erythrocyte membrane, alters the shape of the ce11 and
results in the manifestation of hereditary spherocytosis disease. These proteins, P-gp 1,
CFTR and band 3 are just three of a wider range of medically relevant proteins whose
high resolution structural information would aid in the progress of treatment of disease
using a rational method for drug design. However, despite the relevance of IMPs to
human health, most progress with structure based drug design has been made with
soluble protein targets. For instance, structure-based design is king utilized to generate
inhibitors of a variety of proteins involved in diseases such as diabetes and amyloid
disorders (Iversen, et al., 2000 and Klabunde et al., 2000).
Currently, there are three approaches to solve the 3-dimensionai structure of a protein.
They indude electron microscopy, nuclear magnetic resonance (NMR) and X-ray
crystallography. Al1 of these techniques are limited in some way. Although electron
microscopy has shown promise in elucidating the structure of 2D membrane protein
crystals, it is a technique limited in terms of its inability to achieve atomic resolution
(Hasler et al., 1998). Biological samples undergo extreme radiation damage under an
electron beam. Furtherrnore, the inherent instrument limitations of the electron
microscope itself limit the resolution attainable by this method (Glaeser, 1999). NMR,
on the other hand, is Iimited by protein size restraints to obtain a resolvable spectrum; 40
kDa is generaily accepted as the molecular weight limit. Since most IMPs exist in
complexes exceeding this size restraint and must be solubilized in the presence of
Iiposomes or detergents, NMR is not the ideal approach for solving 3-dimensional
structures of intact IMPs. Recently, however, progress has been made for the NMR
determination of membrane proteins ushg magic angle spinning NMR spectroscopy
(Smith et al., 1996). Finaliy, X-ray crystallography has proven to be the most powerful
technique of al1 the methods of structure determination. Since 1984, it has achieved high
resolution structures of approximately 30 different IMPS. However, this is limited
success as less than 1% of the protein structural data bank are attributed to membrane
proteins. The underlyïng problem with crystauizing membrane proteins is the fact that the
large hydrophobic domains within IMPs must be solubilized with detergents outside of
the lipid bilayer environment. These detergents reduce the success of crystailization due
to the volume they occupy and their inherent flexibility. As a result, a wealth of
knowledge is omitted nom the protein data bank and great efforts are placed into creating
new methods to obtain high-resolution membrane protein structures via X-ray
cry stallography .
1.2 The Problems in Membrane Protein Crystallization
A) Hydrophobic Domains
The bottleneck in solving membrane protein structures by X-ray crystallography fies in
obtaining well-ordered 3-dimensional crystals. The fundamental problem is the fact that
membrane proteins have large hydrophobic domains which cross the width of the
phospholipid bilayer (Figure 1.1). In the absence of a membrane, these hydrophobic
domains interact non-specifically, causing aggregation and consequently, the proteins
precipitate out of solution. These domains, therefore, must be stabilized with detergents
to achieve a soluble system suitable for crystallization trials. In this case, the Iipid
surrounding the hydrophobic domain of the IMP is replaced with a "belt" of detergent
molecules (Ostermeier et al., 1995 and 1997).
B) Detergents
A detergent is an amphiphilic rnolecule used to solubilize hydrophobic compounds in
aqueous solutions (Neugebauer, 1990). Detergents are broadly classified into three
categories depending on the charge on the hydrophilic head group: ionic, zwitterionic and
non-ionic. Examples of commonly used detergents in protein purification and structural
analysis are found in Figure 1.2. Each of these three detergent categories is represented.
Ionic detergents have a charged head group that is usually attached to either an alkyl
chah or a steroid structure. Zwittenonic detergents have a neutrd head group that is also
usually attached to alkyl chahs or steroid structures. Generaily, ionic and zwittenonic
detergents are used to break protein-protein interactions, ofien changing the protein's
conformation. Consequently, many of these detergents are considered denaturants
(Michel, 1983). Nonionic detergents, on the other hand, contain a non-charged head
group and are generally used for breaking iipid-lipid and lipid-protein interactions. These
detergents are capable of maintainhg the native conformation of protein and as a result,
are considered mild detergents. Nonionic detergents, therefore, are the preferred
detergents for solubilizing membrane proteins (Michel, 1983). The size of the
hydrophobic tail also plays a role in the "hanhness" of a detergent. Long tails tend to
stabilize membrane proteins better than shorter tails. Generally, miid detergents have
large, neutral head groups with long alkyl tails, whereas harsh detergents contain small,
charged head groups with short alkyl tails (Michel, 1991).
Figure 1.2: Examples o f commonly used detergents in membrane protein solubilization
and purification. A) sodium dodecyl sulfate (SDS) B) lauryldimethylamine oxide
(LDAO) C) (upper) octylglucoside (OG) and (lower) dodecylmaltoside @DM).
C) Micelles - A Consequence of the Hydrophobie Effect
When an amphiphilic compound is placed in an aqueous solution, the intermolecular
hydrogen bonding of water molecules surrounding this compound is disrupted because
the nonpolar portions of the compound are unable to partake in hydrogen bonding with
water. As a result, the water molecules surrounding this compound rearrange to form a
more ordered cagelike conformation that results in an overall decrease in entropy of the
system (Neugebauer, 1990). Consequently, in order to M z e the decrease in entropy
of the system, water molecules force these compounds to aggregate to occupy minimum
space. The critical micelle concentration (cmc) is the concentration at which monomenc
arnphiphilic molecules cluster to form micelles. Micelles are assemblies in which the
polar moieties of the amphiphitic molecules are exposed to the solvent and the nonpolar
moieties of the compounds form a hydrophobie core shielded fiom solvent (Figure 1.3A)
(Gennis, 1989). In an aqueous solution in which the detergent concentration is above the
cmc, monomers and micelles exist in equilibrium (Figure 1.3B) (Hjelmeland, 1 986).
Monomer
concentration
(monomers
and micelles)
cmc
Micelle
-
monomer
-
f--- micelle
L
total detergent concentration
cmc
Figure 1.3: Micelle formation A) Micelles fonn from monomers when the crnc is reached
(Gemis, 1989) B) Monomer and micelle concentration as a function of total detergent
concentration (Adapted from Hjelmeland, 1986). Cmc is the critical micelle
concentration.
The size and shape of the micelle depends largely on the structure of the monomer itself.
Each monomer has a packing parameter, P, = VJa,,l, ,where V, represents the volume of
the tail, a, represents the optimal surface area of the head group and 1, represents the
length of the tail. When P, is less than 1/3, a sphencal micelle results, and when P, is
greater than 1/2, a bilayer forms (Neugebauer, 1990). Figure 1.4 compares the molecular
shape of a traditionai detergent to that of a phospholipid (Gennis, 1989). Traditional
detergent monomers with a single acyl chah are cone-shaped and form spherical
micelles. Phospholipids, on the other hand, have two acyl chains attached and as a result,
have a much larger nonpolar volume to surface area ratio than traditional detergents.
Consequently, the cyhdrically shaped monomeric phospholipids form a bilayer.
Micelle size and aggregation number 0, the number of monomers in each micelle, is
also affected by the charge of the head group and solvent conditions such as pH, ionic
strength and temperature (Hjelmeland, 1986). Al1 of these factors combined play critical
roles when solubüizing IMPs for the purposes of crystallization. It is desirable to choose
a detergent that will form a relatively small detergent belt around the hydrophobic
domain of the IMP but which also serves as an appropriate mimic of the lipid bilayer and
maintains the stability of the IMP.
Traditional
detergent
Phospholipid
Cone
Phase
nnnn
bilayer
11
Criticai Packing
Parameter
1/2<Pp< 1
(E3ilayer Sheet)
Figure 1.4: Cornparison of phases, molecular shapes and packing parameters of a
traditional detergent and a phospholipid (Adapted fiom Gennis, 1989). The detergent and
phospholipid hydrophilic heads are grey and blue, respectively. The hydrophobie tails
are black.
D) Membrane Protein Crystal Formation
Membrane proteins can form crystais in one of two ways: within a lipid bilayer (Type 1)
or as a detergent solubilized cornplex (Type II) (Figure 1.5). Type I crystals are formed
by havhg two-dimensional crystals ordered in a third dimension. Both hydrophobic and
polar interactions between the protein and lipid stabilize the crystal lattice in two
dimensions but the protein's polar interactions create the lattice contacts in the third
dimension (Michel, 1983). Until recentiy, electron microscopy has proven to be the tool
to analyze these structures to obtain low-resolution structures.
Type II membrane protein crystals, on the other hand, are obtained in the presence of
detergents. In this case, the crystal contacts are due primarily to the polar interactions
between the extramembranous hydrophilic domains. in some cases, polar interactions
between the hydrophilic moieties of the detergent molecules themselves also contribute
to the stability of the crystal laîtice (Ostermeier et al., 1997).
Type II
Figure 1.5: Membrane protein crystal types. The hydrophilic and hydrophobic surfaces of
the IMPs are red and green, respectively. The phospholipid and detergent hydrophilic
moieties are depicted as blue and grey, respectively whereas their hydrophobic moieties
are black. Type 1 membrane protein crystals are formed in the presence of lipid and form
2-dimensional crystais which are ordered in the third dimension. Hydrophilic and
hydrophobic interactions forrn the ordered three dimensions. Type II membrane protein
crystals are formed in the presence of detergent and are formed p r i m d y by the polar
interactions of the hydrophilic protein domains in 3-dimensions. (Adapted nom Michel,
1983).
E) Protein-Detergent Complexes
As mentioned, many factors are taken into consideration when attempting to solubilize an
IMP in detergent for the purposes of crystallization. The detergent rnust be mild so as to
maintain the native conformation and active state of the protein; protein stability is a
prerequisite for crystallization since denatured proteins are poor candidates for forming
crystals. In addition, the detergent rnust be small enough to enable the extramembranous
polar domains of the protein to form the crystal contacts necessary for Type II crystals
(Figure 1.5). Unfortunately, tradi tional detergents have achieved on1 y limited success in
generating well-ordered 3-D crystals of IMPs suitable for high-resoiution structural
analysis for three reasons. First, since type II crystal lattices are formed by rigid polar
interactions between the proteins' extramembranous hydrophilic domains. long tailed
detergents push the individual protein molecules further apart reducing the opportunity
for these polar interactions to be established. Second, the hydrophilic head of the
detergent interferes with the protein's hydrophilic domain thereby reducing the polar
surface area available to establish these crysta1 contacts. Third, due to the size and
flexibility of the detergent, these crystal lattice contacts are ofien not repeated in a regular
three dimensional array which is necessary to achieve a well-ordered crystaI suitable for
structural analysis.
Finding the "right" detergent to obtain properly folded proteins for well-ordered crystals
is a difficult task. As mentioned, the charge and size of the head group as well as the
length and size of the hydrophobie tail al1 play critical roles. In fact, minor changes in
any detergent property can have large implications when crystallizing an IMP. For
instance, cytochrome c oxidase fiom bovine beef heart mitochondria would only form
well-ordered crystals using decy lmaltoside but no other length of maltoside (Ostermeier
et al., 1997).
The concentration of the detergent also plays a role in the stability of the system. Excess
detergent can induce protein denaturation by the dissociation of subunits or solubilization
of hydrophobic cofactors (Tribet et al., 1996). Polar interactions between excess
detergent micelles are the underlying force behind phase separation. Phase separation
occurs when the crystallization solution separates into detergent-rich and detergent-
depleted regions (Figure 1.6). These regions play a role in crystallization as some
proteins will ody crystallize close to the detergent-rich phase (Ostermeier et d., 1997).
In short, it is very difncult to predict what detergent conditions are suitable to generate a
soluble, stable protein that can produce well-ordered crystals. To summarïze, a mild
detergent that maintains the active state of the protein is desired. It must be smail enough
to allow polar interactions to form between the extramernbranous domains of the protein
and ngid enough to mullmize the dynamics of the system to allow these contacts to be
maintained.
Figure 1.6: Phase diagram of an example detergent, CsEs. M represents detergent
monomers in solution, Li represents micelles and Li' and L 1" represent two other micelle
phases. The figures a, p, y and 6 demonstrate that at concentrations below the cloud point
the concentrated lamellar phase has a lower volume than the dilute lamellar phase.
(Adapted fkom Zulauf, 199 1).
1.3 Success in Achieving High-Resolution Structural Information of Membrane
Proteins
Despite the difnculty in obtaining the optimal protein-detergent complexes suitable for
crystallization, several membrane protein families have been solved to high resolution.
Typically, these membrane proteins have crystallized as type LI crystals. Tables 1.1 and
1.2 present the membrane proteins that were solved with the detergent(s) andor lipids
that were used.
These tables are divided into a-helical and f3-barre1 structures because both structures can
satisfi the thermodynamics of the phospholipid bilayer. In both cases, dong the
transmembrane domain of the IMP, the charged and polar amino acids form the interior
of the protein structure whereas the hydrophobic amino acids are oriented to the exterior
of the protein to face the hydrophobic acyl chahs of the phospholipid bilayer. Helical
membrane proteins are found in prokaryotic and eukaryotic inner membrane and plasma
membranes, respectively. fi-sheet membrane proteins, on the other hand, are generally
found in the outer membranes of bacteria but have also been found in mitochondria (Liu
et al., 1999).
Table 1.1 : Crystallized membrane proteins with hi&-resolution a-helicai structures
(adapted fkom http://194.95.28.4/micheU~ublic/me~nprotsct. h l ) References for these
structures c m be found at this web address.
Bacteriorhodopsin
Calcium ATPase
Cyclo-oxygenase (COX- 1 = prostaglandin
H2 synthase 1, COX-2)
Cytochrome bc 1 complex
(respiratory complex III)
Cytochrome c oxidase
(respiratory complex IV)
Fumarate reductase (succinate dehydrogenase/
respiratory complex II)
Halorhodopsin
Light harvesting complex
Mec hanosensitive channel (MscL)
Photosynthetic reaction center
Potassium channel
Squalene cyciase
OG, Lipid
Lipidic cubic phase (Type 1)
OG, OPOE
I DMG or DHPC, OG, DDM or
DDM+rnCG
DDM, UDM, DM, NG
DDNOE,
DDM+DM
Lipidic cubic phase (Type I)
Triton X100, OG, DMUDAO
DDM
DMDAO, OG
DMDDA
OTOE
Table 1.2: Crystallized membrane proteins with high-resolution barre1 structures
(adapted fiom http://194.95.28.4/micheVpublic/mem rotstnict. html) References for eac h
of these structures c m be found at this web address.
PROTEIN DETERGENT 1
1 8-stranded p o ~ s (maltoporin, ScrY)
8-stranded membrane anchor (ompA, ompX)
12-stranded membrane anchor (OMPLA)
1 6-stranded porins (ompF, PhoE, OmpK3 6)
OTOE
OG
OTOE, OHEStDMDAO, OG
Legend for Tables 1.1 and 1.2: DDG = n-dodecyl-8-D-glucopyranoside, DDNOE =
dodecylnonaoxyethylene, DDM = dodecyl-fi!-D-maltoside, DDNOE = dodecyl
nonaoxyethylene, DHPC = diheptanoyl phosphatidylcholine, DM = decyl-PD-maltoside,
DMDDA = N,N-dimethyldodecylamine, DMDAO = N,N-dimethy1dodecylamine-N-
oxide, DMG = decanoyl-N-methyl-glucamide, DMHAO = N,N-dimethylhexylamine-N-
oxide, DMUDAO = N,N-dimethylundecylamine-N-oxide, HG = n-heptyl-p-D-
glucopyranoside, HxG = n- hexyl-p-D-glucopyranoside, MHCG = methyl-6-O-(N-
heptylcarbamoy1)-a-D-glucopyranoside, NG = nonyl-p-D-glucoside, OG = Octyl-p-D-
glucopyranoside, OHES = n-octyl-2-hydroxyethylsulfoxide, OPE =
octylpolyoxyethylene, OPOE = octyIpentaoxyethylene, OTOE = octytetraoxyethylene,
UDM = undecy l-f3-D-maltoside
Outer membrane transporters
22 stranded receptor (FhuA, FepA)
a-hemolysin
Outer membrane protein (ToLC)
DMDAO, OHES, DMDAO
OG
DDG+HGtHxG+OG
1.4 Novel Appmches to Crystaîüzing Membrane Proteins
A vast amount of effort has k e n placed into obtaining membrane protein crystals using
traditional techniques. Tables 1.1 and 1.2 s m a r i z e the less than 40 structures obtained
to date. These structures represent the fimit foliowing many years of failed attempts and
do not reflect the variety of other IMPs that have been pursued. Therefore, as a result of
the limited success ushg traditional means, a variety of novel approaches to crystallizing
membrane proteins have been proposed.
A) Antibody-mediated Crystallization of Membrane Proteins
Since the cntical crystai lattice contacts are made between the polar extramembranous
domains of the proteins themselves, Ostermeier et al. proposed that if the polar domain of
the membrane protein was enlarged, then these critical cystal lattice contacts would be
more easily achieved (Ostermeier et al., 1995). This investigation directed an F,
fragment of an antibody against the extramembranous portion of cytochrome c oxidase.
The F, fragment is a soluble protein that binds noncovalently with high affinity and
specificity to the exposed extramembranous hydrophilic domain of the membrane
protein. F, fiagrnents are good candidates for the crystaliization of membrane proteins
because they are not flexible and can crystallize easily. Ostermeier generated crystals of
the cytochrome c oxidase- F v complex in the presence of dodecylmaltoside that difiacted
to a resolution of 2.8 A. As predicted, the cntical crystal lattice contacts were through
the polar interactions of the F v fragments. A major limitation of this technique for
producing IMP crystals is generating the specific monoclonal antibodies necessary for
each IMP.
B) Interna1 Fusion Proteins
The internal fusion technique is based on the same premise as the antibody-mediated
crystallization of membrane proteins where increasing the overail polar surface area of a
membrane protein wouId facilitate its crystallization. Traditional fusion techniques
generate an N- or C-terminal fusion of two proteins, producing two domains connected
by a flexible linker. This traditional type of fusion does not favour the production of
crystals due to the flexibility between the two domains. The internal fusion technique,
however, involves inserting a ''carrier protein" into an interior loop(s) of a membrane
protein. The inserts or "carrier proteins" are soluble proteins carefully selected based on
the following cnteria: (1) previously crystallized (2) monomenc (3) N- and C- termini
within 5-12 A and at the surface of the molecule (4) soluble and stable under a wide
range of pH and ionic strengths and (5) greater than 30 kDa in size (Rivé et al., 1994 and
Pnvé et al., 1996). A great deal of effort has been invested in designing these fusions for
the 12 transmembrane a-helical protein, lactose permease. Although success has k e n
achieved in terms of generating fusion proteins that are highly expressed, active and
stable, lirnited success has been achieved in obtaining crystals. However, an internal
fusion between two soluble proteins, maltose binding protein and cytochrome b,, has
been crystallized which shows promise that this technique could work for membrane
proteins (Ahn and Pnvé, unpublished).
C) Amphipols \
Amphipols are a new class of surfactants designed to maintain membrane proteins in an
aqueous solution fkee of detergent (Tribet et al., 1996). The amphipol is an arnphiphilic
polymer composed of a hydrophilic backboae grafied with hydrophobic chains. These
amphipols were able to replace the detergent and maintain the stability of several
membrane proteins - bacterïorhodopsin (BR), bacterial photosynthetic reaction center,
cytochrome bsf and matrix porin. Rate zona1 centrifugation showed that these protein-
amphipol complexes were monodisperse, containhg no large aggregates. Despite the fact
that amphipols replace detergents in aqueous solutions, they can also interact with the
polar domains of the membrane proteins. This, coupled with the fact that amphipols are
structurally flexible, makes them poor candidates for crystallization purposes.
D) Peptitergents
A peptitergent is a 24 residue a-helical amphipathic peptide designed to replace
detergents in solubilizing membrane proteins. This approach was based on the premise
that amphipathic helices (helical coiled coils and four-hehc bundles) have been shown to
associate in such a way as to partition hydrophobic groups away fiom solvent (Presnell et
al., 1989). The peptitergent helix was designed to contain a 'Ylat" hydrophobic face that
would interact with the hydrophobic domains of membrane proteins (Schafheister et al.,
1993). It was designed to be superior to traditional detergents by packing around the
membrane proteins in a more rigid, well-ordered manner. More specifically,
Schafineister postulated that a paraliel a-helical arrangement of peptitergents could align
dong the length of the membrane protein.
Peptitergents maintained the solubility of BR and rhodopsin over two days, achieving
85% and 60% solubility, respectively (Schafineister et al., 1993). However, peptitergents
failed to maintain the solubility of the potin, PhoE. This suggests that these helical
peptides show promise in terms of solubilizing helical membrane proteins but not f3-
barre1 structured membrane proteins. A crystal structure of the peptitergent alone was
solved to 2.5 A resolution that revealed an antipardel four-helk bundle in which the
monorners interacted flat surface to flat surface (Schafmeister et al., 1993).
Unfortunately, no crystals of a peptitergent-membrane protein complex have been
reported to date and therefore, no structural information is available to evaluate the
promise of this peptide in crystallizing IMPs.
E) Lipidic Cubic Phases
Lipidic cubic phases are iipidwater mixtures that display cubic symmetry (Gouaux,
1998). These phases are comparable to biological membranes in terms of their
viscoelastic properties. A desùable feature of these phases is that they can incorporate
proteins, detergents and precipitants without perturbing itself or the membrane protein.
Connected aqueous channels are dispersed throughout the lipid matrix that allow proteins
to lateraily diffuse throughout the matrix to form the directional contacts necessary for
crystal formation (Landau et al., 1996).
Type I BR crystals were obtained that difiacted to nearly atomic resolution using this
lipidic cubic phase approach (Landau et al., 1996, Luecke et al., 1999). BR remained
stable within this matrix by partitionîng its hydrophobie domains within the lipidic phase
and its hydrophilic domains within the aqueous channels. BR crystallized in different
forms depending on the lipid used. It formed hexagonal and rhombic crystals in the
monoolein (C18:lc9) and monopalmitolein (C16rl&) matrices, respectively. In fact, this
technique has also shown considerable promise for crystallizing other membrane
proteins; halorhodopsin, a light-driven chloride pump has recently been solved to 1.8 A
resolution (Kolbe et al., 2000).
1.5 Objective
Despite the limited success of the novel approaches mentioned above, an approach has
yet to be discovered for crystallizing membrane proteins that will rapidly produce hi&-
resolution membrane protein structures. Therefore, this thesis presents the preliminary
work behind another novel approach used to solubilize and crystallize membrane proteins
more effectively and efficiently than traditional means. This thesis presents the
characterization of a new class of detergents, calied Iipopeptide detergents (LPDs) and
presents evidence demonstrating that these LPDs are superior in maintaining the
solubility and stability of the model membrane protein bacteriorhodopsin.
CHAPTER 2: EXPEïUMENTAL DESIGN
2.1 Lipopeptide Detergent Design
The ultimate goal behind designing lipopeptide detergents (LPDs) was to create a new
detergent that was a better candidate for crystallizing IMPs. It was designed to be a
better mimic of the phospholipid bilayer, occupy less space and be more rigid than
traditional detergents.
The basic scaffold of an LPD is a 25 residue a-helical amphipathic peptide. The peptide
was designed to be approximately 37 A in length when folded which is long enough to
span the width of a phospholipid bilayer (30-45 A). In order for this detergent to be a
better m e c of the phospholipid bilayer, fatty acyl chahs (12 to 20 methylene units in
length) were designed to be covalently coupled to both ends of the peptide (Figure 2.1).
Phospholipids within a biological membrane generally contain between 16 to 18
methylene units, whereas traditional detergents used to solubiiize IMP rarely exceed 12
allcyl units in Iength due to solubility limitations (Lodish et al., 1995). Furthemore, since
this peptide was designed to fold into an a-helix with the fatty acyl chahs aligning
closely dong the hydrophobic domain of the IMP, it was proposed to form a smaller,
more ngid, well-ordered complex with membrane proteins than traditional detergents.
This charactenstic, in particular, would facilitate crystallization of LMP-LPD complexes.
Figure 2.1: LPD design A) Amino acid sequence of the designed peptide. The
hydrophobic amino acids are green, the charged amino acids are red and the polar, non-
charged amino acids are blue. The O is ornithine and is the site of fatty acid coupling.
The black bars represent the potential Glu-Lys salt bridges at positions i, i+4. B) Helical
wheel diagram of the helical peptide. C) Mode1 of the helical conformation of the LPD.
The C a trace of the peptide scafSold is grey and the fatty acid chah is black.
The sequence of this designed a-helical peptide was Ala-Om-Ala-Glu-Ala-Ala-Glu-Lys-
Ala-Ala-Lys-Tyr-Ala-Ala-Glu-Ala-Ala-Glu-Lys-a-Ala-Lys-Ala-Om-Ala (Figure
2.1A). Upon folding into an a-helix, it was designed to be amphipathic such that all the
hydrophobic amino acids, in this case alanine, aiign dong one face of the heiix and the
charged, hydrophilic amino acids, lysine and glutamic acid, align dong the other face of
the helix (Figure 2.1 B). Alanine was used as the only hydrophobic amino acid because it
is known to be a strong helix former and it is small in size (Chakrabartty et al., 1994).
The size of this hydmphobic residue plays a role in allowing the covalently coupled fatty
acyl chains to align dong the length of the helicai peptide. Lysine and glutamic acid were
included for two reasons. Firstly, they are strong a-helix formers (Chou et al., 1978) and
secondly, they are placed (i and i+4) apart along the helix in order to form Glu-Lys salt
bridges designed to stabilize helix formation (Marqusee et al., 1987). Tyrosine was
included in the helix to accommodate concentration determination using W absorbance.
Finally, ornithines were placed at positions 2 and 24 of the helix as the sites for the
covalent coupling of the fatty acyl chains that align along the hydrophobic face of the
helix (Figure 2.1 B, C). This coupling of the fatty acyl chains to the peptide was a unique
feature of the desigried peptides. Finally, to reduce the destabilizing charge-dipole effects
of a helix, the N-terminus of the peptide was acetylated and the C-terminus was amidated
(Scholtz, 1 992).
The nomenclature of the LPD senes was LPD-n, where n signified the length of each of
the fatty acyl chains attached to either end of the 25 residue peptide. To investigate a
wide range of fatty acyl cbain lengths, the LPD series synthesized included LPD-10,
LPD- 12, LPD-14, LPD-16, LPD-18, LPD-20, LPD-22, LPD-24 and LPD-28. In
addition, a control peptide with no fatty acyl chains attached, C-O, was also synthesized.
2.2 Peptide Synthesis Strategy
The LPD was synthesized by solid-phase methods as outlined in Figure 2.2. In this type
of synthesis, a protected amino acid is covalentiy coupled through its carboxyl group to a
polymeric support, The peptide is then synthesized C-terminus to N-terminus. Bnefly,
the a-amino protecting group of the terminal amino acid on the polymeric support is
removed to facilitate its coupling to an activated carboxyl group of the next N-terminally
protected residue in the sequence. The Boc and Fmoc chemistry methods are two
approaches to protecting the a-amino group of the amino acids. The Boc method uses t-
butyloxycarbonyl to protect the a-amino group and is acid labile whereas Fmùc uses 9-
fluorenyhnethyloxycarbonyl and is base labile.
In the case of LPD synthesis, a combination of Boc and Fmoc chernistries was used. To
start, a Boc-protected alanine methylbenzhydrylarnine (Boc-Ala-MBHA) resin was
chosen as a suitable resin for reasons mentioned below. This resin is typically used with
Boc chemistry and therefore the covalent coupling between the resin and alanine is not
TFA labile. Upon treating the Boc-Ala-MBHA resin with trifluoroacetic acid (T'FA), the
Boc group was removed to allow the coupling of the subsequent amino acids. The fkee
carboxyl group of the incorning Fmoc-protected residue was activated in the presence of
O-(7-azabenzotiazol- 1 -yl)- 1,1,3,3 -tetramethyluronium hexafiuorophosphate (HATU) to
form an ester that c m react with the a-amho group bound to the support. The remainder
of the peptide was then synthesized using Fmoc chemistry by deblocking the Fmoc
protecting group with pipendine. Once the 25 residue peptide was completed, and the
Fmoc protecting group of the N-terminai residue was deblocked, the fiee terminal a-
amino group was acetylated in the presence of acetic anhydride in acetonitrile- The LPD
was then generated fiom the peptide by coupling aliphatic moieties to either end of the
peptide. This was achieved by selectiveiy Boc protecting the ornithine side chains. Shce
the linker to the MBKA resin and al1 the side chah protecting groups of the amino acids
except ornithine are stable under acidic TFA conditions, only the ornithine 6-amino
groups were exposed upon treating the peptide with TFA. To link the aliphatic moieties
to the peptide, saturated fatty acids were activated with HATU and covalently coupled by
their carboxylic group via an amide bond to the Barnino group of the ornithines. The
final step in the synthesis was the cleavage of the lipopeptide fiom the resin and the
deprotection of al1 the side-chain protecting groups in the presence of
trifluoromethanesulfonic acid (TFMSA). When MBHA resin is treated with TFMSA, an
amidated C-terminal alanine results. This amidated C-terminus, in combination with the
acetylated N-temiinus, were designed to reduce the destabilizing charge-dipole effects of
the helix (Scholtz, 1992).
1 50% TFA
NH2-Ala-MBHA 1 FrnoeOrn *l MN
Fmoc-Orne-Ala-MBHA
20% piperidine
Repeat cycle for each successive amino acid
1 20% piperidine
1 acetic anhydndelpyridine
1 50% TFA
Fatty acid / HATU
--Ala-Om.. . .Om-Ala-MBHA
I l Lipid Lipid
Lipid Lipid
Figure 2.2: LPD synthesis flow chart. Ornithine side chains are protected by Boc,
represented as *.
2 3 Physical Characteriurtion
Following synthesis, the peptide had to be purified to remove any organic impurhies that
arose during synthesis. Initially, purification of the LPDs was attempted using Ci*
reverse-phase (RP) HPLC. However, since the LPDs were signXcantly hydrophobic,
they bound irreversibly. Consequently, a Cq RP-HPLC was the purification system of
choice. The typical ion-pairing agent in HPLC is T'FA as it is non-corrosive and has
excellent separation capabilities. However, since our LPDs were so hydrophobic, HCl
was chosen as a superior ion-pairing agent because it decreased the overall hydrophobic
content of the peptide thereby decreasing its retention within the column and it also
increased the resolution of the peaks. Furthemore, the chloride ion is a more simple
counter-ion to be present in the solution following lyophilization than the TFA counter-
ion.
Following purification, the lipopeptide identities had to be confirmed. Electrospray-
ionization mass spectrometry is a high-resolution technique that identifies the molecdar
weight of peptides to within 0.0 1 % (Chait et al., 1992). Other techniques such as amino
acid analysis or protein sequenators could be used to c o d m the identity of the LPDs,
but ESI-MS is a simple and highly accurate technique that uses very little sample. To
confirm the a-helical conformation of the lipopeptides, circular dichroism (CD)
spectroscopy was used to estimate their secondary structure. In fact, CD was also used to
determine the stability of the LPDs under wide pH and concentration ranges.
Investigating the pH stability and concentration dependence of the LPDs was usefbl
information when preparing crystallization trials.
LPD monomers were designed to have a "wedge" shape (Figure 2.3). More specifically,
the diameter of the amphipathic peptide helix was slightly larger than that of the fatty
acyl moiety aligning dong the hydrophobic face of the helix. Consequently, it was
postulated that the LPDs self-associate into cylindricaLly shaped micelles upon reaching
their cmc. This contrasts with traditional detergents and phospholipids which f o m
spherical micelles and bilayers, respectively (Figure 2.4).
The LPD micelle is arranged such that the hydrophilic face of the peptide is exposed to
the sotvent and the fatty acyl chains orient thernselves by hydrophobic interactions to
form a hydrophobic core (Figure 2.5A). Upon comparing the LPD with traditional
detergent, LPDs were presumed to form a more compact and rigid protein-detergent
complex than traditional detergent-protein complexes (Figure 2.5B). In fact, to
investigate the monodispersity and size of these presumed micelles, gel filtration
chromatography and sedimentation equilibrium ultracentrifugation techniques were used.
Gel filtration or size exclusion chromatography (SEC) is a technique that separates
proteins based on theu size and shape. The number and shapes of the eluted peaks as
well as the retention t h e of the peptide within the column estimate the monodispersity
and molecular weight of the associated system, respectively. Sedimentation equilibrium
ultracentrifugation is a method that can determine the apparent molecular weight of an
associated system using the sedimentation equilibrium equation for a single ideal species.
A rigorous statistical analysis of several data sets using different concentrations at
different velocities are used to accurately determine the molecular weight (Ralston,
1993).
Figure 2.3: LPD modelled using RasMol Version 2.6 - ucbl.0. The LPD monomer is
displayed as its A) top view and B) aad C) side views. The peptide s d o l d is grey-white and
the fatty acyl chahs are black. The LPD monomer possesses a %edge"-shaped geometry .
Traditional
detergent
Phospholipid
Lipopeptide
detergent
Monomer shape
Cone
---- Cy Linder
Wedge
Spherical
Bilayer
Figure 2.4: Cornparison of molecular shapes of monomers and micelles. The hydrophilic
heads of the detergents are grey and the hydrophobie tails are black. The hydrophilic
heads of the phospholipids are blue. (Adapted h m Gennis, 1989)
Figure 2.5: A) Proposed self-assembly of LPD monomers into a cylindrically shaped
micelle B) Cornparison of the stabilization of an IMP by (left) LPDs and (right)
traditional detergents. The LPD is depicted as a grey cylinder representing the
amphipathic helical peptide with two black fatty acid chains attached. The traditional
detergent is depicted as a grey box (hydrophilic head) on a black stick (hydrophobic tail).
The IMP polar domains are represented in red and the hydrophobic domain in green.
2.4 Solubilization of Liposomes
A desirable feature of a detergent is its ability to extract membrane proteins fiom the
membrane bilayer. The investigation into the abiiity of LPDs to breakdown the lameilar
structure of a phospholipid bilayer into lipid-LPD mixed micelles was modelled on the de
la Maza investigation in which a traditional detergent, DDM, solubilized
phosphatidylcholine (PC) liposomes (de la Maza et al., 1997). A liposome is a solvent-
filled vesicle composed of a single phospholipid bilayer. The solubilization of a lipid
bilayer by detergents can be monitored using light scattering. The amount of light
scattered decreases as the lipid transitions fiom king within a liposome of large size to
being incorporated into a comparatively smaller mixed micelle.
2.5 Bacteriorhodopsin Stability Trials
Bacteriorhodopsin (BR) is one of the most extensively studied integrai membrane
proteins. As mentioned, it has been solved to nearly atomic resolution of 1.55 A
resolution within lipidic cubic phases (Figure 2.6) (Luecke et al., 2000). It contains 7
trammembrane a-helices connected by three extemal and three cytoplasmic loops. It
functions as a light-driven proton pump that converts photon energy into an
electrochemical potential. In fact, extensive knowledge concerning its mechanism of
action has been elucidated from the structures solved (Kuhlbrandt, 2000).
Figure 2.6: Ribbon diagram of bacteriorhodopsin (Luecke et al., 1999). The 7 trammembrane
a-helice s are depicted in blue, wnnecting loops in grey and kstrands in red. The retinal Enked
via Schiff base to Lys 216 of BR is iflustrated in yeilow.
BR is an ideal candidate to work with because in its active form, a retinal bound via
Schiff base to Lys 216 confers a purple color that absorbs in the visible spectrum at 550
nm. Upon denaturing, BR becomes a yellow protein that absorbs in the visible spectrum
at 380 nm (Mukai? 1999). Consequentiy, the solubility and stability of BR is easily
monitored by analyzing changes in its absorption spectnun. This thesis uses BR as a
mode1 integral membrane protein to investigate the effectiveness of the lipopeptide
detergents in solubilizing and stabilizing a membrane protein.
A common technique when working with membrane proteins involves an initial
extraction of the membrane proteins fiom the lipid bilayer in a mild, inexpensive
detergent followed by an exchange of this detergent for one more suitable for
crystallization purposes (Michel, 199 1). Since synthesizing LPDs via solid phase
synthesis is a costly process, a standard, reIatively inexpensive detergent, octylglucoside
(OG) was used to extract BR fiom the purple membranes of Hahbacteriurn salinarium
(Landau et al., 1996). This thesis then set out to exchauge the OG for LPD in order to
monitor and compare the solubility and stability of LPD-solubilized BR to OG-
solubilized BR. Traditional methods of exchanging detergents include ion exc hange,
affinity chromatography, sucrose gradient centrifugation or dialysis (Michel, 1991).
However, since the LPDs are synthesized on a small scale and most of these methods
require copious amounts of detergent, an alternative exchange method using very little
LPD was utilized. This alternate exchange method, ultrafiltration, involved a
simultaneous dialysis of OG and concentration of the BR-LPD complex. An
ultrafiltration membrane was used which, upon centrifbgation, allowed OG to pass
through but not the BR or LPD micelles. Not ody did this method minimize the quantity
of LPD required, but it also significantly reduced the time necessary for the dialysis
process. The goal of this investigation was to survey various concentrations of LPDs in
association with BR in order to determine the optimal concentration of LPD necessary to
maintain the solubility and stability of the native BR complex over time. Once BR was
detennined to be stable in the LPD solution, crystallization trials of the complexes began.
2.6 Crystallization Trials
Crystallizing membrane proteins in the presence of detergents is done in essentially the
same marner as with water-soluble prote&. Any number of crystallization methods can
be used including vapour diffision, microdialysis or batch methods (McPherson, 1989
and Michel, 1991). However, the hanging drop vapour difision technique was used in
this thesis. Essentiaily, a concentrated solution of a purified protein was mixed 1 : 1 with a
precipitating solution and hung over a reservoir containing the precipitating solution.
Over time, the protein was brought to supersaturation by the process of vapour diffision
producing either protein precipitate or protein crystals. A sparse rnatrix screen (Jancarik
et al., 1991) was used to test a wide variety of different pH, salts, and precipitants. The
final steps in crystallization included optimizing "leads" fiom the crystal screen by setthg
up a matrix varying pH, or protein, sait or precipitant concentrations.
CHAPTER 3: MATERLUS AND METHODS
3.1 Peptide Synthesis
LPD peptides were synthesized on a 0.2 mmole scale using solid-phase synthesis on a
9050 Plus Pepsynthesizer (PerSeptive Biosystems). t-Butoxycarbonyl-alanine-4-
methylbenzhydrylamine (Boc-Na-MBHA) resin (Advanced ChemTech) was inçubated
for 15 minutes in 50% trifluoroacetic acid (TFA) in dichloromethane @CM) to remove
the Boc amino protecting group. After rinsing with ethanol and filtering the resin, the
dned resin was combined 1 :2 with 150-2 12 Fm glass beads to maintain the integrity of
the resin under high pressure conditions. The peptide was synthesized using 9-
fluorenyLrnethoxycarbonyI (Fmoc) chemistry on the Millipore 9050 Plus Pepsynthesizer.
Each cycle of peptide synthesis consisted of a 2 minute Fmoc deblocking step using 20%
piperidine in dimethylformarnide @MF) at 6.6 mL/& followed by 10 minutes at 3
mlh in . The column was washed with DMF for 14 minutes at 6.6 mL/min to remove
any remaining piperidine. The arnino acid to be coupled was dissolved in DMF in the
presence of the activator, O-(7-azabenzotiazol-1 -yl)- l,1,3,3-tetramethyluronium
hexafluorophosphate (HATU). This solution was recycled at 6.6 mL/min through the
deblocked Ala-MBHA resin for 1 hour and 1 5 minutes. Foliowing an 8 minute wash at
6.6 mL/min with DMF, the cycle was repeated with the subsequent amino acids.
M e r the last amino acid was coupled to the resin and deblocked to remove the tenninal
Fmoc group, the amino terminus of the peptide was acetylated over two hours in 0.5 M
acetic anhydride, 0.5 M pyridine in DMF. To prepare the peptide for coupling with the
fatty acid chahs, the Boc protecting groups on the ornithine side chains at positions 2 and
24 of the peptide were deblocked in the presence of 50% TFA in DCM over 20 minutes-
This deblocking step had no effect on the side chah protecting chains of glutamic acid or
lysine because they contained protecting groups typical of an Fmoc synthesis; a-benzyl
ester and a-2-chloro-benzyloxycarbnyl, respectively. The desired fatty acid (decanoic
acid, dodecanoic acid, tetradecanoic acid, hexadecanoic acid, octadecanoic acid,
eicosanoic acid, docosanoic acid, tetracosanoic acid or octacosanoic acid) (Sigma) was
coupled via an amide bond to the free amùio side chah group of ornithine in the presence
of a 3 fold excess of HATU, 0.17 M DIPEA in DMF over one hour.
As a final step, the LPD was cleaved fiom the resin and the side chah protecting groups
were removed using a cooled solution containing 0.9 mL TFMSA, 1.05 mi, thioanisole,
1 -1 mL m-cresol, and 0.8 mL ethane dithiole in 1 0 mL TFA. After mixing slowly for 2
hotus, the LPD was precipitated overnight in ethyl ether at -20°C. The LPD was washed
several tirnes in cold ethyl ether using a 5 minute centtifiigation step (1000 x g at 4°C).
The washes were complete once the supernatant was colorless and the LPD was a white
precipitate. M e r evaporating the ethyl ether, the LPD was dissolved in approximately
10 rnL water, separated from the resin by filtration, lyophilized and redissolved in water.
A control peptide, C-O, was synthesized in the sarne marner as above with the exception
of omitthg the fatty acid coupling step. This peptide, therefore, contains the same 25
amino acid sequence as the LPDs but does not have any fatty acids attached to it.
3.2 LPD Purification
To desalt and remove some of the organic irnpurities fiom the synthesis mixture, an
initial purification was done using a PD40 Sephadex G-25 M gel filtration column in 0.1
mM ammonium bicarbonate buffer (Amersharn P harmacia Biotech). The hctions whose
spectra had a single peak at 276 nm were pooled and immediately lyophilized. Afier
resuspending the lyophilized LPD in a minimal volume of water, the LPD was M e r
purified ushg a Delta Pak Cq reverse phase (RP) HPLC column (Waters, 300 A pore, 15
pm particle, 25 x 100 mm). The LPD was eluted at 20 ml/min in a 10%-90% gradient at
1% per minute using a 20 mM HCI b e e r A solution and a 30 mM HCl in acetonitrile
buffer B solution. The eiuted fiactions were collected in approximately 15 mL fractions.
3.3 Electrospray lonization Mass Spectrometry @SI-MS)
15 pL of the eluted Cq RP-HPLC fiactions were analyzed by electrospray ionization mass
spectrometry using a PE Sciex API III Plus triple quadrupole mass spectrometer
(perforrned by Dr. Lingjie Meng at the Molecular Medicine Research Centre, Mass
Spectrometry Lab, University of Toronto). The spectrometer was operated at unit
resolution (50% valley definition). Full scan mass spectra were acquired over the mass
range of m/z 500-1500 by s c d g the first m a s spectrometer, QI, using a m/z 0.2 step
size and a 1 ms dwell time. Those fiactions with the caiculated mass correspondhg to
appropriate molecular weight were pooled and immediately lyophilized. The LPD was
redissolved in water to achieve a 1-2 m M concentration.
3.4 LPD Concentration Determination
The concentration of the LPD was determined using the tyrosine absorbance at 276 m
( ~ 4 4 5 0 M%m-'). Absorption spectra nom 200 nrn to 800 nm were obtained using the
Ultrospec 2000 WNisible Spectrophotometer (Pharmacia Biotech). LPD in a denaturing
guanidinium hydrochloride solution had the same absorption as samples in buffer.
Consequently, the concentration of LPD was determined in the appropriate experimental
buffers.
3.5 Circular Dichroism
CD spectra were recorded on a Circular Dichroism Spectrometer mode1 62DS (Aviv) at
25°C. Spectra were obtained using a I mm quartz cuvette fiom 200 to 290 nm with a 0.5
nm bandwidth, 1 nm between points and a 5 second averaging time. In addition to
observing spectra for ail the LPDs at 100 pM in 50 rnM KP04, 200 mM NaCl, pH 7.4, a
pH study (50 p M at pH 3-10) and concentration dependence study (20pM to 140 PM)
were done. A reference baseline was generated in each experiment by subtractuig the
reference buffer data fiom the sample data. Error bars were generated to account for the
uncertainty in the peptide concentrations due to the 0.003 absorbance unit error attributed
by the Ultrospec2000 UVNisible Spectrophotometer.
3.6 Micelle Size Determination
A) Gel Filtration Analysis
100 pL diquots of the 100 PM LPD series in 50 mM KP04,200 mM NaCI, pH 7.4 were
nui on a prepacked Superdex 75 HR10/30 gel filtration column (Amersham Pharmacia
Biotech) at 1 W m i n to detennine the aggregate size of the LPD series. The molecular
weights of the LPD micelles were calculated fiom the equation of the curve defïned by
molecular weight standards.
B) Sedimentation Equilibrium Anaiysis
Sampfes at three different concentrations (-0.2 mM, 0.4 mM and 0.8 mM) were analyzed
using sedimentation equilibrium ultracentrifugation (Beckman - Optima XL- 1 Analytical
Ultracentrifiige) to determine the monodispersity and size of the LPDs (performed by
Sandy Go, Ontario Cancer Institute, Department of Medical Biophysics, University of
Toronto). The LPDs were dissolved in 50 mM IWO4, 200 mM NaCl, pH 7.4. The
sarnples were analyzed at 20°C at 280 nm wavelength, measurements king made every
0.001 cm using 10 replicates. n i e SedNterp software was used to calculate the solvent
density and the partial specific volumes of the peptides. The rotor speeds used for each
LPD data set are found in Table 3.1. Data fitting and analysis were performed with
Microcal Ongin 4.1. Global analysis of the nine data sets for each LPD was fit to an
equation for a single ideal species to yield an apparent molecular weight.
Table 3.1 : Rotor speeds used for sedimentation equilibrium analysis
SAMPLE ROTOR SPEEDS (rpm)
C-O (control peptide)
LPD- 12, LPD- 14, LPD- 16
35000,40000,44000,48000
20000,25000,35000
LPD- 18
LPD-20
25000,30000, 35000
15000,20000,25000
LPD- 12 solubilized BR
LPD-20 solubilized BR
6000,9000,12500
6000,9000,12500
3.7 Liposome Solubilizrition
A) Liposome Preparation
Egg phosphatidylcholine (PC) (Avanti Polar Lipids) dissolved in chloroform was dried
under a strearn of nitrogen. The lipid was hydrated to 1 mM using DLS buffer (10 mM
N-2-hy&oxyethylpiperazine-N'-2-ethanesdfonic acid (HEPES), 200 mM NaCl, pH 7.2)
using a I hour incubation on a Nutator. Unilamellar liposomes were obtained by
extrusion through a 100 nm polycarbonate membrane (Avestin) using a LiposoFast-Basic
mini-extruder (MacDonald et al., 1991). PC liposomes were diluted to 0.1 m M using
DLS buffer and incubated overnight (20-24 hours) in varying concentrations of
dodecylmaltoside (DDM) detergent, C-O or LPDs.
B) Phosphate Determination
The phospholipid concentration was determined by measuring the phosphate
concentration of the liposome solutions (Ames, 1960). Samples (25-100 PL) were mixed
with 0.3 mL of 10% magnesium nitrate in ethanol. The solvent was slowly evaporated to
dryness over a flame until the brown fumes disappeared. Care was taken to ensure the
ethanol did not ignite during this drying process. After the test tube had cooled, 0.3 mL
of 0.5 M HCl was added. The test tube was then heated in a boiling water bath for 15
minutes to hydrolyze any pyrophosphate that was formed in the ashing procedure to
inorganic phosphate. A marble was placed over the test tube to prevent evaporation
during boiling. M e r cooling the test tubes to room temperature, 0.7 mL of a cooled
ascorbic acid - moiybdate mixture (1 part 10% ascorbic acid to 6 parts 0.42% ammonium
molybdate in 1 M sulfiiric acid) was added and the solution was heated at 37°C for one
hour. Finally, the solution was cooled to room temperature and the optical density (OD)
at 820 nm was read. The phospholipid concentration in the liposome solutions were
extracted fiom the equation of the line of a standard potassium phosphate curve (0-75
nmol).
C) Light Scattering
The DynaPro-801 Dynamic Light Scattering / Molecular Sizing Instrument (Protein
Solutions) was used to monitor the ability of detergent or LPDs to solubilize the PC
liposomes by rneasuring the total amount of light scattering as well as the size
distribution (Rh) and polydispersity of the samples determined by dynamic light
scattering. Approximately 10 readings were recorded for each sample at 100% APD bias,
5 second maximum acquisition t h e and a S N threshold of 1. DynarnicsB software was
used to analyze the data. Those values with a polydispersity below 25% were considered
monodisperse and those with a baseline iess than 1.005 and a SOS less than 5.000 were
considered monomodal according to the manufacturer's recommendations.
3.8 Purple Membrane Isolation and Bacteriorhodopsin Purification
A) Purple Membrane Isolation
The purple membranes were isolated fiom Halobacterium salinarium essentially as
described (Oesterhelt et al., 1974) with a few exceptions. A Microbank bead (Pro-lab
Diagnostics) containing Halobacterium salinarium (a gift fiom Dr. L. Lanyi, UC I d e )
was transferred to 5 mL standard growth media (SGM) containing 1 pg/mL of
novobiocin (Sigma). Standard growth media at pH 7 consisted of 4.3 mM sodium
chlonde, 8 1 mM magnesium sulfate heptahydrate, 1 0.2 mM sodium citrate, 26.8 mM
potassium chloride, lOg/L bacteriological peptone (Oxoid), 1.36 pM calcium chloride,
27.5 pM zinc sulfate heptahydrate, 12 PM manganese sulfate, 12 FM ferrous ammonium
sulfate hexahydrate, 3.36 pM cupric sulfate pentahydrate. After a 5- 10 day incubation at
40°C, 3 rnL of this preculture was used to inoculate 300 mL SGM containhg 1 pg/mL
novobiocin. After a 3 day incubation at 40°C, 16 mL of this culture was used to
inoculate 800 mL SGM containing no novobiocin. After 5-10 days, when the cdture
reached the end of exponential phase and the media had a purplish hue, the cells were
harvested by centrifuging at 16000 x g for 10 minutes at 4OC. The cells were
resuspended in LOO mL 4 M NaCl and 0.5 mg/L DNaseI (Sigma). The cells were lyzed
by osmotic shock using an overnight dialysis against 12 L 0.1 M NaCl at 4OC with a 12-
14 kDa molecular weight cut off membrane (Spectrum). 'Ihe membranes were washed 2-
3 times in 0.1 M NaCl by centrikghg at 1 00,000 x g for 60 minutes and homogenizing
with a Teflon pestle in 0.1M NaCl. Finally, the purple membranes were isolated by
overlaying the PM over a 40%/60% sucrose density gradient and centrifuging at 75,000
x g overnight at 4OC. The PM formed a band at the interface of the two sucrose
solutions. After extracthg the purple membranes fiom the sucrose gradient, the samples
were stored at -80°C.
B) Bacteriorhodopsin Purification
BR was purifed fiom the purple membrane as described with a few exceptions (Landau
et al., 1996). The sucrose concentration was reduced fiom the purple membranes by
diluting the solution 1 :20 using 0.1 M NaCl. Following centrifugation (1 hour, 100,000 x
g at 4"C), the purple pellet was homogenized using a teflon pestle in 25 mM NaP04, pH
6.9 and adjusted to a final concentration of 1.5% (51 mM) f3-octylglucoside (OG)
(Anatrace) using a detergent-to-protein ration of 30: 1. The BR concentration was
detennined at 550 MI (E = 54000 M-' cm-')* For maximal BR solubilization, the solution
was incubated for at leas 36 hours in the dark. m e r adjusting the pH to 5.5 with 0.1 N
HCI, the soluble BR was retrieved fiom îhe supernatant foiiowing centrifugation for 45
minutes at 200,000 x g and 4OC. BR was then concentrated to approximately 5 mg/mL
using a PM 10 Arnicon membrane at 50 psi and purified at 1 mWmin in 25 mM NaP04
pH 5.5 in 1 -2% (4 1 mM) OG over a Superdex 75 gel filtration column. The purified BR
sarnple was assayed for purity using a 10%-20% SDS polyacrylamide îricine gel
incubating the samples 1: 1 with Tricine sample buffer.
3.9 Bacterio thodopsin Stabilization
A) Detergent Exchange
LPD in water was exchanged into 50 mM NaP04, ISO mM NaCl, pH 7.4 using
centrifugation through a Biomax 5K NMWL membrane (MiIlipore) at 7500 x g for 5
minutes. The appropriate v o b e of purified BR in 25 mM NaP04, 1.2%OG, pH 5.5 was
added to the exchanged LPD to achieve a final LPD:BR concentration ratio of 0.5 mM
LPD:22 pM BR. The sample was centrifûged at 7500 x g for 5 minutes through the
Biomax 5K NMWL membrane. The buffer volume was adjusted to that of the desired
fmal volume to achieve a 22 pM BR concentration and centrifuged again. To ensure the
total removal of OG, this process was repeated 4 times. A negative control was prepared
by exchanging the BR into a bmer containhg no detergent. Likewise, a positive control
was prepared by exchanging the BR into the exchange b a e r containing 1.2% OG.
Monitoring the absorbance of the retentate at 276 n m demonstrated that essentidy 100%
of the LPD samples were retained following the exchange procedure. However, when
exchanging the OG for C-O, a significant amount of the C-O flowed through the Biomax
membrane. To accommodate for this, the appropriate amount of C-O was added after
each centrifugation step to ultimately reach the desired ratio of C-O to BR.
B) Detergent Assay
The concentration of OG remaining in the exchanged samples was determined using the
colorimetric assay for carbohydrates as descnbed (Dubois et al., 1956). A standard curve
of OG (0-200 m o l ) and 5-25 PL of the exchanged samples were prepared to a fmal
volume of 1 mL, adjusting the volume with water. To detect the carbohydrate groups, 50
pL phenol was mixed in followed by 2.5 mC concentrated sulfùric acid. After mixing,
the sarnples were cooled to room temperature before reading the OD at 490 m. The
detergent content was calculated based on a standard curve of a stock OG solution (0-200
nmol).
C) Solubility and Stabiiity Analysis of Bacteriorhodopsin
The exchanged BR samples were stored at room temperature in the dark. They were
monitored for solubility and stability by centrifuging at 130,000 x g for 30 minutes on a
Beckman AirfÛge at O, 1, 4, 7, 14, 21 and 28 days. The soluble supernatant was
monitored using an absorption spectnim nom 200 to 800 nm obtained using an Ultrospec
2000 UVNisible Spectrophotometer (Pharmacia Biotech).
D) Sedimentation Equilibrium Analysis of the LPD Solubilized BR
LPD-12 and LPD-20 solubilized BR samples were prepared as in the detergent exchange
procedure above. Three different BR concentrations (-4 PM, 8 PM, 16 CrM) in 50 mM
NaPO,, 200 mM NaCl, pH 7.4 were analyzed using sedimentation equilibrium
ultracentrifugation (Beckman - Optima XL-1 Analytical Ultracentrifuge) to determine
the aggregate size of the LPD-BR complex (performed by Sandy Go, Ontario Cancer
Institute, Department of Medical Biophysics, University of Toronto). Data was collected
at 20°C at the rotor speeds found in Table 3.1. The data was analyzed at 550 nm
wavelength, measuring every 0.001 cm using 10 replicates. The SeciNterp software was
used to calculate the solvent density and the partial specific volume. Data fitting and
analysis were performed with Microcal Origin 4.1. Global analysis of the nine data sets
for each LPD-BR complex was fit to an equation for a single ideal species to yield an
apparent molecular weight.
3.10 Crystallization Trials
Crystallization trials of the LPDs and LPD-solubilized BR complexes were prepared by
the vapour diffusion technique using sparse-matrix screens (Jancarik et al., 1991), Crystal
Screen 1 and il (Hampton Research). 1+1 pL drops were prepared of 48 mg/mL C-O over
an 800 PL reservoir for Crystal Screen 1. Optllnization of condition #46 (0.2 M calcium
acetate hydrate, 0.1 M sodium cacodylate pH 6.5, 18% w/v polyethylene glycol 8000)
was done in 0.5 + 0.5 pL drops placed above 800 pL reservoirs by reducing the
concentration of C-O in 5 mg/mL increments fiom 40 mg/mL to 5 mg/mL. Similarly, l+l
pL drops were prepared of 5 mg/mL LPD- 14 over a 1 mL reservoir for Crystal Screen 1.
No optimi;rsition was done using LPD-14. Lastly, 0.5 + 0.5 pL drops of 7 mg/mL LPD-
16 were prepared of Crystal Screens 1 and II over 800 pL reservoirs. Optimization of
condition #26 (0.2 M ammonium acetate, 0.1 M tri-sodium citrate dihydrate pH 5.6,30%
V/V a-methyl-2,4-pentaaediol) was done using 0.5 + 0.5 pL drops over 1 mL reservoirs
varying the MPD in 2% increments fiom 33% to 43% and varying the ammonium acetate
in 0.05 M increments fiom O. 1 M to 0.25 M.
LPD-20 solubilized BR samples were prepared as descnbed (3 -9A Detergent Exchange).
The BR was concentrated to 7 mg/mL BR and 1+1 pL drops of the LPD-20 solubilized
BR were prepared over 1 rnL reservoirs using Crystal Screen 1. AL1 trays were incubated
in the dark at room temperature.
CHAPTER 4: RESULTS
4.1 LPD Synthesis and Purification
An LPD series with fatty acyl chain lengths successively increased by 2 acyl units was
generated. The series synthesized extended fiom LPD- 1 O to LPD-28.
Following synthesis and a PD-1 O Sephadex G-25 M gel filtration purification, a number
of organic i m p d e s remained in solution (Figure 4.1A). However, RP-HPLC using a C4
column removed these impurities (Figure 4. IB). Fractions fiom the major peak off the
HPLC column were analyzed by ESI-MS and oniy those fiactions with greater than 90%
purity were pooled. The observed ESI-MS identities of the synthesized LPDs were within
0.025% of their calculated values (Figure 4.2, Table 4.1).
4.2 Characterization of the LPD Series
A) Hydrophobicity and Solubility
RP-HPLC works on the premise that hydrophobic peptides interact more strongly to a
nonpolar aliphatic stationary phase (C4 silica) than the polar mobile phase. Upon
introducing increasing concentrations of a nonpolar solvent such as acetonitrile, the
solvent cornpetes with the hydrophobic peptide for the nonpolar stationary phase causing
the peptide to desorb fiom the stationary phase and elute fkom the column (Liu et al.,
1999). W-HPLC, therefore, can be used as a means to determine the relative
hydrophobic content of peptides.
% acetonitrile % acetoni trile
Figure 4.2: ESI-MS spectrum for LPD- 12 following C, RP-HPLC purification. The
molecular weight is determined by identifying a fmily of m/z peaks and attributing
the appropriate charge states to them (table insert). The mass i s calculated according
to the foilowing equation: m = m/z (2) - z, where m is the rnass and z i s the charge. In
this case, the observed mass was 2839.39.3 ghol which is within 0.01% of the
calculated mass of 2839.1 ghol.
Table 4.1 : Cornparison of LPD calculated and observed molecular weights
l C-O I 2474.5 I 2474.37 + 0.0 1 I
ESI-MS M.W. LPD
M. W. = molecular weight (@mol)
Calculated M- W.
LPD- 16 1
295 1.3 295 1.65 + 0.05
In this case, RP-HPLC demonstrated that an increase in the fatty acyl chah length of our
LPD series, resulted in an overall increase in hydrophobie content of the LPDs. This was
demonstrated by the increased concentration of acetonitrile required to desorb the LPD
fiom the coIumn (Figure 4.3).
In fact, this increase in hydrophobicity of the LPD series with increased fatty acyl chah
length correlated with an increased difficulty in maintaining their solubility. The LPD
senes that was synthesized included LPD-IO through to LPD-28. However, LPD-22 and
LPD-24 were only partly soluble in water pnor to the gel filtration step. Similady, LPD-
28 was cornpletely insoluble in water prior to the gel filtration step. Consequently, the
LPD series that was extensively investigated was LPD-12 to LPD-20. Foilowing RP-
HPLC purification and lyophilization of this LPD series, some difficulty was also
observed in maintaining the solubility of highly concentrated LPD solutions in water.
However, these LPD solutions were initially fiozen at extremely hi& concentrations of
40 mM or greater. Upon thawing, a "gel-üke" phase formed which would not redissolve
upon dilution with water. Organic solvents such as trifluoroacetic acid and
trifluoroethanol were used to facilitate solubilization but were unsuccessful at
maintaining the solubility over t h e . Once the LPD series was diluted with water to
concentrations of 2 mM or less and stored at 4OC the LPD solubility was maintained over
a long period of tirne.
C-O LPD-12 LPD-14 LPD-16 LPD-18 LPD-20
Figure 4.3: C, RP-HPLC elution of the LPD series. The retention time of the LPD
increases with increasing lipid acyl chain length indicating that an increase in fatty acyl
chah length confered an increase in the hydrophobie content of the LPD.
B) Secondary Structure
To confirm the design of the desired a-helical peptide, the secondary structure of the
LPD series was chmcterized using circular dichroism (CD) spectroscopy. A wavelength
scan fkom 200 nm to 270 nm Vigure 4.4) demonstrated that the LPDs were a-helical as
they displayed CD minima characteristic of a-helices at 208 nm and 222 nm. The
control peptide, C-O, however, contained significantly less helical content as
dernonstrated by its reduced eilipticity. Furthemore, the spectral minima at 208 nm
observed for the LPDs was replaced by a minimum at 205 nm for C-O.
Using CD spectra to view LPDs exposed to a wide pH range (pH 3-10) demonstrated that
LPD secondary structure was essentially stable over this pH range but experienced a
slight decrease in helical content at neutral pH (Figure 4.5). Observations of the LPDs in
solutions of greater than 2 mM demonstrated a similar pH optimum of 4 to 6 and 8 to 10
such that maintainhg the LPD in a pH between 6 to 8 reduced the solubility of the LPD.
CD spectra were also used to determine if there was a concentration dependence on
secondary structure. Figure 4.6 demonstrates that LPD secondary structure wits not
dependent on concentration over the range of 20 p M to 120 pM.
-35000 ! I . 1
200 220 240 260
wavetength (nm)
Figure 4.4: CD wavelength scan of the LPD series. CD spectra were obtained using 100
LPD Ui 50 mM KPO,, 200 mM NaCl, pH 7.4. The LPDs contain strong helical
secondary structure, as demonstrated by the minima at 208 nm and 222 nm. The control
peptide (C-O) contains Iittle helical secondary structure as demonstrated by the reduced
ellipticity and shiM 208 nm minimum.
Figure 4.5: pH dependence on secondary structure. CD spectra were obtained for
(+) 50 pM C-O and (B) 50 ph4 LPD-14 in 50 mM KPO,, 200 mM NaCl.
C-O concentration (PM)
O 20 40 60 80 100 120 140
LPD- 12 concentration (PM)
Figure 4.6: Concentration dependence of LPDs on secondary structure. CD spectra were
obtained using 50 mM K m , 200 rnM NaCl, pH 7.4.
C) Mice lie Size
SEC and sedimentation equilibrium ultracentrifugation were used to investigate whether
the LPDs form a micelle of a defined size or simply aggregate into ill-dehed assemblies.
A comparison of the elution profiles of the LPD series fiom the Superdex 75 SEC
col- clearly indicated that an increase in the fatty acyl chain length on the LPD
decreased its retention tirne within the column (Figure 4.7). Furthemore, a single peak
was observed for every member of the LPD series. This suggests that oniy one moIecuIar
species was present for each LPD. Estimates of the molecular weights for these LPD
species were achieved fiom rnolecular weight standards (Figure 4.8). This molecular
weight detennination indicated that the molecular species present was a micellar form of
LPD rather than its monomeric form. This data also demonstrated that an increase in the
fatty acyl chah length of the LPD also constituted an increase in aggregation number, the
number of monomers per micelle (Table 4.2). The aggregation number ranged from 9 to
13. The control peptide, C-O, however, eluted as an apparent trimer.
The micelle sizes fiom sedimentation equilibrium ultracentrifugation were obtained using
partial specific volumes, vO, calculated using SedNterp software. The calculated v* for
the control peptide, C-O, was 0.7463 cm3/g and the LPD values ranged from 0.7990 crn3lg
for LPD-12 to 0.7946 cm3/g for LPD-20. The sedimentation equilibrium
ultracentrifugation results were striklligly similar to the SEC results. A global analysis of
al1 the data sets for each LPD was fit to the sedimentation equilibrium equation for a
single ideal species to yield an apparent molecular weight (Table 4.2).
O 200 400 600 800 1 O00 1200
time (seconds)
Figure 4.7: Elution profile of LPDs on Superdex 75 HR 1 OB0 SEC column. 100 pM LPD in
50 mM KPO,, 200 mM NaCl, pH 7.4 were nui at 1 mlfmin.
i C-O i
LPD-12 !
LPD-14 1
Eluti on volum e ( d )
Figure 4.8: Estirnate of LPD micelle size using the calibration molecular weight
standards. 100 pM LPD in 50 mM KPO,, 200 mM NaCl, pH 7.4 were run on a Superdex
75 HR10/30 SEC column. The molecular weight standards mcluded vitamin B12
(1.3554 kDa), ribonuclease A (13.7 ma), chymotiypsmogen A (25 kDa) and bovine
semm al bumin (67 kDa).
Table 4.2: Cornparison of gel filtration chromatographie and sedimentation equiiibrium
ultracentrifugation detedat ion of micelle size
LPD
C-O
LPD- 12
LPD-14
M. W. = molecular weight (&mol) N = aggregation number; number of monomers per micelle * = represents samples that were non-ided or heterogeneous
Gel Filtration Determination
LPD- 16
LPD- 18
Sedimentation Equiiibrium Analysis
Micelle M.W.
7969
25454
30045
33563
36129
N
3 -22
8.97
10.3 8
Micelle M. W.
1979
23228
26858'
N
0.8
8.18
9.28
9.78
12.33
1 1.37
12.0 1
28438'
37095'
Figure 4.9 represents the best fit of the data for LPD-12. The LPD micelles were
considered monodisperse if the in (abs) versus radius squared plot produced a Linear
relationship. Although LPD- 14, LPD- 16 and LPD- 1 8 solutions were considered slightly
non-ideai and heterogeneous, the general trend that an increase in fatty acyl chain leogth
constituted an increase in aggregation nurnber could still be drawn fiom the data. The
aggregation numbers of the LPD series ranged f?om 8 to 12. Furthemore, modelling the
monodisperse sedimentation data in terms of a monomer to micelie equilibrium produced
association constants in the order of 1oL9' to 1o2I4 which indicates that essentially no
monomenc species was detectable in the solution.
The control peptide, C-O, appeared to be monomeric according to the sedimentation
equilibrium ultracentrifugation data which contrats with the trimenc form determined by
SEC. However, neither technique accurately determined the molecular weights of the
LPDs and C-O. For SEC, which is dependent on the shape of the macromolecde, the size
of the macromolecules were estimated from the equation of the line generated by
sphericaïly shaped molecuiar weight standards. Similarly, for the sedimentation
equilibrium ultracentrifugation analy sis, the calculated partial speci fic volumes may
differ from the actual experimentally determined values. In fact, figure 4.10
dernonstrates that a srnaIl change in partial specific volume significantly affects the
molecular weight of the species.
Radius Figure 4.9: Sedirnentation equilibrium ultracentrifugation of LPD-12 run at 25000 rpm.
The LPD series were analyzed at 280 nm in buffer containing 50 mM KPO,, 200 mM NaCl,
pH 7.4.
Figure 4.10: Caiculation of the apparent moleculat weights of (+) LPD-12 and (i) LPD-20 with
respect to changing partiai specific volume, v O .
D) Liposome Solubilization
The solubilization of 0.1 mM PC liposomes by DDM, C-O peptide and LPDs was
andyzed through the changes in the iight scattered by these systems 24 hours after the
addition of the detergents. Figure 4.1 1 shows the solubilization curves of liposomes
titrated with increasing concentrations of detergents, DDM and LPD-14. In generai, the
total light scattered is proportional to the concentration and hydrodynamic radius of the
particles in the solution. An increase in the total light scattered and hydrodynamic radius
were observed initially. This increase was due to the initial incorporation of the detergent
molecules into the lipid bilayers making the liposomes larger as well as liposome fusions.
However, upon the addition of increasing detergent concentrations, the lipid bilayer was
solubilized which caused the phosphotidylcholïne monomers to transition fkom their
bilayer state to form stable mixed micelles with detergent. This was demonstrated by
lower levels of light scattering and significantly smaller hydrodynamic radii compared to
the original liposomes. The hydrodynamic radius of the 0.1 mM PC liposomes was 33.8
nm with a 9.6 nm polydispersity, as determined by dynamic light scattering.
Polydispersity represents the particle size distribution. Since the polydispersity was
greater than 25%, the liposome suspensions were considered to be slightly
heterogeneous.
Figure 4.12 displays the summary of results for the dynamic light s c a t t e ~ g data obtained
for the solubilization of the 0.1 mM PC liposomes. The positive control, DDM,
completely solubilized the bilayer at a concentration of approximately 0.8 mM producing
a micelle with a hydrodynamic radius of 3.7 nm. The control peptide, C-O, at
concentrations up to 2 mM, however, had essentidy no effect on the size of the liposome
vesicles. In fact, the average hydrodynamic radius remained in the same range of the
initial liposomes but the polydispersity of the systern increased. In cornparison, LPD-12,
LPD-14 and LPD-16 demonstrated the ability, similar to DDM, to fully solubiiize the PC
lipid bilayer into mixed micelles using LPD concentrations beyond 1 mM. These mixed
micelles had hydrodynamic radii of 2.3 nm, 2.53 n m and 2.5 nm, respectively. In
addition, these mixed micelle systems had polydispersities of less than 25% indicating
that these systems were monodisperse. LPD-18 and LPD-20, on the other hand,
produced heterogeneous solutions in the presence of the PC liposomes with LPD
concentrations up to 2 mM; the hydrodynamic radii were larger than the liposomes alone
and had larger polydispersities (data not shown). This suggests that LPD-18 and LPD-20
interact with the liposomes but not enough to break up the vesicles. Further
investigations with higher concentrations of LPDs were not pursued because using
concentrations beyond 2 mM would have consumed unreasonable amounts of the LPD
solutions.
O 0.25 0.5 0.75 1
DDM concentration ( m m
O 0.5 1 1.5
LPD C- 14 concentration ( m m
Figure 4.1 1 : Cornparison of concentration tieation of 0.1 mM PC liposomes with A) DDM
and B) LPD-14. These data are single trials that are representative of repeated trials. The
photons scattered represents the tdal scattering of iight and the hydrodyrnunic radius (Rh) was
detenni ned by the autocorrelation func tion fiom the dy nam ic 1 ight scattering data.
liposomes DDM C-O LPD-12 LPD- 14 LPD-16
Figure 4.12: Summary of 0.1 mM PC liposomes solub il ization. 0.1 mM PC liposomes
were incubated in the presence of detergents for 24 hours before obtaining the iight
scattering data The error bars represent the polydispersity of the DLS readings.
DDM serves as the control detergent, achieving complete solubilization at 0.8 mM.
The negative control, C-O, f~ led to solubilize the PC liposomes at 2 mM. LPD- 12,
LPD-14 and LPD- 16 solubil ized PC liposomes at concentrations of 1.25 mM, 1.5 mM
and 1 mM, respectively. This data represent one data trial but is representative of
repeated trial S.
4.3 Bacteriorhodopsin Purification and Stability Triais
Purple membranes (PM) containhg BR were isolated fiom H. salinarium cells (Figure
4.13A). The purification of BR in the presence of octylglucoside (OG) nom the PM
produced pure protein that has an apparent molecular weight on an SDS polyacrylamide
gel of approximately 20,000 kDa (Figure 4.13B).
To determine the stability of BR in the presence of LPDs, the OG had to be replaced with
LPDs. Detergent exchange was achieved using 5 successive dilution/concentration steps
using ultrafiltration membranes (Figure 4.14). After the 5 cycles, no OG was detected in
the retentate. This ensured that the effects observed when BR was in the presence of
LPDs was attributable to the LPD and not due to residuai OG following the exchange.
The first step in determining the effectiveness of a detergent in solubilizing a membrane
protein is to determine the optimal concentration of LPD to achieve complete
solubilization. While insufficient detergent concentration results in precipitation of the
protein, excess detergent can result in denaturation of the protein andor phase separation
(Tribet et al., 1996). Figure 4.15 displays the SDS polyacrylamide gel of a LPD- 12
concentration gradient (0.05 mM to 2.5 mM) in association with 22 pM BR immediately
following detergent exchange. This gel demonstrates that LPD- 1 2 at concentrations
greater than or equal to 0.25 mM fully solubilized 22 y M BR. Furthemore, the absence
of the LPD-12 band in the gel for concentrations of 0.1 mM or lower suggests that LPD-
12 interacted with BR and precipitated out of solution dong with the BR followbg the 30
minute 130,000 x g centrifùgation step.
Eluted BR fiactions
+-l
Figure 4.13 : PM isolation and BR purification A) 40%/60% sucrose density gradient
isolation of PM B) GELCODE Blue Stained 109&20% SDS polyacrylamide Tricine gel
of BR purification using the Superdex 75 gel filtration column eluting in 25 mM NaPO,
and 40 mM OG.
1 2 3 4
Number of exchange washes
Figure 4.14: OG remaining in retentate following the exchange wash steps. The 50 mM OG
used to solubilize 22 pM BR was replaced with the appropriate concentration of LPD in 50
mM NaPO,, 150 mM NaCl, pH 7.4. The OG content remaining in the retentate was
determined using the coIorimetric assay for carbohydrates (Dubois et al., 1956). The dashed
horizontal Iine at 25 rnM OG represents the cmc of OG.
Figure 4.15 : Coomassie stained 10%-20% SDS polyacrylamide tricine gel de termining the
minimum LPD-12 concentration required to fully solubilize BR. Samples were centnfuged
at 130,000 x g for 30 minutes and the supernatants were mixed 1:l with 2x Tricine sample
buffer.
These LPD-12 solubilized BR samples were also monitored spectrophotometrically to
determine the conformational state of the BR over the. Figure 4.16 displays this time
course in terms of its absorption at 550 nm for different LPD-12 concentrations. Within
the concentration range of 0.25 mM to 2.5 mM, al1 the LPD-12 concentrations
maintained BR in its native state over the 32 day period.
Similady, concentration gradients were prepared for al1 the other members of the LPD
series, LPD-14 to LPD-20 (data not shown). This data determîned that 0.5 mM LPD was
the minimum concentration required for al1 the LPDs to maintain maximal solubiiity and
stability of 22 FM BR over time. Figure 4.17 displays the time course monitoring the
stability of the 0.5 mM LPD-solubilized BR in cornparison to BR in the absence of
detergent, in the presence of OG or in the presence of the controI peptide, C-O. This
figure demonstrates that BR in the absence of detergent or in the presence of C-O was not
soluble; no protein was recovered in the supernatant following the 130,000 x g
centrifugation step. In fact, purple pellets were obtained for these conditions which
confirmed that the precipitation was due to insoluble native BR rather than denatured BR-
LPD-IS to LPD-20, however, were d l capable of maintainhg the native soluble state of
BR over the 32 day period. Finally, this tirne course also demonstrated that OG-
solubilized BR quickly destabilized within one week of storage. Furthermore, upon
analyzing the spectra fkom this time course (Figure 4.18), it is evident that the OG-
solubilized BR denatured over time but remained soluble; the native state Abssso peak
decreased with a concomitant increase in the denatured Abs380 peak. The LPD-12
solubilized BR spectra, on the other hand, changed minimally over the month time fiame
which concludes that LPD-12 solubilized BR was a very stable system. Although data is
not shown for LPD-24 and LPD-16 at 0.5 mM, these LPDs also proved to stabilize BR
over the 32 day penod. In contrast, figure 4.19 demonstrates that the longer fatty acyl
chah length LPD-solubilized BR samples, LPD-18 and LPD-20 (data not shown),
aggregated slightly over tirne. One of two situations occurred. If the aggregates were
insoluble, a decrease in the AbssSo peak accompanied by a decrease in baseline occurred.
However, if the aggregates remained soluble, an increase in the Abssso peak was
observed that was accompanied by an increase in the baseline. Ln addition, aggregation
was more pronounced when higher concentrations of LPD were used (Figure 4.19 B).
Sedimentation equilibrium ultracentrifugation anaiysis was performed on LPD-12
solubiiized BR (Figure 4.20) and LPD-20 solubilized BR (data not shown). Calcuiating
the partial specific volume based on one molecule of BR complexed with one molecule
of LPD and performing a global fit to all the 9 sedimentation equilibrium profiles yielded
apparent molecular weights of 172,918 glmol 2 3.4% and 214,129 g/mol + 4.8% for
LPD-12 and LPD-20 solubilized BR, respectively. Based on the assumption that the BR
formed a trimer as in its native state (Landau et al., 1996), 30.7 to 34.8 LPD-12
molecules and 40.5 to 47.1 LPD-20 molecules were available to solubilize the BR trimer.
Preliminary modelling of this assumption using Swiss PDB Viewer and displaying it
using RasMol Version 2.6, demonstrated that this stoichiometry is feasible (Figure 4.21).
Approximately 25 LPD monomers fit around the BR trimer. In fact, it is possible that the
excess LPD, in the form of micelles, could have contributed to some of the heterogeneity
that was detected in these systems.
! + no detergent 1 j
i+49mMOG f 1-+0.05 mM : '-0.1 mM i
i+-0.25mM I ! I
'-0.5mM ,
j t l m ~
a
Figure 4.16: Time course monitoring the stability of LPD-22 solubilized BR to
determine the minimum LPD-12 concentration required to maintain the solubi E î y of
BR. 22 jM BR in 50 mM NaPO,, 150 mM NaCl, pH 7.4 was solubilized in various
concentrations of LPD- 12.
tirne (days)
Figure 4.17: Time course monitoring the stability of 0.5 mM LPD solubilized BR. 22 pM
BR in 50 mM NaPO, 150 m M NaCl, pH 7.4 was solubilized in 75 mM OG or 0.5 mM
LPDs. Spectra were taken foilowing a 30 minute centrifugation at 130,000 x g using a
Beckman airfuge.
250 350 450 550 650
wavelength (nm)
250 350 450 550 650
wavelength (MI)
-Day O 1 -Day4 i
-Day 7 1 1
-Day 14/ l - Day 21 l - Day 32 j
-DayO - Day 2
, Day 7 ;- Day 15 i I
l- Day 21 i 1
i-Day 31 '
Figure 4.1 8: Spectni monitoring the stabi li ty of BR solubil ized over a 30 day period. 22 phi BR in
50 m . NaPO, 150 mM NaCL, pH 7.4 was solubilized in A) OG or B) 0.5 rnM LPD-12. Spectra
were taken over a 30 &y period following a 30 minute centrifugation at 130,000 x g using a
Beckman airfûge prior to each reading.
250 350 450 550 650
wavelength (nm)
;- ~ a y 2 i i Day 7 i-Day 15 ' - ~ a ~ 22 - Day 32
2.5 - Day O
2 l
- Day 2 I
1.5 Day 7 s m s - Day 15 6 1 - Day 22
- Day 32 l
0.5
O 250 350 450 550 650
wavelength (nm)
Figure 4.19: Cornparison of LPD- 18 spectra monitoring the stability of BR solubilized
over a 32 day period. 22 pM BR in 50 mM NaPO,, 150 mM NaCl, pH 7.4 was
solubilized in A) 0.5 mM LPD-18 and B) 2.5 mM LPD-18. Spectra were taken
foIlowing a 30 minute centrifugation at 130,000 x g using a Beckman airfige prior to
each reading.
Figure 4.20: Sedimentation e q d ibrium ultracentrifugation of LPD-12 solubi lized BR run at
9000 rpm . The LPD-solubi lized BR samples were anal yzed at 550 nm in buEer containing 50
mM NaPO,, 200 mM NaCI, pH 7.4
Figure 4.2 1 : Preliminary model h g of an LPD solubi lized BR trimer viewed normal to the
membrane dong the BR three-fold axis (model led using Swiss PDB Viewer). The BR
carbons are dark grey, mtrogens are Mue and oxygens are red. The LPD peptides are
depic ted in light grey with the fatty acy 1 chahs in Mack. 25 LPD monomers align around
the BR trimer.
As in the case of the determination of LPD micelle size using sedunentation equiiibrium
ultracentrifugation, the apparent molecular weights derived for LPD-BR complexes fiom
this ultracentrifugation experiment were not accurate because these molecular weight
determinations were calculated based on estimated partial specinc volumes of the LPD-
BR complexes. In this case, these calculated v O values were largely umeliable due to lack
of information regarding the stoichiometry of the LPD-BR complex as well as the
difficulty predicting the effects of the fatty acyl moieties on the partial specific volume.
In the end, these apparent molecular weights with their respective stoichiometry can only
be taken as an overall estimate of the tme system.
4.4 Crystallization Trials
Sparse matrix screens (Jancarik et al., 1991) of C-O, LPD-14 and LPD- 16 as well as LPD-
20 solubilized BR were prepared using the vapour diffusion technique. These initial
sparse matrix screens served primarily as usefid starting points in terms of preparing
subsequent crystallization trials of both the LPDs alone and in association with BR.
Crystal screen 1 (Hampton Research) produced leads for LPD C-O as it produced
"fuzzballs" in 0.2 M calcium acetate hydrate, 0.1 M sodium cacodylate pH 6.5 and 18%
PEG 8000 and LPD-16 producing thin "egg-shell" crystals in 0.2 M ammonium acetate,
0.1 M trisodium citrate dihydrate, pH 5.6, 30% v/v a-methyl-2,4-pentanediol. However,
no success was achieved upon optimizing either of these two leads.
Figures 4.22 and 4.23 illustrate the spectnim of results obtained fkom the LPD-20
solubilized BR crystallization attempts. Microcrystals were obtained in some conditions
(Figure 4.22). Some of these crystals appear purple which suggests that they could be
BR-LPD cystals. However, other crystals were not intensely purple and were thought to
be either LPD, salt or denatured BR-LPD crystals. When working with detergents in
crystallization trials, a phenornenon cded phase separation often occurs as a result of the
polar interactions between the detergent molecules. As a result, a detergent-rich phase
and a detergent-depleted phase form within the crystallization drop (Michel, 1990).
Phase separation and precipitation of BR were comrnon observations with this sparse
matnx screen. In some instances, BR maùitained its native state within a detergent-rich
(Figure 4.23A) and a detergent-depleted phase (Figure 4.23C). In other cases, however,
BR denatured (Figure 4-23 B). Evidently , more crystallization conditions need to be
explored in order to identi& "leads" that c m be optimued.
Figure 4.22: LPD-20 solubi Ezed BR cry stallization trials in A) 1.5 M E thium sulfate, 0.1
M sodium HEPES pH 7.5 B) 0.1 M potassium fluoride, 6.13% PEG 4000 and C ) 20%
isopropanol, 0.1 M sodium acetate pH 4.6,0.2 M calcium chloride
Figure 4-23 : Phase separation examples of LPD-20 solubil k d BR A) 30% PEG 4000,O. 1
M sodium citrate pH 5.6, 0.2 M ammonium acetate B) 30% isopropanol, 0.1 M sodium
cacodylate pH 6.5, 0.2 M sodium citrate and C) 0.2 M ammonium sulfate, 0.1 M sodium
cacodyl ate pH 6.5,30% vlv PEG 8000
CHAPTER 5: DISCUSSION AND FUTURE WORK
AIthough a great amount of effort has k e n invested in creating novel approaches for
generating high-resolution membrane protein structures, no measurable success has k e n
achieved in generating these structures in an effective and efficient manner. This thesis,
therefore, set out to develop a novel class of detergents, iïpopeptide detergents (LPD), to
facilitate the crystallization of membrane proteins. LPDs are 25 residue a-helical
amphipathic peptides covalently coupled to fatty acids at either end. They were designed
to occupy less space and be a better mimic of the phospholipid bilayer than traditional
detergents used for crystallizing membrane proteins. in addition, upon solubilizing a
membrane protein by aligning its fatty acid moieties dong its hydrophobic domains, the
LPD was designed to provide rîgid polar surface areas available to make the critical
crystal contacts necessary to fonn well-ordered 3-dimensional crystals.
5.1 Characterization of LPDs
An LPD peptide series which varied the length of the fatty acyl moieties fiom C- 12 to C-
20 by 2 acyl units was synthesized and purified using solid phase synthesis and Cd RP-
HPLC, respectively. Its identity and evaluation of purity was confiïrmed using ESI-MS.
Obtaining a pure peptide following organic synthesis is of utmost importance for two
reasons. First, the characterization of the peptide in association with the IMP must be
attributed to the peptide itself and not organic impurities. Secondly, a pure peptide is
required when preparing crystallization triais of the LPD-IMP complex.
The a-helicai design of the peptide was confirmed using CD spectroscopy. Despite the
fact that the control peptide, C-O, was designed to be a-helical, it demonstrated
significantly less helical content than the lipopeptides. This suggests that the fatty acid
moieties induce a conformational change upon the peptide. Sedimentation equilibrium
ultracentrifugation analysis demonstrated that at very low concentrations, less than 0.2
mM, the fatty acyl ch- drive LPD monomers to assemble into their micellar form in
order to satisQ thennodynamic requirements. In fact, in the monodisperse LPD solutions,
the association constants were so large that the monomeric form was undetectable.
Furthermore, CD measurements for LPD concentrations as low as 20 pM demonstrated
that the secondary structure was independent of concentration. In contrast, the
sedimentation equiiibrium analysis indicated a monomeric system for the control peptide
C-O. Therefore, it is postulated that both hydrophobicity of the fatty acyl chains and the
consequent association into their miceff ar form induce the a-helical conformation of the
peptide (Liu et al., 1999). Consequentiy, it is thought that the LPD monomers would be
similar to C-O in their monomeric form such that the LPD monomers would also possess
more of a random coi1 secondary structure. This would dlow the fatty acid chains and
the peptide backbone to sample greater degrees of conformations. Furthermore, it is
thought that these LPD monomers may form a monolayer at the air-water interface in
order to remove the aliphatic chains from the bulk aqueous phase.
The hydrophobic content of the LPDs also played a role in terms of its stability such that
increasing the fatty acyl chah lengths beyond 16 reduced the solubility of the system.
However, the concentrations at which the LPDs remained soluble was impressive in
comparison to traditional detergents with shi lar or smaller acyl chah lengths. LPDs-12
to 16 were stable in water at concentrations far greater than 10 mM while LPDs-18 and
20 were also initially soluble at these concentrations but quickly becarne insoluble with
time. In the end, it was discovered that the LPD series &PD-12 to LPD-20) was well
behaved over time at concentrations of approximately 2 mM in water. In contrast, the
solubility of traditional detergents is greatly reduced to impractical levels for detergents
with hydrophobic moieties of Cl4 or greater. For instance, DDM with a hydrophobic
acyl chain length of 12 has a solubiIity b i t at concentrations of 0.4 mM or lower
(Anatrace 1999-2000 Catalog).
RP-HPLC demonstrated that an increase in the fatty acyl chain length increased the
peptide's hydrophobic content. SEC and sedimentation equilibrium ultracentrifugation
demonstrated that it increased the size of the micelle as well. The increase in micelle size
can be attributed to the fact that the extra hydrocarbons occupy space. In order to achieve
equai packing density within the core of the micelle, an increase in the volume of the
interior core of the micelle must occur. This increase in volume is accomodated by
increasing the number of monomers within the micelle. LPDs with fatty acyl chain
lengths up to 20 formed well-defined micelles of reasonably small volume. In contrast,
native phospholipids fiom biological membranes that contain acyl chains of 16 to 18
hydrocarbons in length cannot form micelles but rather form bilayers. As mentioned, the
geometry of the detergent monomer plays a significant role in the size and shape of the
micelle. Phospholipids bear two fatty acyl chains resulting in a monomer tbat assembles
into a bilayer. LPD monomers, on the other haad, have a more balanced hydrophilic to
hydrophobic ratio than phosphoiipid or traditional detergents do. LPDs have a large
hydrophilic domain that aligns alongside a slightly srnaller hydrophobic domain. LPD
monomers, therefore, possess a wedge shaped or eliiptical geometry.
In fact, modelling the LPDs in terms of their sedimentation equilibrium derived
aggregation nurnbers, 8 to 13, suggested that the micellar form has an oblate elliptical
shape because their axial ratios are approximately equal to 1 (Cantor et al., 1980). Oblate
ellipses are more spherical in nature than prolate ellipses which are comparatively more
cylindrical in shape. For C-O, on the other hand, sedimentation analysis indicated a
monomeric form of the peptide whereas SEC indicated a trimeric form of the peptide. It
is postulated that the C-O monomer forms a prolate elliptical shape. Consequently, since
rod-like macromolecules elute more quickiy from a SEC colurnn than spherically shaped
molecules, it is dmcult to accurately determine the molecular weight of non-spherical
macromolecular samples by SEC.
Likewise, it is also important to emphasize that the sedirnentation equilibrium analysis
determination of micellar size was not entirely accurate either. In this case, the partial
specific volumes of the systems were based on a calculated rather than an experimentally
determined value. In order for one to be assured of the accuracy of these molecular
weights, a more rigorous experimentai determination of the partial specific volumes of
the LPD senes must be performed. Experirnentally, the partial specific volumes can be
determined using a mechanical oscillator densimeter (Kratky et al., 1973) or using
different concentrations of D20/Ht0 solutions (Fless et al., 1997).
In terms of comparing LPDs to traditional detergents, several parameters of the LPDs
imply that they are superior to traditional detergents. First, the length of the fatty acyl
chains attached to the LPDs are in the same order of those native to a biological
membrane. Traditional detergent alkyl chab lengths rarely extend beyond 12 acyl units
due to solubility constraints. Second, the two fatty acyl chains of the LPDs align dong
the face of the amphipathic peptide thereby allowing these chains to align dong the
longitudinal axis of the membrane protein. Traditional detergents also align dong the
longitudinal axis of the IMP but because the chah lengths are too short, more detergent
monomers are required to align in the form of a sphere around the rest of the hydrophobie
domain. As a result, the dimensions of a LPD-protein complex are smaller than that of a
traditional protein-detergent complex. In fact, the aggregation number of the LPD
micelles range fiom 8 to 13 in contrast to traditional detergents which range fiom 10 to
130 (Hjelmeland, 1986).
Although the micelle size was determined using both gel filtration chromatography and
sedimentation equilibrium ultracentrifugation, this data only provided information
regarding molecular weight and not structure. To determine the structure of the micelles,
X-ray crystallography, electron microscopy or NMR techniques can be used.
Crystallization trials of the LPDs alone were prepared but failed to generate any
favourable crystals. Extensive efforts should be invested into obtaining well-dmcting
LPD crystals. This can be achieved by setting up crystallization trials in a factonai
manner varying LPD, salt a d o r precipitant concentrations, pH, and temperature.
The pH parameter was investigated using CD spectroscopy. This technique indicated
that the LPDs are essentially stable over the pH range of 3-10 but demonstrated a slight
decrease in stability over the neutral pH range. A more pronounced destabilization effect
at neutral pH was observed for the C-O peptide than the LPD series. This could be due to
the repulsion of like charges at the amino terminus of the peptide. At neutral pH, the fiee
6-amino group of the uncoupled ornithine is positively charged and hteracts unfavorably
with the positive helix dipole charge at the N-terminus of the peptide. The repulsion of
like charges destabilizes the helical conformation of the peptide. LPDs, however,
experience less instability at neutral pH because the &amino groups of the ornithines are
coupled to fatty acids. In this case, the instability could simply be a result of the
destabilizing helix dipole effects. For friture work, a cornparison of the LPDs with LPD-
2, a peptide with acetylated ornithines, would serve as a superior comparative control
rather than or in addition to C-O simply for this stabilization reason. Nevertheless, the
LPDs are pH stable and this is a desirable trait for sampling a wide range of
crystallization conditions. Fwthermore, this trait provides a broader application for the
LPDs in terms of their ability to solubilize other membrane proteins with different pH
optima.
5.2 LPD Solubilization of P hosp bolipid Bilayers
The extraction of membrane proteins by detergents in their native state from the
phospholipid bilayer is a critical step in membrane protein crystallization in the presence
of detergents (Knol et al., 1998). The traditional detergent DDM achieved solubilization
of the PC Liposomes in the ratio of 8 DDM to 1 phospholipid. Similarly, LPD-12, LPD-14
and LPD-16 demonstrated the ability to solubilize a phospholipid bilayer in a ratio of 10
LPDs to 1 phospholipid. The data obtained for the DDM solubilization of PC liposomes
was essentially consistent with the de Ia Maza investigation. However, the definitions of
complete solubilization in the two investigations were different. This investigation
sampted fewer concentrations and defined solubilization once the rnixed micelles
produced a consistent light scattering and hydrodynamic radius. In contrast, the de Ia
Maza investigation, sampled a wider range of concentrations and defined complete
solubilization when the total light scattenng was 10% of the original light scattered.
Consequently, the molar ratios obtained in this investigation appear artificially higher
than the de la Maza investigation. Nevertheless, the fact that LPDs can fully solubilize a
lipid bilayer makes them potential candidates to solubilize membrane proteins directly
from their biological membranes. However, with the current methoci of synthesis, this is
an unredistic approach due to the unreasonable amounts of LPD that the extraction
would consume.
Solubilization of a lipid bilayer depends on the cmc of the detergent as well as the
hydrophilic-lipophilic balance of the detergent (de la Maza et al, 1997). DDM is more
effective at soiubilizing a phospholipid bilayer than OG because its hydrophobic to
hydrophilic moieties are more balanced. In addition, its longer hydrophobic tail is
correlated to an increased propensity of the detergent to adsorb to the outer leaflet of the
liposome and then subsequently incorporate into the lipid bilayer (de la Maza et al.,
1997). Other factors such as the charge and size of the hydrophilic head of the detergent
as well as the charge and composition of the phospholipid bilayer play roles in the
efficiency of detergents or a-helical peptides to penetrate a phospholipid bilayer (Dathe
et al., 1999). In our case, the longer tailed detergents, LPD-18 and LPD-20, demonstrated
a trend contrary to expected in that they f d e d to solubilize the PC liposomes at similar
concentrations to the shorter chain length LPDs. However, this codd be explained by the
fact that the shorter chah length LPDs are insufncient to partition effectively into the
liposome bilayer. Instead, they disperse the phospholipid bilayer causing an eventual
phase transition of the bilayer into mixed detergent - lipid micelles-
De la Maza investigated the permeability of the liposomes upon addition of detergent by
monitoring the release of the fluorescent dye, S(6)-carboxyfluorescein (CF), fiom the
interior of the liposomes (de la Maza et al., 1997). To understand our unexpected
fmdings, a similm experiment could be performed with our LPD series to investigate if
LPD-18 and LPD-20 embed into the liposome without disrupting the liposome's
permeability. Monitoring the permeability of the liposomes in the presence of the smaller
chain length LPDs would indicate the concentration of LPD necessary to produce these
changes. One of two situations rnay be plausible to explain our findings. Fust, LPD-18
and LPD-20 could have embedded into the liposome altering its permeability but because
insufficient LPD concentrations were applied, the liposomes failed to disperse. Second,
the permeability of the liposomes was unaltered due to the similarity of the lipid chain
lengths and LPD-18 and LPD-20 simply incorporated into the liposome producing a
iarger mixed LPD-PC liposome. For a more intensive approach to understand these
results, cryotransmission electron microscopy couid be used to visualize the structural
transitions of the liposome upon titration with LPDs (Knol et al., 1998).
5.3 Bacteriorhodopsin Stability and Crystallization in Association with LPDs
First, our data cleariy demonstrated that in the absence of detergent or in the presence of
the control peptide C-O, BR was not soluble and precipitated out of solution. This
confirms that detergents are necessary to maintain the solubility of BR. Second, LPD-12
through to LPD-20 were capable of initially solubilizing BR to the same extent as the
traditional detergent OG. Since C-O failed to solubilize BR but LPDs could, this
confums that the fatty acyl chains of the LPDs are the critical components of the peptide
which facilitate the solubilization of the membrane protein. Over time, however, the
LPDs proved to be far superior to OG in maintaining the stability of the solubilized BR.
The increased stability of the LPD-BR cornplex compared to the OG-BR complex can be
attributed to the size and shape of the complexes as well as the packing constraints of the
fatty acyl chains dong the hydrophobic domain of BR. First, the fatty acyl chah lengths
of the LPDs range f?om 12 to 20 whereas OG has only 8 acyl units which is insufncient
to span the length of the hydrophobic domain. Consequently, there is a mismatch
between the hydrophobic surface provided by OG compared to that the IMP itself. As a
result, a "ring" of OG molecules must surround this hydrophobic domain of the IMP to
maintain the solubility of the protein. As predicted, IMPs are more stable in
environments which better mimic the native phospholipid bilayer. Secondly, the fatty
acyl chains on the LPDs have less fieedom of motion within a micelle compared to
traditional detergents because of the structural hindrances imparted on these fatty acyl
chains by the peptide they are attached to. Traditional detergents, on the other hand, are
more flexible and can sample a wider range of conformations to achieve the appropriate
packing density within the hydrophobic core to satisQ thermodynamic requirements.
LPDs with fatty acyl chah lengths of 16 or smaller were most effective at maintainhg
the solubility and stability of native BR over time. LPDs with chah lengths longer than
16 were capable of initially solubilizing the BR to a similar extent as the shorter chain
length LPDs, but over tirne, the solubilized BR aggregated slightly in the presence of the
longer chab iength LPDs. This could be attributed to the packing of the fatty acyl chains
within the hydrophobic core. In the case of the shorter chah lengths, the chains are short
enough to not overlap each other dong the peptide. However, with the longer chains, the
c h a h could overlap causing the chains to sample different conformations to achieve
equal packing density as the shorter chain LPDs. This could result in the instability of
the LPD-BR complex because of the way the fatty acyl chains align along the
transmembrane domain of BR.
In fact, a preliminary mode1 of how LPD solubilize a BR trimer was proposed based on
the sedimentation equilibrium analysis of the LPD-12 and LPD-20 solubilized BR
sy stems. Again, this mode1 which proposed approximately 25 LPD molecules aligning
along the longitudinal axis of the BR trimeric transmembrane domains is purely
preliminary especially siuce the partial specific volume was a highly inaccurate estimate.
As in the case of micelle size determination, the partial special volume of the associated
LPD solubilized BR system must be experimentally determined to more accurately
propose a mode1 for the LPD-solubilized BR trimeric complex.
Furthemore, the LPD-12 and LPD-20 solubilized BR systems were not perfectly
monodisperse when monitored by sedimentation equilibrium analysis. The heterogeneity
of these systems codd be partially attributed to excess LPD micelies present in the
solution as well as to the method of detergent exchange. Perhaps using such a quick
method of exchange did not facilitate the production of a well defined protein-detergent
complex Removing the OG so quickly without allowing the LPD to slowly equilibrate
into the system could have produced some slight BR aggregation. Perhaps a more gentle,
equilibrium based method of detergent exchange such as dialysis would facilitate the
exchange of detergents to produce a more hornogeneous system.
Obtaining stnictural data of these complex systems would be invaluable. Information
regarding the oligomeric state of BR as well as the number of LPD molecules
surroundhg BR could be deterrnined. A monodisperse LPD-protein system is required
when setting up crystallization trials. Nevertheless, despite the lack of monodispersity of
the LPD-solubilized BR samples, a sparse matrix crystallization screen was prepared for
the LPD-20 solubilized BR system. A variety of results were obtained ranging fiom
precipitation to phase separation to mini-crystals. Therefore, M e r crystallization trials
rnust be sampled to identiQ the appropriate parameters such as pH, temperature and
concentration of protein, salt anaor precipiîant that yield well-ordered membrane protein
crystals. In addition, the method of crystallization could be investigated. Although the
"hanging-drop" vapour diffusion technique seems to be the most convenient method for
preparing sparse matrix screens, other techniques such as microdialysis, batch method or
"sitting-drop" vapour diffusion could be explored (Michel, 1990). In fact, microdialysis
is a desirable approach for crystallizing BR. It can perfonn two functions
simultaneously. First, a dialysis membrane can be used which is large enough for the
diffusion of OG molecules but not the LPD micelles; this would facilitate the detergent
exchange. Second, the dialysate conditions can be easily monitored and altered to
facilitate the crystaliization of the BR-LPD complex.
5.4 Future Work
Although this designed LPD achieved success in providing superior stability of BR than
the traditional detergent OG, variations to the design could potentially improve the LPDYs
ability to achieve solubility or crystallization. Extending or shortenhg the length of the
peptide or changing the diphatic moiety attached could accommodate different sized and
shaped proteins such as monotopic a-helical membrane proteins or bstrand containing
membrane proteins. For example, the peptide could range fiom 20 to 30 residues and
still satisQ the requirement of spanning the width of the average membrane bilayer.
Furthemore, by changing the aiiphatic moiety to a branched or cyclical structure would
change the size and shape of the micelle; this could also facilitate crystallization of
different sized and shaped membrane proteins.
Synthesizing peptides using the solid phase synthesis method is an expensive procedure
that produces low yields upon purification. Therefore, in the interest of cost, the peptide
sequence could be modified to accomodate the production of the peptide in vivo (ie.
recombinant Escherichia coli). The ornithines, the sites of the covalent coupling of the
aliphatic moiety, couid be replaced with cysteine, for instance, and could be coupled by a
thio-ester bond under the appropriate conditions to the aliphatic moieties following
purification of the peptide fiom the recombinant cells. In addition, it would be necessary
to create a fusion between this peptide and a purification tag to prevent degradation by
proteases in vivo as well as simplifjr the purification procedure. The purification tag
should be easily cleaved and removed to yield pure peptide. This recombinant
production of the peptide may greatly reduce the cost of synthesis and may produce much
higher yields.
One interesthg physicd characteristic of LPD that was not determined in this thesis was
its cmc. In fact, sedimentation equilibrium ultracentrifugation demonstrated no
monomenc species to be present at concentrations as low as 0.2 mM, thus indicating that
the critical micelle concentrations of the LPD senes were very low. A simple method of
determining this would be to utilize the fluorescent probe, anilinosulfonic acid (ANS).
This probe has the property to produce greater fluorescence in a hydrophobic
environment than in a polar environment. Upon reaching its cmc, detergent monomers
assemble into a micelle creating a hydrophobic core. Therefore, if ANS is included in the
detergent solution, upon reaching the cmc of the detergent, a sudden increase in
fluorescence would be observed (Walter et al., 1990).
Finally, extensive solubility and crystallization investigations of the current LPDs in
combination with other membrane proteins should be pursued. A broad range of types
and sizes of integral membrane proteins could be explored including other poiytopic a-
helical transmembrane proteins, monotopic proteins or diierent sized p-barre1 integral
membrane proteins. In fact, once a mode1 membrane protein is solved in association with
the LPDs, an integral membrane protein of unknown structure could be attempted.
On a fmal note, although this thesis has focused on the role of LPDs as detergents for
solubilizhg membrane proteins for crystailization, other applications for the role of LPDs
could be investigated. One application, in particular, is the potentiai for LPDs to serve as
cytolytic agents. a-helical peptides have been shown to be hemolytic and bacteriolytic
agents (Shai, 1999, Dathe et ai., 1999). In fact, great efforts have been placed into
expandhg the large group of antibiotic peptides which fold into an amphiphatic a-helical
conformation upon binding to and inserting into the phospholipid bilayer of target cells
(Dathe et al., 1999). This insertion of the peptide into the membrane disrupts the normal
functioning of the cell by breaking down the transmembrane potential and creating a
le* membrane resulting in death of the cell. It wodd be hopefül that the LPDs would
be superior antimicrobial agents, providing increased activity and microbial selectivity
than the existing peptides.
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