Julian Langer Dissertation
I
Dissertation
submitted to the
Combined Faculties for the Natural Sciences and for Mathematics
of the Ruperto-Carola University of Heidelberg, Germany
for the degree of
Doctor of Natural Sciences
presented by
Julian David Langer, Diplom-Chemiker
born in Heidelberg
Oral examination:
.
II
Conformational dynamics of coatomer:
functional and structural studies.
Referees:
Prof. Dr. Felix Wieland
Prof. Dr. Irmgard Sinning
I
Table of contents
Table of contents.......................................................................................................................I
Abstract (english).....................................................................................................................IV
Abstract (german).....................................................................................................................V
Abbreviations...........................................................................................................................VI
List of figures..........................................................................................................................VII
List of tables............................................................................................................................IX
1.
1.1.
Introduction.................................................................................................................... 1
The Secretory Pathway.............................................................................................. 1
1.2.
Vesicular Transport: The three coating systems ........................................................ 5
1.2.1.
Clathrin-coated vesicles ..................................................................................... 5
1.2.1.1.
Structural studies on clathrin-coated vesicles ............................................. 6
1.2.1.2.
Conformational changes in adaptor proteins............................................... 9
1.2.1.2.1. The adaptor protein 2 (AP-2) system....................................................... 9
1.2.1.2.2. The adaptor protein 1 (AP-1) system......................................................11
1.2.1.2.3. Golgi-localizing, γ-adaptin ear homology domain, Arf-binding proteins ...12
1.2.2.
COPII-coated vesicles.......................................................................................12
1.2.3.
COPI-coated vesicles........................................................................................14
1.2.3.1.
The COPI budding process .......................................................................15
1.2.3.2.
Summary: Comparison of COPI with the other vesiculation systems.........20
1.3.
2.
Aims of present work ................................................................................................23
Results .........................................................................................................................24
2.1.
Conformational dynamics of coatomer ......................................................................24
2.1.1.
A conformational change in γ-COP: Screening p24-family members.................24
2.1.2.
Limited proteolysis and screening of other coatomer subunits...........................26
2.1.3.
Labeling of coatomer with fluorescent dyes.......................................................27
2.1.4.
Labeling with amine-reactive dyes ....................................................................28
2.1.5.
Functionality of labeled coatomer......................................................................31
2.1.6.
Specific immobilization of labeled coatomer ......................................................34
2.1.7.
Surfaces with Cy3-Cy5-labeled coatomer..........................................................39
2.1.8.
Inter- or intramolecular FRET? ..........................................................................40
2.1.9.
Assessing the number of attached dyes............................................................41
2.1.10. Approximating the FRET efficiency Eapp. ...........................................................42
2.1.11. Interaction of coatomer with cytoplasmic tail domains of ligand proteins ...........43
2.1.12. ELISA-like binding assay...................................................................................46
2.1.13. Rare events.......................................................................................................48
2.2.
Electron Microscopic investigation of COPI vesicles .................................................49
2.2.1.
Preparation of COPI vesicles in vitro .................................................................49
2.2.2.
Embedding of chemically fixed vesicles into a matrix ........................................52
2.2.3.
Cryo-electron microscopy of COPI vesicles generated in vitro ..........................53
2.2.4.
Using "backed"-Quantifoil..................................................................................55
2.2.5.
Direct preparation and imaging of COPI vesicles. .............................................57
II
3.
Discussion ....................................................................................................................64
3.1.
Conformational dynamics of coatomer ......................................................................64
3.1.1.
Data presented in this work...............................................................................64
3.1.2.
Membrane protein capture and coat lattice formation........................................66
3.2.
4.
Electron microscopy and tomography of COPI vesicles ............................................69
Materials and Methods .................................................................................................70
4.1.
Materials ...................................................................................................................70
4.1.1.
Reagents...........................................................................................................70
4.1.2.
Peptides............................................................................................................70
4.1.3.
Beads................................................................................................................71
4.1.4.
Molecular weight standards for SDS – PAGE....................................................71
4.1.5.
Protease – Inhibitors .........................................................................................72
4.1.6.
Antibodies .........................................................................................................72
4.1.6.1.
Primary antibodies.....................................................................................72
4.1.6.2.
Secondary antibodies ................................................................................73
4.1.7.
Activated fluorophores.......................................................................................73
4.2.
Equipment ................................................................................................................74
4.2.1.
FPLC-Anlagen ..................................................................................................74
4.2.2.
SMART .............................................................................................................74
4.2.3.
Spectrophotometer............................................................................................74
4.2.4.
Single molecule-sensitive confocal setup ..........................................................74
4.2.5.
Electron microscopes........................................................................................77
4.3.
Methods....................................................................................................................77
4.3.1.
SDS – PAGE.....................................................................................................77
4.3.1.1.
Stock solutions for SDS - PAGE ................................................................77
4.3.1.2.
Separation gels .........................................................................................78
4.3.1.3.
Stacking gels .............................................................................................79
4.3.2.
Sample preparations .........................................................................................79
4.3.2.1.
Sample preparation for SDS – PAGE ........................................................79
4.3.2.2.
Tri-Chloro-acetic acid (TCA) – precipitation ...............................................79
4.3.2.3.
Immunoprecipitation (IP) ...........................................................................80
4.3.3.
Staining of proteins in SDS – gels .....................................................................80
4.3.3.1.
Coomassie-Staining ..................................................................................80
4.3.3.2.
Silver stain.................................................................................................81
4.3.4.
Western blot analysis ........................................................................................81
4.3.4.1.
Transfer of proteins separated by SDS-PAGE onto PVDF-membranes.....81
4.3.4.2.
Ponceau-Staining of proteins on PVDF-membranes..................................82
4.3.4.3.
Immunochemical detection of proteins on PVDF-membranes ...................82
4.3.5.
Bradford assay..................................................................................................83
4.3.6.
Isolation of coatomer from rabbit liver cytosol....................................................83
4.3.6.1.
Isolation of rabbit liver cytosol....................................................................83
4.3.6.2.
Ammoniumsulfate precipitation of rabbit liver cytosol.................................83
4.3.6.3.
DEAE anion exchange chromatography of ASP ........................................84
4.3.6.4.
SourceQ anionic exchange chromatography of the DEAE pool .................84
4.3.6.5.
Concentration of the SourceQ-Pools .........................................................84
4.3.6.6.
Buffers for coatomer preparation ...............................................................85
4.3.7.
Labeling coatomer.............................................................................................86
4.3.8.
Analysis of labeled coatomer subunits ..............................................................86
4.3.9.
Isolation of rat liver Golgi...................................................................................87
4.3.10. Pull down experiments ......................................................................................87
III
4.3.11. Membrane binding assay ..................................................................................88
4.3.12. In vitro COPI vesicle budding assay ..................................................................88
4.3.12.1.
17h gradient purification ............................................................................88
4.3.12.2.
1h cushion centrifugation...........................................................................89
4.3.12.3.
Sucrose-free preparation of crude vesicles................................................90
4.3.13. Grid preparation for negative staining................................................................90
4.3.14. Grid preparation for cryo electron tomography ..................................................90
4.3.15. Precipitation assay ............................................................................................91
4.3.16. Limited proteolysis ............................................................................................91
4.3.17. Antibody purification..........................................................................................91
4.3.18. Surface preparation...........................................................................................92
4.3.19. Calculation of FRET efficiency Eappr...................................................................92
4.3.20. ELISA-like binding assay...................................................................................94
4.3.21. Electron microscopy and tomography ...............................................................95
4.3.21.1.
Aquisition of tilt series................................................................................95
4.3.21.2.
Tomographic reconstruction ......................................................................95
4.3.21.3.
Segmentation of tomograms......................................................................96
5.
References ...................................................................................................................97
Acknowledgments.................................................................................................................113
IV
Abstract
In my PhD thesis I have investigated molecular mechanisms in the biogenesis of membrane
vesicles.
Formation of transport vesicles involves polymerization of cytoplasmic coat proteins. In COPI
vesicle biogenesis, the heptameric complex coatomer is recruited to donor membranes by
the interaction of multiple coatomer subunits with the budding machinery. Specific binding to
the trunk domain of coatomer's subunit γ-COP of the Golgi membrane protein p23 induces a
conformational change in the γ-subunit, leading to polymerization of the complex in vitro.
Using a combination of biochemical assays and an assay based on single-molecule, singlepair fluorescence resonance energy transfer, we find that this conformational change is only
induced by dimers of the p24-family proteins p23 and p24, and neither by the other p24family members nor by cargo proteins. This conformational change takes place in individual
coatomer complexes, independent of each other, and the rearrangement induced in γ-COP is
transmitted within the complex to its α-subunit. α-COP is one of coatomer's subunits capable
of binding to dibasic cargo motifs, and also shows analogy to the Clathrin molecule. We
propose a model in which capture of membrane protein machinery triggers cage formation in
the COPI system.
At the nanometer resolution I started investigating the structure of the lattice of
conformationally changed coatomer on COPI vesicles generated in vitro from purified Golgi
membranes and coating machinery, using cryo electron tomography. Initial data on coated
vesicles and coated buds is presented.
V
Zusammenfassung
In meiner Doktorarbeit habe ich die molekularen Mechanismen der Biogenese von COPIVesikeln untersucht.
Der Transport von Proteinen und Membranen in einer eukaryotischen Zelle erfolgt über
vesikuläre Träger. Zur Bildung dieser Vesikel polymerisieren Hüllproteine, die sowohl in einer
löslichen, cytosolischen Form als auch in einer membrangebundenen Form vorliegen. In der
Biogenese eines COPI Vesikels bindet der heptamere Hüllkomplex Coatomer an die
Donormemrbanen über multiple Interaktionen mit der Maschinerie zur Abknospung der
Vesikel. Die Interaktion der γ-Untereinheit von Coatomer (γ-COP) mit dem transmembranProtein p23 induziert einen Konformationswechsel in γ-COP, der zur Polymerisation des
Komplexes in vitro führt.
In dieser Arbeit wurden biochemische und biophysikalische Methoden verwendet, um diesen
Konformationswechsel in Coatomer zu untersuchen. Dabei wurde ein Verfahren zur
Untersuchung der Konformation individueller Coatomer-Komplexe mit EinzelmolekülFluoreszenz-Resonanz-Energie-Transfer etabliert. Der beschriebene Konformationswechsel
in γ-COP wird nur durch dimere der Proteine p23 und p24 induziert, und nicht durch andere
Mitglieder der p24-Familie oder Frachtproteine. Er findet in einzelnen Coatomer-Komplexen
statt, und wird in die periphere Untereinheit α-COP weitergeleitet. α-COP ist eine der beiden
Untereinheiten von Coatomer, die Frachtmoleküle mit dibasischen Signalsequenzen binden;
zudem zeigt α-COP Analogien zu Clathrin über α-solenoide und β-Propeller-Domänen. In
dieser Arbeit wird ein Modell vorgeschlagen, in dem die Bindung von spezifischen
Transmembranproteinen die Polymerisierung der Hüllproteine und die Ausbildung der COPIVesikelhülle induziert.
In einem zweiten Projekt wurde die Struktur von Coatomer in der COPI-Hülle auf in vitro
generierten COPI Vesikeln durch cryo-Elektronenmikroskopie und Tomographie untersucht.
Erste
tomographische
Rekonstruktionen
Donormembranen werden gezeigt.
von
Vesikeln
und
COPI-Knospen
an
VI
Abbreviations
AA (aa)
AP
APD
APS
Arf
ATP
BFA
BSA
COP
DMMA
DMSO
DNA
DTT
EApp
EDTA
ER
ERGIC
FRET
GAP
GDP
GEF
GST
GTP
h
Hepes
HPLC
HRP
IgG
IMAC
IP
IPTG
KD
kDa
kHz
MALDI
min
NP-40
nt
OD
PBS
PBS-T
PCR
PMSF
PVDF
rpm
s
SDS-PAGE
SNARE
TCA
TEMED
TGN
TMB
wt
Amino Acids
Adaptor protein
Avalanche photo diode
Ammonium persulfate
ADP-ribosylation factor
Adenosin tri-phosphate
Brefeldin A
Bovine serum albumin
Coat protein
Dimethylmaleic acid anhydride
Dimethyl sulfoxide
Desoxyribonucleic acid
Dithiothreitol
Approximated energy transfer efficiency
Ethylendiaminetetra-acetic acid
Endoplasmic reticulum
ER-Golgi intermediate compartment
Fluorescence Resonance Energy Transfer
GTPase activating protein
Guanosine diphosphate
Guanosine nucleotide exchange factor
Glutathione-S-transferase
Guanosine tri-phosphate
hour
4-(2-hydroxyethyl)-1-piperazin-ethansulfonic acid
High pressure liquid chromatography
Horse radish peroxidase
Immunoglobulin class G
Immobilized metal affinity chromatography
Immunoprecipitation
Isopropyl-1-thio-β-D-galactopyranoside
Dissociation constant
kilo-Dalton
kilo-Hertz
Matrix assisted laser desorption/ionization
minute
Nonidet® P40 (Nonylphenylpolyethylene glycol)
Nucleotides
Optical density
Phosphate buffer saline
Phosphate buffer saline + Tween 20
Polymerase chain reaction
Phenylmethulsulfonyl fluoride
Polyvinyldifluoride
Revolutions per minute
second
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
Soluble N-ethylmaleimide sensitive factor attachment protein receptor
Trichloroacetic acid
N, N, N’, N’-Tetramethylethylenediamine
Trans Golgi network
Tetramethylbenzidin
wild type
VII
List of figures
Figure 1:
Schematic representation of the Secretory Pathway.
Figure 2:
Scheme of the Clathrin triskelion and designs of Clathrin lattices.
Figure 3:
Structural similarities of coat proteins.
Figure 4:
Scheme of the COPI budding process.
Figure 5:
Scheme of p24-family members structure and sequences of cytoplasmic
domains.
Figure 6:
Precipitation of coatomer by p24-family proteins.
Figure 7:
Limited proteolysis of coatomer.
Figure 8:
Analysis of coatomer after labeling with different amounts of Cy3-NHS esters.
Figure 9:
Analysis of coatomer after labeling with NHS-esters of Cy3 and Cy5.
Figure 10:
Functionality of labeled coatomer: Binding to the cytoplasmic domains of p23
and OST48.
Figure 11:
Functionality of labeled coatomer: Membrane binding assay.
Figure 12:
Functionality of labeled coatomer: In vitro formation of COP vesicles.
Figure 13:
Immobilisation strategies to specifically tether labeled coatomer to glass
surfaces.
Figure 14:
Low intensity surface scans of glass surfaces coated with a BSA-antibody
solution.
Figure 15:
Low intensity surface scans of glass surfaces coated with a BSA-antibody
(CM1) solution and incubated with Cy3-Cy5-labeled coatomer.
Figure 16:
Low intensity surface scans (20µm x 20µm) of glass surfaces coated with a
BSA-antibody solution and incubated with a 1:1 mixture of Cy3-labeled
coatomer and Cy5-labeled coatomer.
Figure 17:
Single-pair FRET in coatomer.
Figure 18:
Normalized histogram for Cy3-Cy5-labeled coatomer, no peptide added.
Figure 19:
Normalized histogram for Cy3-Cy5-labeled coatomer, after addition of OST48.
Figure 20:
Normalized histogram for Cy3-Cy5-labeled coatomer, after addition of p23d.
Figure 21:
ELISA binding curves to monitor the KD and the total number of available
binding sites of coatomer for cargo proteins in the presence and absence of
dimeric p23.
Figure 22:
Fluorescence intensity trace of a Cy3-Cy5-labeled coatomer that oscillates
between two FRET states.
Figure 23:
Images of GTPγS-COPI vesicles, purified by sucrose gradient and submitted
to negative stain.
VIII
Figure 24:
Images of GTPγS-COPI vesicles purified either via a 17h-gradient (A-C) or
centrifugation on a sucrose cushion).
Figure 25:
Images of COPI vesicles (2.5mg of Golgi) prepared with GTPγS, purified by
centrifugation on a sucrose cushion, and submitted to negative stain.
Figure 26:
Images of COPI vesicles prepared with GTP in vitro, embedded in a polymer
matrix and stained with Ruthenium Red and OsO4.
Figure 27:
Images of COPI vesicles (2.5mg of Golgi) prepared with GTPγS, purified by
centrifugation on a sucrose cushion, and submitted cryo electron microscopy.
Figure 28:
Low (A) and high (B) magnification Images of COPI vesicles, deposited on a
carbon-backed Quantifoil grid, embedded in vitreous ice.
Figure 29:
Low (A) and high magnification (B) Images of COPI vesicles with 6nm gold
dots deposited on carbon-backed Quantifoil, and embedded in vitroues ice.
Figure 30:
Section of a tomogram of a sample of purified GTPγS-COPI vesicles
deposited on a thin carbon film, embedded in vitreous ice.
Figure 31:
TEM image of GTP-COPI vesicles embedded in vitreous ice.
Figure 32:
Section of a tomogram recorded of COPI samples containing COPI vesicles
directly applied to a Quantifoil grid after preparation.
Figure 33:
Sections of a tomogram recorded of COPI samples containing COPI vesicles
directly applied to a Quantifoil grid after preparation.
Figure 34:
Rendering of coated vesicle boxed in Figure 33 using the Amira software
package.
Figure 35:
Sections of a tomogram of a sample directly applied to a Quantifoil grid after
preparation, containing a coated bud.
Figure 36
Rendering of the coated bud boxed in Figure 35 using the Amira software
package.
Figure 37:
Model for putative role of the conformational change in α-COP during COPI
vesicle formation.
Figure 38:
Structure of NHS-ester-derivative of the fluorophores
Figure 39:
Scheme of the custom-built confocal setup used in this study.
Figure 40:
Properties of filters and dichroic mirrors employed in the single-molecule
sensitive setup.
Figure 41:
Fluorescence intensity trace of a typical singly donor- and acceptor-labeled
coatomer complex, and scheme of data extraction.
Figure 42:
Schematics of steps involved in calculation of EApp.
IX
List of tables
Table 1:
Coatomer subunits: Size, motifs and interaction partners.
Table 2:
Sequences of peptides used in this study.
Table 3:
Antibodies used in this study.
Table 4:
Gel compositions used for the Protean III system.
Table 5:
Buffers used for the preparation of coatomer from rabbit liver cytosol.
X
Declaration of primary authorship
I hereby declare that I have completed this thesis independently and without any assistance
from third parties. Furthermore, I confirm that no sources have been used in the preparation
of this manuscript other than those indicated in the thesis itself.
Heidelberg, 28. 07. 2008
1. Introduction
1.
1
Introduction
Eukaryotic cells are highly specialized and compartimentalized systems, generating different
intracellular micro-environments. By their individual protein and lipid composition, each
compartment allows for specific, characteristic reactions (i.e. disulfide bond formation,
glycosylation or protein degradation). These compartments ("organelles") have to
continuously exchange proteins and lipids to maintain the specific composition required for
the individual reactions, in order to both replenish lipids, proteins and substrates required for
the reaction, and to transport products to their destination within or outside the cell. The bulk
of this cargo is transported via small vesicles with a diameter of 60-100nm.
1.1.
The Secretory Pathway
In Eukaryotic cells, the majority of the proteins synthesized in the Endoplasmic Reticulum is
transported to their respective destinations via an intracellular route comprising several
organelles (Figure 1). This pathway, termed the Secretory Pathway, was described first in
exocrine pancreatic cells (Caro and Palade, 1964).
Figure 1: Schematic representation of the Secretory Pathway. Some of the pathways
indicated are still under debate.
Unlike cytosolic proteins or proteins targeted to compartments like the nucleus or
mitochondria, proteins transported along the Secretory Pathway bear distinct signal
sequences, targeting them initially to the ER (Blobel and Dobberstein, 1975a; Blobel and
1. Introduction
2
Dobberstein, 1975b). They are comprised of secreted proteins, lysosomal/vacuolar proteins
and membrane proteins. During their transition through the Secretory Pathway leading to
their respective destinations, these proteins also undergo post-translational modifications
(i.e. disulfide bond formation (Freedman, 1989), N-glycosylation (Kornfeld and Kornfeld,
1985), prolin-hydroxylation and attachment of GPI-anchors (Caras and Weddell, 1989)).
Proteins that travel along the Secretory Pathway begin their journey in the ER. As soon as
the nascent protein chain bearing the signal sequence leaves the ribosome, further
translation is arrested (Walter and Blobel, 1981a; Walter and Blobel, 1981b; Walter et al.,
1981). The signal sequence then binds to the signal recognition particle (SRP), and this
complex, in turn, binds to the ER-bound SRP receptor (Ganoth et al., 1988; Gilmore et al.,
1982a; Gilmore et al., 1982b; Walter and Blobel, 1980). Here, translocation through the ER
membrane is mediated by the "translocon", a trimeric protein complex comprising the
proteins Sec61α/β/γ (Deshaies et al., 1991; Deshaies and Schekman, 1989).
In the ER, two independent housekeeping systems exist that target un- and misfolded
proteins (Friedlander et al., 2000; Travers et al., 2000): The unfolded-protein-responsesystem (UPR) and the ER-associated-degradation-system (ERAD). Proteins identified as unor misfolded (for example by free thiol groups or incorrect N-glycosylations) are retranslocated into the cytoplasm (Plemper and Wolf, 1999), polyubiquitinated (Hershko et al.,
1979) and subjected to proteasome-mediated proteolysis (Haass and Kloetzel, 1989).
Proteins that have successfully folded and have passed ER quality control are exported from
the ER by COPII vesicles (Barlowe et al., 1994). The COPII coat is comprised of the small
GTPase Sar1p, that governs the coating process by its nucleotide state, and the coating
protein complexes Sec23/24 and Sec13/31 (Salama et al., 1993), that sequentially bind to
Sar1p once it has been activated by GDP-to-GTP exchange on the donor membrane to form
the COPII coat (see also: 1.2.2. COPII-coated vesicles).
Two classes of proteins transported via COPII vesicles have been characterized: First,
membrane proteins with exposed, cytosolic domains that typically bear diacidic (DxE) or
diaromatic signal sequences have been identified (Barlowe, 2003). Both motifs bind to one
member of the cytosolic coating protein complex Sec24 on two distinct sites (Miller et al.,
2003; Mossessova et al., 2003). The second class of cargo proteins is comprised of soluble
cargo proteins and GPI-anchored proteins that cannot access the cytosolic coating
machinery. Two different models describing anterograde transport are currently discussed:
The bulk-flow model and the receptor-induced-transport model.
In the bulk-flow model, no signal sequences are required for uptake of these proteins into
COPII vesicles, and all soluble proteins are transported from the ER to the Golgi. Proteins
1. Introduction
3
destined for the ER bear distinct ER-retrieval signals that allow them to recycle to the ER via
COPI-mediated transport (Karrenbauer et al., 1990; Wieland et al., 1987). This model was
supported by studies analysing the secretion rate of glycosylated acyltripeptides (Wieland et
al., 1987) and that of amylase and chymotrypsinogen by quantitative EM (MartinezMenarguez et al., 1999).
The receptor-induced-transport model postulates binding of soluble, lumenal cargo proteins
to transmembrane receptors as a key step in ER-to-Golgi transport. These transmembrane
receptors would, in turn, via their cytoplasmic domains, bind to the COPII machinery and sort
cargo into the vesicles. Supporting this hypothesis, deletion of the yeast membrane protein
Erv29 resulted in reduced uptake of the soluble pheromone α-factor into COPII vesicles
(Malkus et al., 2002). However, direct interaction of soluble cargo with a transmembrane
receptor has not been proven to date.
On their route to the Golgi, proteins exiting the ER pass through the ERGIC (ER-Golgi
Intermediate Compartment) that is defined by the presence of the protein ERGIC-53 (Hauri
et al., 2000). It is yet unclear if the ERGIC is formed by homotypic fusion of COPII vesicles or
if it is a distinct compartment that binds COPII vesicles by heterotypic fusion (Bannykh and
Balch, 1998; Bannykh et al., 1998). The ERGIC presumably acts as an interface between
COPII- and COPI-mediated transport, as it can be decorated with markers specific for both
COPII and COPI (Scales et al., 1997). These markers localize to distinct regions: Those that
can be decorated with markers for proteins cycling between the ER and the Golgi and for
COPI, and regions that can be decorated with markers for secreted proteins. Thus, the
ERGIC has been put forward as the location where the first sorting steps take place
(Martinez-Menarguez et al., 1999). This hypothesis was further strengthened by in vivo
studies showing a differential behaviour of ERGIC-53 and anterograde cargo (Ben-Tekaya et
al., 2005). Transport from the ERGIC to the Golgi is COPI-independent, as structures with
anterograde cargo leaving the ERGIC are both negative for COPI (Martinez-Menarguez et
al., 1999) and are significantly larger than typical COPI vesicles (Presley et al., 1997; Scales
et al., 1997).
The next step for proteins in the Secretory Pathway consists of intra-Golgi transport from the
cis-side to the trans-side. In eukaryotic cells, a Golgi stack consists of four to six cisternae,
that differ in both protein and lipid composition, and form a gradient from the cis-side (the
entry point from the ERGIC), the cis-Golgi-network (Rambourg and Clermont, 1990) to the
trans side, the trans-Golgi-network (Griffiths and Simons, 1986).
Transport within the Golgi is mediated by COPI vesicles. The coating process is, again,
governed by a small GTPase, Arf1. The proteinous coat of COPI vesicles is formed by the
heptameric protein complex coatomer that is recruited to the donor membrane en bloc (see
1. Introduction
4
also: 1.2.3. COPI-coated vesicles, (Hara-Kuge et al., 1994)). COPI vesicles mediate
transport in both the anterograde and in the retrograde direction within the Golgi, and take up
proteins bearing specific ER-retrieval signals (Cosson and Letourneur, 1994; Letourneur et
al., 1994), and transfer them back to the ER. Thus, in contrast to COPII vesicles, that transfer
cargo in an anterograde direction only, COPI vesicles mediate bidirectional transport (Orci et
al., 1997; Pelham and Rothman, 2000).
Two of coatomer's subunits, γ- and ζ-COP, were found to be isotypic (Blagitko et al., 1999;
Futatsumori et al., 2000; Wegmann et al., 2004). The two isotypes of each subunit were
termed γ1 (ζ1) and γ2 (ζ2), respectively. The different isotype combinations are present in
diverging amounts within the cell: γ1ζ1 (53%), γ1ζ2 (16%), γ2ζ1 (26%) and γ2ζ2 (5%), and
localize differentially within the Golgi (Moelleken et al., 2007). These findings strongly
suggest that the different isotypes mediate specific transport steps within the Golgi, however,
no distinct binding partners interacting with one of the isotypes have been characterized to
date.
Proteins following the Secretoy Pathway through the Golgi undergo repetitive shuttling
processes, and, in each cisterna, are sorted for their respective destination. This model was
first described in the "Destillation-hypothesis" (Dunphy et al., 1981; Rothman, 1981), that
implied that in each step, anterograde cargo is preferentially incorporated in COPI vesicles
moving in direction of the TGN, and retrograde cargo is transported back in direction of the
ER. Thus, in each transport step from one cisterna to the next on the route through the Golgi,
cargo is further enriched by selective uptake into the vesicles going in the correct direction.
Thus, proteins traversing the Golgi via the Secretory Pathway are both modified by the
corresponding enzymes in the different cisternae, and sorted for their corresponding
destinations.
At the next station, the Trans Golgi Network (TGN), transport is mediated mainly by clathrincoated vesicles (CCVs), that are made up from three core components: A small GTPase, an
adaptor protein complex and the scaffolding protein clathrin (see 1.2.1 Clathrin-coated
vesicles). The TGN comprises the second big sorting station in intracellular trafficking. Here,
on one hand, endocytic cargo is transported from the plasma membrane via CCVs. On the
other hand, anterograde cargo is distributed to its respective destinations by two different
types of transport: regulated exocytic transport via secretory granules, and constitutive
transport via clathrin-coated vesicles.
Constitutive transport is mediated by various forms of clathrin-coated vesicles. Cargo
destined for organelles or the plasma membrane is sorted by different adaptor protein
complexes. Two classes of these complexes take part in this process: First, adaptor protein
complexes that recognize cargo by distinct peptide sequences or monoubiquitination (i.e.
1. Introduction
5
AP-complexes, GGAs or β-Arrestins), and second, lipid binding complexes (AP180/CALM,
Epsins and Amphiphysins).
For example, lysosomal hydrolases are recognized via attached mannose-6-phosphate
(M6P) groups by M6P-receptors, and incorporated into vesicles destined for endosomes and
subsequently lysosomes. Monoubiquitinated proteins are taken up into vesicles that are
targeted towards late endosomes, and lysosomes. Mis- or unfolded proteins are targeted to
the lysosomes for degradation.
Regulated exocytic transport only takes place in endocrine or neuroendocrine cells. Here,
prohormones are taken up into secretory granules that allow release of the mature hormones
after an extracellular stimulus (Tooze et al., 2001). In the formation of secretory granules no
coating proteins have been described to date. However, clathrin-coated vesicles are involved
in the removal of mislocalized proteins (i.e. secretory proteins) from the secretory granules.
1.2.
Vesicular Transport: The three coating systems
As described above, three types of transport vesicles have been characterized in eukaryotic
cells so far:
Clathrin-coated vesicles (CCVs) that shuttle between the trans-Golgi network, endosomes,
lysosomes and the plasma membrane; COPII-coated vesicles that mediate exit from the ER;
and COPI-coated vesicles that function within the early Secretory Pathway (reviewed by
(McMahon and Mills, 2004)). All intracellular vesiculation systems share certain homologies
and the following chapter focuses on the characteristics of each system.
1.2.1.
Clathrin-coated vesicles
Clathrin-coated vesicles are structured in three layers. The innermost layer is comprised of
the membrane bilayer containing the transmembrane cargo proteins. It is linked to the outer,
scaffolding clathrin layer by a middle layer of adaptor proteins. These adaptor proteins are a
disparate group of proteins with no obvious sequence homology (see 1.2.1.1 Structural
studies on clathrin-coated vesicles). They are termed adaptor proteins as they bind to both
transmembrane components and clathrin, and hereby mediate a link ("adaptor") between the
cargo in the innermost layer and the scaffolding on the outside.
Clathrin-coated vesicles were first described by Roth and Porter in 1964 (Roth and Porter,
1964), studying the yolk uptake of mosquito oocytes. First electron microscopy images of the
soccerball-like structure of a clathrin-coated vesicle were obtained by Kanaseki and Kadota
1. Introduction
6
1969 (Kanaseki and Kadota, 1969), and the outer scaffolding protein, clathrin, was
subsequently discovered by Pearse (Pearse, 1975).
At the same time, the adaptor proteins, the middle layer of CCVs, were identified (Keen et
al., 1979). Further biochemical characterization revealed that there are at least two classes
of these proteins (Keen et al., 1987): The tetrameric adaptor protein complex 1 (AP-1),
localized to the trans-Golgi network and the tetrameric adaptor protein complex 2 (AP-2),
localized on the plasma membrane (Ahle et al., 1988). As different kinds of cargo are
transported at the TGN and the plasma membrane, it was speculated that not clathrin, but
the adaptor protein complexes confer specificity for sorting distinct cargo to the different
CCVs. Indeed, Ohno et al found that the adaptor complexes bind to short targetting
sequences present in cargo molecules for the CCVs (Ohno et al., 1995). Subsequently, other
adaptor protein complexes were identified (AP-3, (Dell'Angelica et al., 1997); and AP-4,
(Dell'Angelica et al., 1999)). However, direct interaction of these adaptor complexes with
clathrin has not been demonstrated to date.
Other, monomeric adaptor proteins were discovered later: Golgi-localizing, γ-adaptin ear
homology domain, Arf binding proteins (GGAs, (Dell'Angelica et al., 2000)), AP180/CALM
and the epsins. Like the tetrameric adaptor proteins, the monomeric adaptor proteins are
able to cross-link the outer scaffolding clathrin layer to the innermost transmembrane
components.
Like in the other vesiculation systems, recruitment of the adaptor proteins to the membrane
is governed by a small GTPase. Arf1 has been described in both AP-1 (Austin et al., 2000),
AP-3 (Ooi et al., 1998), AP-4 (Boehm et al., 2001) and the GGAs (Dell'Angelica et al., 2000),
and Arf6 was implied in mediating AP-2 recruitment (Paleotti et al., 2005)). However, in
contrast to the other vesiculation systems, coat release seems to be independent of the
nucleotide state of the small GTPase, but to be ATP (Holstein et al., 1996) and
phosphorylation-dependent, as phosphorylation of AP-2 in its α and β2 subunits, and AP-1 in
its β1-subunit, leads to dissociation of the clathrin coat (Jha et al., 2004; Wilde and Brodsky,
1996).
1.2.1.1.
Structural studies on clathrin-coated vesicles
Clathrin has an unusual structure, termed "Triskelion": It is made up of three extended
subunits, which radiate from a central hub (Ungewickell and Branton, 1981). A small protein
was identified that was attached to these "legs", when isolating clathrin from CCVs. The two
1. Introduction
7
proteins were termed clathrin "heavy chain" (190kD) and "light chain" (25kD; (Kirchhausen
and Harrison, 1981)).
Each leg, i.e. each heavy chain, is about 475nm long (Kirchhausen et al., 1986), and the
elongated structure is achieved by a heptad repeat of a central 71-residue motif, termed αsolenoid structure (clathrin heavy chain repeat). Each leg is comprised of the following
subdomains: A proximal segment (adjacent to the central hub), a knee-like subdomain, a
distal segment, and an ankle, a linker and a terminal domain, that is comprised of WD-40
repeats forming a β-propeller domain (Fotin et al., 2004a). The light chains are attached to a
heavy chain via each proximal segment close to the central hub (Figure 2, taken from Fotin
et al., 2004).
Figure 2: Scheme of the Clathrin triskelion and designs of Clathrin lattices (adapted from
Fotin et al, 2004).
Upon coat assembly, clathrin assembles into a lattice of open hexagonal and pentagonal
faces (Musacchio et al., 1999). The resulting coats show various designs, ranging from small
assemblies with 28 ("mini-coats"), 36 ("hexagonal barrels") and 60 ("soccer balls") clathrins
to even bigger aggregates (Crowther et al., 1976). In vitro reconstitution of clathrin-coated
vesicles from the isolated components yield relatively homogeneous preparations, with a
large proportion of hexagonal barrels (Vigers et al., 1986a; Vigers et al., 1986b). These
vesicles have been studied extensively using negative stain (Crowther et al., 1976) and cryo
EM (Fotin et al., 2004a; Fotin et al., 2004b; Fotin et al., 2006; Musacchio et al., 1999; Smith
1. Introduction
8
et al., 1998; Vigers et al., 1986a; Vigers et al., 1986b). The clathrin triskelions assemble in a
pattern where the clathrin legs make up the edges, and the triskelion hubs lie at the vertices
(Smith et al., 1998). The legs radiate from the central hub, project slightly inward, and bend
at both the knee and the ankle segments. Thus the interaction of different clathrin molecules
to form a lattice is mediated by the α-solenoid structures interdigitating in the edges of the
coat, and the interaction with the middle layer, i.e. the adaptor proteins, is mediated by the βpropeller in the terminal domain (Figure 2, taken from Fotin et al., 2004).
Figure 3: Structural similarities of coat proteins. In the table, protein subunits of the various
coats are listed along with their functional homologies.
The structure of the adaptor proteins has also been extensively studied. The crystal
structures of the core of both the adaptor protein AP-1 ((Heldwein et al., 2004)) and AP-2
(Collins et al., 2002) have been solved. Both tetrameric complexes exhibit a characteristic
design, with two domains (the α and the β subunits) containing a trunk and an appendage
domain linked by a flexible hinge, as illustrated in Figure 3 (modified from (Langer et al.,
2007)), and interact with a wide range of binding partners. For example, AP-2 binds directly
to phosphoinositides (Beck and Keen, 1991; Collins et al., 2002; Gaidarov and Keen, 1999;
Rohde et al., 2002), cargo proteins (Ohno et al., 1995), cargo receptors (Claing et al., 2002)
and coating machinery, like GTPases (Paleotti et al., 2005) and the scaffolding clathrin
(Zaremba and Keen, 1983), reviewed by (Sorkin, 2004).
1. Introduction
9
Monomeric adaptors, like the GGAs, show a similar design only by presence of the
characteristic trunk-hinge-appendage subdomain structure (Figure 3), binding to cargo via
their VHS (VPS-27, Hrs and STAM) domain (Puertollano et al., 2001a; Zhu et al., 2001), to
the small GTPase Arf via its GAT (GGA and Tom1) domain (Boman, 2001; Boman et al.,
2000), and to clathrin via clathrin boxes in the hinge region (Puertollano et al., 2001b).
The adaptor proteins as the linker between cargo proteins in the membrane layer and the
outer scaffold clathrin have a special role during the coating process. Their affinity for cargo
must be tightly controlled, allowing specific binding and recruitment to clathrin-coated pits
that must be activated during the budding process only. This is achieved by conformational
changes of the adaptor proteins, that are selectively triggered upon either phosphorylation or
interaction with specific ligands. The following chapter focuses on these conformational
changes in adaptor proteins.
1.2.1.2.
Conformational changes in adaptor proteins
1.2.1.2.1. The adaptor protein 2 (AP-2) system
The AP-2 complex and its vital role in clathrin-mediated endocytosis is the most extensively
studied vesicle coating system. It is also the first coating protein in which a conformational
change was described (Matsui and Kirchhausen, 1990). Using limited proteolysis, Matsui et
al. observed that the µ2-subunit of soluble AP-2 was more resistant to trypsin digestion than
that of clathrin-coated vesicle-bound AP-2. Investigating this, Fingerhut et al. found that
binding of AP-2 to cargo can be modulated by phosphorylation (Fingerhut et al., 2001).
Phosphorylation induced a 30-fold increase in affinity of AP-2 to Yxxφ-based cargo-sorting
signals, and phosphorylated AP-2 exhibited an increased binding to membranes. It was later
shown that phosphorylation of µ2 is also essential for endocytosis (Olusanya et al., 2001).
In 2002, the x-ray structure of the core of AP-2 (without the ear domains of α- and β2adaptin) was solved by Owen and coworkers (Collins et al., 2002). In this crystal structure,
the binding pocket for Yxxφ motifs is inaccessible, as it is covered by β2-adaptin. It was
concluded that the crystallized form corresponds to the inactive, cytosolic form of AP-2, and
that a conformational change would be required to open the binding site on µ2. It is likely that
phosphorylation of Thr156 in the µ2-subunit, located in the linker connecting two µ2-domains,
induces this conformational change, and that the increased affinity of phosphorylated AP-2
for cargo is due to the conformational change opening its binding pocket. Thus AP-2's affinity
for cargo is directly modulated by a phosphorylation-induced conformational change.
1. Introduction
10
Later, in 1999, Haucke et al. found a conformational change induced in AP-2 upon binding of
Yxxφ-based cargo signals. The authors investigated the interaction of AP-2 with
synaptotagmin, a transmembrane protein that binds to AP-2 via its C2B domain and may act
as an AP-2 recruitment site at the plasma membrane (Jorgensen et al., 1995; Zhang et al.,
1994). In the presence of Yxxφ-signal-bearing peptides, binding of synaptotagmin to AP-2
and recruitment of AP-2 to membranes was increased (Haucke and De Camilli, 1999). A
conformational change of AP-2 upon binding of a Yxxφ-signal-containing peptide was shown
by altered susceptibility to limited proteolysis of AP-2's α-adaptin core domain. Later studies
revealed that synaptotagmin binds AP-2 via µ2- and α-adaptin, and that the binding affinity of
isolated µ2 (outside the context of the AP-2 complex) to a C2B-domain was not increased in
the presence of a Yxxφ-signal-containing peptide (Haucke et al., 2000). Therefore, it was
proposed that the conformational change observed within AP-2 (Haucke and De Camilli,
1999) includes other subunits of the complex.
The conformational change induced by binding of a Yxxφ-signal was interpreted as an
activation of the AP-2 complex for cargo binding. This hypothesis was further supported by
Boll et al. (Boll et al., 2002), who investigated binding of AP-2 to the low density lipoprotein
(LDL) receptor via its signal sequence FDNPVY (consensus sequence: FxNPxY). They
showed that LDL receptor signal-containing proteins bound the µ2 subunit on a site different
from both synaptotagmin's C2B binding site and the Yxxφ-signal binding site, and that the
presence of Yxxφ-peptides increased AP-2's binding to a FDNPVY protein. Like Haucke
(Haucke and De Camilli, 1999), they detected a conformational change in AP-2 upon
addition of a peptide containing a Yxxφ-sequence. Again the susceptibility of β-adaptin to
proteases changed in the presence of Yxxφ-peptide. However, the binding site for FxNPxYmotifs remains unknown.
It is generally believed that the AP-2 complex undergoes a structural rearrangement during
membrane recruitment that regulates its ability to bind to various cargo signals. It is of note
that the conformational change observed by Haucke (Haucke and De Camilli, 1999) and Boll
(Boll et al., 2002) is not induced by phosphorylation, as no kinases were present in their
experiments. Thus, it may be speculated that two sequential conformational changes of the
AP-2 complex take place during its recruitment to the donor membrane: the first one by
phosphorylation to recruit the complex to the membrane and allow Yxxφ-signal binding, and
the second one, induced by ligand binding to the now-accessible Yxxφ-binding site, to allow
for subsequent binding of additional proteins.
1. Introduction
11
The role of Yxxφ-signal-containing proteins in clathrin-mediated endocytosis was further
analyzed by Haucke (Haucke and Krauss, 2002). Using a liposome-based system, peptides
bearing these signals were shown to induce AP-2 clathrin cage formation. Cargo proteins
have been proposed to be crucial parts of the vesicle formation machinery, based on in vivo
data showing that, of the large number of clathrin-coated pits formed spontaneously on the
plasma membrane, only a small fraction proceed to form clathrin-coated vesicles (Ehrlich et
al., 2004). The authors proposed that the presence of cargo proteins, e.g. Yxxφ-signal
containing proteins, stabilizes coated pits and allows them to proceed to form vesicles.
1.2.1.2.2. The adaptor protein 1 (AP-1) system
The AP-1 complex is thought to mediate transport from the TGN to endosomes/lysosomes
(Traub and Kornfeld, 1997), although recent studies have proposed a role for AP-1 in
recycling from endosomes to the TGN (Meyer et al., 2001; Meyer et al., 2000). The minimal
machinery for AP-1 recruitment has been characterized (Crottet et al., 2002; Meyer et al.,
2005; Wang et al., 2003). It is composed of the small GTPase Arf1 in its active, GTP-bound
form, phosphatidylinositol phosphates as lipid components, and Tyr-based cargo signals
present on the donor membrane. Like AP-2, AP-1 was shown to bind to certain cargo motifs
(e.g. the cytoplasmic tails of cation-dependent and cation-independent mannose-6phosphate receptors (MPR; (Glickman et al., 1989; Mathews et al., 1992; Meresse and
Hoflack, 1993))), and to interact with the coating machinery GTPase Arf1 (Nie et al., 2003)
and with clathrin (Zaremba and Keen, 1983).
Cargo binding to AP-1 is, at least in part, also regulated by phosphorylation (Ghosh and
Kornfeld, 2003a). Like in the AP-2 system, AP-1's affinity to cargo peptides decreased
dramatically upon dephosphorylation of AP-1 (Ghosh and Kornfeld, 2003a). Limited
proteolysis of phosphorylated and dephosphorylated AP-1 revealed a conformational
change, as the µ1-subunit exhibited higher susceptibility to trypsin in the phosphorylated
form. The observed increases in susceptibility to limited proteolysis and cargo binding upon
phosphorylation is analogous to the behaviour of AP-2 described by Fingerhut et al.
(Fingerhut et al., 2001) and Ricotta et al. (Ricotta et al., 2002), indicating that, in both
systems, a phosphorylation-induced conformational change is employed to activate cargobinding of the adaptor protein complex. A likely candidate to mediate phosphorylation of AP1 in vivo is GAK (Umeda et al., 2000).
Upon interaction of AP-1 with cargo peptides, i.e. the CI-MPR internal dileucine-type cargo
motifs, a conformational change occurs within the core complex (Lee et al., 2008). This
1. Introduction
12
conformationally changed AP-1 showed an increased affinity for Tyr- based cargo motifs
(Yxxφ), and the functional cross-talk between the binding sites for the dileucine-type motif
and the tyrosine-based motif was found to be bidirectional. In addition, the conformational
change also greatly stimulated binding of AP-1 to the small GTPase Arf1 in its GTP-bound
form. Thus, in the AP-1 clathrin system, transmembrane cargo proteins serve as a part of the
machinery to stabilize the Adaptor complexes on the membrane (Lee et al., 2008).
1.2.1.2.3. Golgi-localizing, γ-adaptin ear homology domain, Arf-binding proteins
Similarly, binding of cargo to the monomeric adaptor proteins GGAs (Golgi-localizing, γadaptin ear homology domain, Arf-binding proteins) is modulated by phosphorylation (Ghosh
and Kornfeld, 2003b).
In GGAs, phosphorylation induces an effect opposite to that in adaptor complexes. Binding of
GGA1 to cargo proteins (MPR) increases upon dephosphorylation of GGA1, whereas the
inactive, phosphorylated form is cytosolic (Ghosh and Kornfeld, 2003b). According to Doray
et al. (Doray et al., 2002), an internal AC-LL motif in GGAs 1 and 3 binds to and blocks the
VHS binding site upon phosphorylation of Ser355, and dephosphorylation relieves
autoinhibition. In order to investigate the phosphorylation-dependent conformational
dynamics of GGA1/3, phosphorylated and dephosphorylated GGA1 were submitted to gel
filtration and sucrose gradient analysis. A striking 2nm change in the Stokes radius of GGA1
was found upon phosphorylation, indicating a conformational change. The authors attributed
this change to the AC-LL motif-containing hinge domain moving out of the VHS domainbinding pocket (Doray et al., 2002; Ghosh and Kornfeld, 2003b).
1.2.2.
COPII-coated vesicles
Like in the clathrin system, a COPII coated vesicle is made up from two major coat building
units: The adaptor protein-like Sec23/24 dimer, and the clathrin-like Sec13/31 dimer, that are
recruited sequentially to the donor membrane. The structure of Sec23/24 and Sec13/31
heterodimers purified from yeast cells was first characterized by Lederkremer et al
(Lederkremer et al., 2001) using negative stain EM. They described both the characteristic
bone-like shape of the Sec23/24 heterodimer, and the elongated shape of the Sec13/31
dimer. The Sec13/31 complex is comprised of 5 globular domains arranged like pearls on a
string, and make up the 28-30nm sized building unit of the outer scaffold of the COPII cage.
The pre-budding complex of Sec23/24 with bound Sar1p was co-crystalized in 2002 by Bi
and Goldberg (Bi et al., 2002), showing an intrinsic curvature of the lipid binding face
matching that of COPII vesicles.
1. Introduction
13
In 2006, the structure of minimal COPII cages consisting of Sec13/31 heterodimers was
described by Stagg et al (Stagg et al., 2006). Using cryo-EM and single-particle averaging,
they reconstructed a 30Å resolution map of self-assembled Sec13/31 cages. These cages
contain 24 heterodimers of Sec13/31, and exhibit octahedral symmetry. In a cuboctahedron,
four edges intersect at each vertex, which is strikingly different from the clathrin system,
where a vertex is defined by crossing of three edges (see also 1.2.1 Clathrin coated
vesicles). In 2007, the structure of the Sec13/31 complex was solved by Fath and Goldberg
(Fath et al., 2007), and fitted into the 30Å map previously described by Stagg et al (Stagg et
al., 2006). Although the Sec13/31 dimer resembles clathrin (it features an α-solenoid, rod-like
structure, with a terminal domain comprised of a β-propeller motif), the relative orientation of
the domains of the coating protein in the assembled coat is strikingly different from CCVs.
Whereas in CCVs clathrin's β-propeller domains are pointing inward and are interacting with
the adaptor proteins, in COPII cages the β-propellers of the Sec13/31 dimers actually
comprise the 12 vertices of the cuboctahedron, and the α-solenoid structures do not
interdigitate (Fath et al., 2007; Stagg et al., 2006). Thus, although clathrin and COPII coats
utilize similarly designed building units (α-solenoid and β-propeller motifs) they do not seem
to share a common construction principle.
Like in the other vesiculation systems, the coating process is governed by a small GTPase in the COPII system, Sar1p (Nakano and Muramatsu, 1989). The GTPases' localization is
determined by the bound nucleotide: The GDP-form is a soluble protein, whereas the GTPbound form is bound to the corresponding donor membrane by an anchor located in the Nterminal domain. In the case of Sar1p, an amphiphatic helix is exposed upon Sec12mediated GDP-to-GTP-exchange, that inserts into the lipid bilayer and effectively locks
Sar1p on the membrane. Sec12 localizes to the ER (Barlowe et al., 1993; Barlowe and
Schekman, 1993), and Sec12-mediated GTP-exchange is thought to convey specificity of
COPII budding to ER membranes. Sequentially, Sar1p recruits the coating protein
complexes, first heterodimeric Sec23/24, and then Sec13/31 (Matsuoka et al., 1998). Sar1p
is then released from the membrane by Sec23 (the corresponding GAP), whose activity is
increased by Sec13/31 on the donor membrane (Antonny et al., 2001; Yoshihisa et al.,
1993). Sar1p dissociation from the membrane also leads to coat disassembly.
1. Introduction
1.2.3.
14
COPI-coated vesicles
COPI-coated vesicles were discovered in a cell-free assay that was used to study transport
between Golgi cisternae (Malhotra et al., 1989). Subsequently, the proteins comprising the
coat of these vesicles were identified: A large protein complex termed "coatomer" (Waters et
al., 1991), and the small GTPase Arf1 (Serafini et al., 1991).
Arf1 is a member of the Ras family of small GTPases, and is myristoylated on its N-terminal
glycine residue, a modification that is prerequisite for membrane binding and activity (Kahn et
al., 1991).
Coatomer is a heptameric complex with a molecular weight of ~600kD (Waters et al., 1991).
It is comprised of a trimeric and a tetrameric subcomplex (Figure 3, table 1, and reviewed in
(McMahon and Mills, 2004)), and the whole complex seems to remain fully assembled during
its life time in the cell (Lowe and Kreis, 1996).
Table 1: Coatomer subunits: Size, motifs and interaction partners.
In the budding process, coatomer is recruited from the cytosol in a single step in an Arfdependent manner (Hara-Kuge et al., 1994). This recrutiment en bloc is contrary to the
clathrin and the COPII system, where two sequential steps are required to assemble first the
adaptor proteins (or Sec23/24) and second the scaffolding proteins (clathrin or Sec13/31).
This means that the proteins analogous to those forming the CCVs' inner shell and outer
shell are recruited at the same time.
1. Introduction
15
Structural similarities exist between the trimeric subcomplex comprised of the subunits α, β'
and ε COP and the clathrin proteins, and between the tetrameric subcomplex consisting of β,
γ, δ and ζ-COP and the 4 subunits of the adaptor complexes AP-1 to AP-4 of clathrin-coated
vesicles (Eugster et al., 2000; Gurkan et al., 2006; McMahon and Mills, 2004; Schledzewski
et al., 1999). Like in clathrin, the trimeric subcomplex contains α-solenoid structures and βpropeller domains (table 1, reviewed in (Devos et al., 2004)). The tetrameric adaptor proteinlike subcomplex of coatomer also contains two subunits with the characteristic trunk-hingeappendage-domain design also observed in the APs ((Eugster et al., 2000; Schledzewski et
al., 1999; Watson et al., 2004), reviewed by (McMahon and Mills, 2004)). However, only
limited structural data has been obtained on coatomer architecture. Only the structure of the
appendage domain of γ-COP has been solved, showing a similiar overall fold as the αappendage domain of AP-2 (Watson et al., 2004).
Thus, coatomer contains building units similar to those in the COPII / clathrin system. Their
functions, though, seem to differ from those found in the other systems, as described below.
1.2.3.1.
The COPI budding process
The COPI budding process can be divided up into 4 steps (Figure 4). In the following
chapter, the most important aspects of each step will be discussed.
1. Arf1 recruitment (Figure 4, step 1).
In its inactive, GDP-bound form, the small GTPase Arf1 resides in the cytosol. Arf1 has been
described to take part in the biogenesis of AP-1 (Austin et al., 2000), AP-3 (Ooi et al., 1998),
AP-4 (Boehm et al., 2001) and COPI (Donaldson et al., 1992a; Palmer et al., 1993; Serafini
et al., 1991) vesicles. Specificity for the respective donor membrane is presumably conveyed
by two factors: transmembrane proteins that bear cytoplasmic receptor motifs and the
corresponding nucleotide exchange factors (GEFs).
1. Introduction
16
Figure 4: Scheme of the COPI budding process.
Two members of the p24-family of class I transmembrane proteins, p23 and p24, have been
characterised to recruit Arf1-GDP to the donor Golgi membrane (Contreras et al., 2004;
Gommel et al., 2001; Majoul et al., 2001), proposing these cytoplasmic domains as primary
Arf1-GDP receptors. In addition, the early Golgi SNARE protein membrin has been put
forward to act as an Arf1-GDP receptor (Honda et al., 2005).
The nucleotide exchange factors (GEFs) catalyze the exchange of GDP to GTP on Arf1, and
share a characteristic Sec7 domain (Chardin et al., 1996). Sec7-binding to Arf1 is modulated
by interactions with the lipid bilayer (Itoh and De Camilli, 2004; Terui et al., 1994). The GEF
involved in COPI biogenesis is presumably GBF1, a large Sec7 domain GEF localized to the
cis-Golgi (Niu et al., 2005; Zhao et al., 2002). GBF1 also showed sensitivity to Brefeldin A
(BFA, (Niu et al., 2005; Zhao et al., 2002)), a fungal metabolite that was previously shown to
inhibit nucleotide exchange on Arf1 (Donaldson et al., 1992b).
Binding of GTP leads to a conformational change in Arf1, exposing the N-terminal myristoylanchor and an amphiphatic helix, effectively anchoring Arf1-GTP on the donor membrane
(Antonny et al., 1997; Franco et al., 1996). This conformational change also ensures the
1. Introduction
17
specificity of the GDP-GTP-exchange to the donor membrane, as the exposure of the
myristoyl-anchor is energetically unfavored in the cytosol.
2. Coatomer recruitment (Figure 4, step 2).
Arf1, in the activated GTP form, dissociates from the transmembrane receptors, i.e. p24family proteins. A "priming complex" is formed, consisting of membrane-bound Arf1-GTP and
oligomers of p24-family members. This complex is competent to recruit coatomer to the
donor memrbane.
Coatomer binds to Arf1 via multiple interactions: ε-COP (Eugster et al., 2000), β- and γCOP (Zhao et al., 1997; Zhao et al., 1999), and β' and δ-COP, as recently described (Sun et
al., 2007b). The interaction with δ-COP was found to be nucleotide-dependent only in the
context of the whole coatomer complex, as isolated δ-COP bound both Arf1-GDP and Arf1GTP. Thus the binding pocket for Arf1 in δ-COP may be opened upon GTP-dependent
membrane recruitment of coatomer, and may be reminiscent of the induced activation of AP2 by phosphorylation during membrane recruitment.
The coating complex coatomer is, as described above, recruited en bloc. On the membrane,
coatomer binds to cytoplasmic domains of transmembrane proteins, that are described
below.
3. Interaction of coatomer with transmembrane proteins: Cargo binding and
conformational change (Figure 4, step 3).
On the donor membrane, coatomer interacts via multiple subunits with the cytoplasmic tails
of transmembrane proteins (Bethune et al., 2006; Dominguez et al., 1998; Letourneur et al.,
1994; Sohn et al., 1996). Two classes of interaction partners have been characterized to
date:
1. The p24-family proteins: coat receptors.
The p24-family are class I transmembrane proteins, termed p23, p24, p25, p26 and p27.
They are thought to cycle in the early Secretory Pathway (Blum et al., 1999; Fullekrug et al.,
1999; Nickel et al., 1997; Rojo et al., 2000), with distinct localizations for the individual
members (Dominguez et al., 1998; Fullekrug et al., 1999; Nickel et al., 1997; Rojo et al.,
2000; Rojo et al., 1997; Sohn et al., 1996; Wada et al., 1991). Members of the p24-family all
consist of N-terminal signal sequence, a pair of conserved cysteins and a sequence
predicted to form a coiled-coil domain in the lumenal domain, a transmembrane segment,
and a small cytosplasmic domain (Figure 5).
1. Introduction
18
Figure 5: Scheme of p24-family members structure and sequences of cytoplasmic domains.
The coiled-coil domain is presumably responsible for the formation of homo- and heterooligomers of the p24-family members (Emery et al., 2000; Fullekrug et al., 1999), a
prerequisite for the distinct interaction of their cytoplasmic tails with cytoplasmic proteins. The
cytoplasmic domains of all p24-family members contain two distinct motifs: A diphenylalanine
(FF, colored in red in Figure 5) motif and a dibasic motif (KKxx, KKxn or KRxn with n >3) at
their C-termini (imaged in blue in Figure 5). Binding to coatomer is dependent on both motifs,
as mutation of either the KKxx- or the FF-residues significantly reduces binding, and
mutation of both motifs abolishes the interaction (Bethune et al., 2006; Cosson and
Letourneur, 1994; Reinhard et al., 1999; Sohn et al., 1996).
p24-family members bind to caotomer not via α- and β'-COP, but via the subunit γ-COP
(Bethune et al., 2006; Fiedler et al., 1996; Goldberg, 2000). The interaction of coatomer with
the dimeric cytoplasmic domain of the transmembrane protein p23 was shown to induce a
conformational change and aggregation of the complex in vitro (Reinhard et al., 1999). This
aggregation likely represents coatomer polymerization during COPI vesicle formation, as
COPI vesicle-bound coatomer and coatomer polymerized by the cytoplasmic domain of
dimeric p23 in vitro showed identical patterns after limited proteolysis, indicating similar
conformations (Reinhard et al., 1999). Recently, it was shown that all members of the p24family proteins and some additional cycling membrane proteins bind as dimers to two
independent sites in the appendage and the trunk domain of the coatomer subunit γ-COP,
and that dimers of p23 induce a conformational change in the trunk domain of γ-COP
(Bethune et al., 2006).
Although COPI vesicles can be generated from liposomes with a certain lipid composition
using coatomer and Arf1 alone (Spang et al., 1998), the presence of lipopeptides
representing cytoplasmic domains of p24-family members allows COPI vesicle budding
independent of the lipid composition of the donor membranes (Bremser et al., 1999). They
were also found to be a stoichiometric component of a population of COPI vesicles (Fiedler
1. Introduction
19
et al., 1996; Sohn et al., 1996). However, recently a population of COPI vesicles devoid of
p24-family members has been described (Malsam et al., 2005). Here, other, yet unidentified
transmembrane proteins may take over the role of the p24-family members.
2. The KKxx / KxKxx motif: Cargo.
The dibasic ER retrieval signals are the best characterized cargo sorting motifs binding to
coatomer (Cosson and Letourneur, 1994; Letourneur et al., 1994). KKxx motifs were shown
to bind to the coatomer subunits α- and β'-COP (Letourneur et al., 1994), and recently the
distinct but overlapping binding sites for both KKxx and KxKxx motifs were identified in the βpropeller / WD-40 repeats at the N-termini of α- and β'-COP (Eugster et al., 2004). Most
KKxx / KxKxx-motif bearing proteins bind with an affinity in the low micromolar range
(Bethune et al., 2006). Thus, the β-propeller domains on α- and β'-COP serve a function
distinct from or additional to the one observed in both the clathrin system (interaction with the
adaptor proteins) and the COPII system (formation of vertices).
3. Other cargo binding motifs:
Proteins cycling between the ER and the early Golgi contain a specific signal sequence that
allows them to exit the ER to travel to the Golgi, and then be captured and transported back.
A KDEL-motif acting as an ER retrieval signal in the C-terminal of lumenal ER resident
proteins was identified by Munro and Pelham (Munro and Pelham, 1987; Pelham et al.,
1988). The corresponding receptor ("KDEL-receptor", (Lewis and Pelham, 1990; Semenza et
al., 1990) was characterized as an integral membrane proteins with 7 transmembrane spans
(Scheel and Pelham, 1998), and binds to coatomer via a dilysine-motif and a phosphorylated
serine (Cabrera et al., 2003). Binding of ligand (i.e. KDEL proteins) is pH-dependent (Wilson
et al., 1993), induces oligomerization of the receptor (Majoul et al., 1998) and leads to
retrieval of the receptor-protein complex from the Golgi to the ER (Lewis and Pelham, 1992)
by the COPI pathway (Majoul et al., 2001).
Two other binding motifs in cytoplasmic domains of transmembrane proteins have been
identified to date: The δ-L-motif with the consensus sequence WxxW/Y/F, mediating binding
to the coatomer subunit δ-COP, identified in yeast in the ER-localized Sec71p protein
(Cosson et al., 1998); and the RxR motif, first described in the cytoplasmic loops and Cterminal tails of the ATP-sensitive K+ channel (Yuan et al., 2003; Zerangue et al., 1999). The
binding site for RxR-motifs on coatomer has recently been identified on β- and δ-COP
(Michelsen et al., 2007), and was found to show a striking resemblance to the Yxxφ-binding
site of the adaptor complexes in the clathrin system.
1. Introduction
20
4. Uncoating and fusion with the target membrane (Figure 4, step 4).
COPI coat hydrolysis is mediated by nucleotide exchange on Arf1, i.e. GTP- to GDPexchange on Arf1 leads to coat dissassembly (Tanigawa et al., 1993). After complete
budding, GTP hydrolysis is induced by interaction with an ArfGAP (Cukierman 1995), a
process that could be reconstituted in vitro by addition of the purified catalytic domain of
ArfGAP1 to COPI vesicles generated from purified Golgi membranes (Reinhard et al., 2003).
Two other ArfGAPs have been shown to interact with coatomer: ArfGAP2 (Randazzo, 1997)
and ArfGAP3 (Liu et al., 2001). Thus, for preparation of stable, coated COPI vesicles in vitro
using GTP, no ArfGAP proteins must be present.
Fusion of a COPI vesicle with its target membrane is governed by two classes of proteins:
SNAREs (soluble NSF attachment protein receptors, (Sollner et al., 1993)) and tethering
proteins. Initial binding of the vesicles to their respective target membranes is mediated by
tethering proteins. There are two classes of tethering proteins: Elongated coiled-coil proteins
(Barr and Short, 2003) and large multiprotein complexes (Whyte and Munro, 2002). While
multiprotein complexes have mostly been described in exocytosis (Whyte and Munro, 2002),
coiled-coil proteins are involved in Golgi and endosomal fusion steps: The coiled-coil proteins
Giantin and Golgin 84 are taken up into COPI vesicles, and mediate together with p115,
GM130 and GRASP65/CASP docking of the vesicle to the target membrane.
The specificity of vesicle-membrane fusion is ensured by complementary pairing of SNARE
proteins (McNew et al., 2000): The target membranes contain three-helical target(t)SNAREs, that form a tetrahelical bundle with the vesicle(v)-SNARE present in the COPI
vesicle (Parlati et al., 2000; Weber et al., 1998).
1.2.3.2.
Summary: Comparison of COPI with the other vesiculation systems
The different coating proteins share some striking structural and functional homologies.
However, in each system, similar motifs are put to different use, and the budding process
differs on the molecular, functional level.
All vesiculation systems exhibit a three-layered structure: The inner layer is comprised of the
lipid bilayer, and contains the transmembrane cargo and transmembrane coat recruitment
machinery. The middle layer containing the adaptor- and adaptor-like proteins mediates the
interaction between the outer, scaffolding coating protein layer and the transmembrane layer.
There are two conserved building motifs found in the coating proteins of all three vesiculation
systems: α-solenoid structures and β-propeller domains. The α-solenoid structures are
curved, elongated domains comprised of alpha helices arranged in a manner similar to a
Jelly fold. While functions of the α-solenoid structures in the COPI system have not been
1. Introduction
21
elucidated to date, these domains have distinct functions found in the COPII and the clathrin
system: In clathrin, these domain form the legs of the clathrin molecule, and interdigitate to
form the clathrin lattice (see 1.2.1.1 Structural studies on clathrin-coated vesicles). In the
COPII system, in the Sec13/31 dimer, the α-solenoid structures do not interdigitate. Here,
they serve as the linkers, forming the edges of the Sec13/31 lattice (see 1.2.2 COPII-coated
vesicles).
The second building unit, the β-propeller domains, also have distinct functions in the three
vesiculation systems: In the clathrin system, the terminal β-propellers serve as an interaction
hub to mediate binding to the adaptor proteins and accessory proteins (see 1.2.1.1 Structural
studies on clathrin-coated vesicles). In the COPII system, these domains form the vertices of
the Sec13/31 cage (see 1.2.2. COPII-coated vesicles), and in the COPI system, they contain
the binding sites of the coatomer complex to dibasic ER retrival signals (see 1.2.3.1 The
COPI budding process).
The COPI system shows additional structural homology to the adaptor proteins. Two
subunits of the adaptor-like subcomplex of coatomer, γ-COP and β-COP, display a subunit
design similar to the one found in the adaptor proteins and the GGAs: The subunits contain a
trunk and an appendage domain that are connected by a flexible linker, a hinge region.
However, the COPI coat exhibits a striking difference to the other two vesiculation systems:
In both the COPII and the clathrin system, coat recruitment is mediated in two sequential
steps. The COPI coating protein, coatomer, though, is recruited to the membrane en bloc.
In each system, recruitment of these coating proteins is governed by a small GTPase: In the
COPII system Sar1p, in the clathrin system Arf1 and Arf6, and in the COPI system Arf1.
The minimal machinery to obtain generate vesicles in vitro has been defined in each system:
In the COPII system, minimal, empty cages comprised of Sec13/31 dimers were obtained in
the absence of Sar1p and Sec23/24 (Stagg et al., 2006). Clathrin coated vesicles could be
obtained from liposomes, in both the AP-1 and the AP-2 system using certain lipid
components (only for AP-1), tyrosine-based sorting signals as transmembrane component,
adaptor proteins and clathrin. The minimal machinery required for COPI vesicle budding is
comprised of p24-family members as transmembrane components, the small GTPase Arf1,
and the coating protein coatomer (Bremser et al., 1999).
In the various forms of the clathrin adaptor proteins, conformational changes are employed to
activate the protein coat for cargo binding and to trigger coat lattice formation. A distinct
conformational change in the coating protein coatomer has been also described in the COPI
system: Upon interaction with the dimeric form of the p24-family member p23, the subunit γ-
1. Introduction
22
COP changes its conformation (Reinhard et al., 1999). This conformational change may
represent a crucial step in COPI coat lattice formation, as the conformation of coatomer
polymerized by dimeric peptides of p23 corresponds to the one found on COPI vesicles
generated in vitro.
γ-COPs homologue in the AP-2 complex, the µ2-subunit, also undergoes a conformational
change during the budding process. Here, phosphorylation induces a conformational change
that opens a binding pocket for Yxxφ-based motifs (Matsui and Kirchhausen, 1990).
Interaction with these Yxxφ-motifs then induces a second conformational change in AP-2 that
modulates binding of AP-2 to other cargo proteins (Haucke and De Camilli, 1999). This event
was found to involve other subunits of the AP-2 complex, indicating that the conformational
change in the AP-2 system is not a local event in the µ2-subunit (Boll et al., 2002), but that
the whole adaptor protein complex changes into a membrane-bound form that is activated for
cargo binding.
The structural details of the conformational change in the µ2-subunit are relatively well
understood, and the crystal structure of the core of AP-2 has been solved. Only limited
information exists on the structural and functional details in the COPI system, as only a
fragment of one subunit of the coating protein, the trunk of γ-COP, has been crystallized to
date (Watson et al., 2004). And although the key steps in the COPI vesicle budding process
have been characterized, only little is known about the structural details and functional
dynamics of the coating protein coatomer in the formation of a COPI vesicle.
Aims of present work
1.3.
23
Aims of present work
In the present work, the structural dynamics of coatomer were addressed by:
1. Characterisation of the conformational change in coatomer using single-pair, singlemolecule FRET:
The conformational change in coatomer described by Reinhard (Reinhard et al., 1999) and
Bethune (Bethune et al., 2006) represents a key step in the formation of a COPI vesicle. The
polymerized, membrane bound form of coatomer undergoes a conformational change in
gamma-COP upon recruitment to the membrane, that is triggered by binding of members of
the p24-family, the transmembrane machinery found in COPI vesicles (Sohn et al., 1996).
In other coating systems, conformational changes are used specifically to activate the
complexes for cargo capture, or formation of the coat lattice (reviewed by (Langer et al.,
2007)). Here we ask if the conformational change induced in gamma-COP can be triggered
by all p24-family members, and if it is a local event or affects the conformation of other
subunits, i.e. the whole complex. To this end, we employed a combination of biochemical
methods and a new biophysical method that allows direct observation of one subunit of
individual coatomer complexes: Single-molecule, single-pair fluorescence resonance energy
transfer (FRET).
2. Cryo-electron microscopic studies on COPI vesicles:
The structures of a clathrin vesicles and COPII vesicles have been solved, and recently cryoEM studies have provided new insights into structural details and the architecture of these
vesicles. Compared to the other vesiculation systems, relatively little is known about the
architecture and structure of COPI vesicles, with only limited information obtained recently
(Donohoe et al., 2007). In the present work, we generated COPI vesicles in vitro from
purified Golgi membranes, coatomer and Arf1, and submitted these preparations to cryoelectron tomography.
2. Results
24
2.
Results
2.1.
Conformational dynamics of coatomer
2.1.1.
A conformational change in γ-COP: Screening p24-family
members
Previous studies revealed that the coatomer complex has multiple binding sites for its
different interaction partners (Bethune et al., 2006; Eugster et al., 2004; Sun et al., 2007a),
and that a conformational change is triggered in the γ-subunit upon interaction with dimeric
p23, a p24-family member (Reinhard et al., 1999). This conformational change causes
coatomer to shift into an unsoluble, membrane-associated conformation, that is also found
on COPI vesicles generated in vitro (Reinhard et al., 1999).
All members of the p24-family in their dimeric form bind to coatomer via two independent
binding sites in the trunk and the appendage domain of γ-COP (Bethune et al., 2006), in
contrast to the di-basic motif-containing cargo proteins that bind to α- and β'-COP (Eugster et
al., 2004). We therefore investigated if binding to coatomer of all the p24-family proteins in
either their monomeric or dimeric form would induce the conformational change observed
with p23, and if cargo proteins can also trigger the conformational change. The sequences of
the peptides used in this study are depicted in table 2.
Table 2: Sequences of peptides used in this study. They represent the cytoplasmic tails of
the p24-family members and of a model cargo protein, a subunit of the oligosaccharyl
transferase 48 (OST48). The specific signal sequences are highlighted: The di-aromatic motif
(FF, blue) and the dibasic-like motifs (red) in the p24-family members, and the KxKxx-ERretrieval signal in OST48. The corresponding dimeric peptides were generated by
introduction of an additional cystein at the C-terminus of the peptide, and mild oxidation.
2. Results
25
To this end, we made use of the previous finding that the conformational change in γ-COP
leads to precipitation of the complex, and that preciptated coatomer can be separated by
centrifugation from the conformationally unchanged complex (Reinhard et al., 1999).
Coatomer was incubated at increasing concentrations of the various monomeric and dimeric
peptides for 1h at room temperature, and centrifuged. The supernatants were discarded, and
the pellets analyzed by SDS-PAGE and Coomassie staining, as shown in Figure 6.
Only small amounts of coatomer are precipitated in the absence of peptides (lane 2), or in
the presence of monomeric p23 (lane 3) and monomeric p24 (lane 7). Dimeric p23 induced
polymerization of coatomer in a dose-dependent manner, as expected (lane 4-6, (Reinhard
et al., 1999)). Similarly, dimeric p24 caused precipitation dependent on its concentration
(lanes 8-10). In contrast, only marginal polymerization of coatomer is caused even by highest
concentrations of dimeric p25, p26 and p27 (lanes 11-13).
OST48, known not to bind to γ-COP but to α- and β'-COPs, did not cause significant
polymerization (lane 14).
Figure 6: Precipitation of coatomer by p24-family proteins. Rabbit liver coatomer was
incubated with peptides corresponding to the cytoplasmic tails of the proteins investigated, at
the concentrations indicated. After incubation at room temperature for 30min, the samples
were centrifuged and the pellets submitted to SDS-PAGE and stained by Coomassie.
2. Results
26
As a result, of the dimeric cytoplasmic domains of p24-family proteins that all bind to γ-COP
with similar affinity (Bethune et al., 2006), only p23 and p24 induce a conformational change
in coatomer, and none of the peptides cause a conformational change when added in their
monomeric form.
We now proceeded to ask whether this conformational change is a local event in γ-COP, or if
the whole complex changes its conformation upon recruitment to the donor membrane and
vesicle formation.
2.1.2.
Limited proteolysis and screening of other coatomer subunits
The conformational change in γ-COP induced by dimeric p23 was first analyzed using limited
proteolysis of the precipitated complex, showing an increased susceptability of a fragment of
γ-COP in the polymerized form, that was also found on COPI vesicles (Reinhard et al., 1999).
These findings imply that the conformational change in γ-COP, induced by dimeric
cytoplasmic domains of p23, is an event that takes place during vesicle formation.
Here we used a similar approach, digesting coatomer with the protease Thermolysine, and
screening the cleavage patterns of different subunits of coatomer. This allows screening of
cargo proteins and all p24-family members in their monomeric and dimeric forms in a single
set of experiments.
To this end, coatomer was incubated with both monomeric and dimeric peptides representing
the cytoplasmic tails of p23 and p24, and dimeric peptides of the respecitve tails of p25, p26
and p27, and monomeric OST48. After allowing for ligand binding, the samples were treated
with low concentrations (0.008µM) of thermolysine, and were incubated at room temperature
for the times indicated. The reaction was then stopped by addition of EDTA (40mM), and the
samples were submitted to SDS-PAGE and analyzed by Western blotting with an antibody
directed against α-COP (Figure 7).
Strikingly, the susceptability of α-COP to proteolysis by Thermolysine was only decreased in
the presence of both dimeric p23 and p24 (lanes 7,8 (p23d) and 11,12 (p24d)), as a doubleband at 150kDa is still present even after 2h of protease treatment (lanes 8 and 12). The
monomeric peptides of p23 and p24 were unable to alter the susceptibility of α-COP to
Thermolysine (lanes 5,6 (p23m) and 9,10 (p24m)). In addition, neither monomeric OST48
nor the other members of the p24-family members in their dimeric form were able to modify
the susceptability of α-COP for the protease (lanes 13-20).
2. Results
27
Figure 7: Limited proteolysis of coatomer. The complex was incubated with the peptides
indicated at a concentration of 25µM for dimeric peptides and 50µM for monomeric peptides,
and submitted to limited proteolysis by Thermolysine. The samples were then submitted to
SDS-PAGE and analysed by Western Blotting with an antibody directed against α-COP.
Thus, the susceptatibility of α-COP changes only in the presence of the two p24-family
members that are also capable of precipitating the complex in vitro. Interestingly, p23 and
p24 are the only two p24 proteins that are able to recruit Arf1-GDP to the membrane, forming
the priming complex in the early stages of COPI vesicle budding (Contreras et al., 2004;
Gommel et al., 2001).
To validate these findings, we used an independent method that allows direct observation of
the subunit investigated: Fluorescence resonance energy transfer (FRET). By selectively
monitoring the conformation of the labeled α-subunit, we can analyze if the observed
differential susceptability of α-COP is induced by γ-COP changing its conformation and
covering up or revealing domains of α-COP after the conformational change. To this end,
coatomer has to be specifically labeled on a single subunit with two fluorophores forming a
FRET-pair.
2.1.3.
Labeling of coatomer with fluorescent dyes
Various labeling approaches were considered to achieve specific labeling of one subunit of
coatomer:
Coatomer is a heptameric complex that cannot be recombinantly expressed in prokaryotic
expression systems. The individual subunits are not soluble by themselves, and only
oligomeric subcomplexes displayed limited solubility (α-β'-ε-COP), which were not suitable
for specific labeling.
A well-established approach that has been successfully employed to study the
conformational dynamics of proteins is directed mutagenesis of all cysteins in the target
protein, and the subsequent, site-specific introduction of two cysteines in different positions,
2. Results
28
followed by specific labeling of their sulfhydryl-groups by thiol-reactive derivatives of the
dyes. For coatomer, however, this approach is not feasible, as coatomer contains too many
cysteine-groups (human α-COP (P53621) contains 27 cysteine-residues).
We also tested an approach based on the reversible dissociation of the coatomer complex
(Pavel et al., 1998). Coatomer can be dissociated using an amine-reactive reagent, di-methyl
maleic acid anhydride (dmma). Dmma reacts with primary amines, forming an amide bond
and a free carboxylic acid moiety, thus "inverting" the charge at the position of the amino acid
with the primary amine. This leads to a step-wise dissociation of the complex, with level of
dissociation dependent on the excess of dmma employed. In addition, the dissociation was
found to be subunit-specific, i.e. subcomplexes split up in a characteristic pattern implying
differential reactivity to amine-reactive reagents.
The dissociation of coatomer did not yield satisfying amounts of soluble, dissociated
subunits. However, these findings lead us to examine amine-reactive derivatives of the
fluorophores in order to introduce the dyes into the coatomer complex.
2.1.4.
Labeling with amine-reactive dyes
To this end, we incubated the purified complex with N-Hydroxyl-succinimidyl-esterderivatives (NHS-esters) of the dyes Cy3 and Cy5. To ensure deprotonation of reactive εamino-groups we increased the pH to 8.0, and conducted the reaction at 0°C (ice water).
Labeled complex was separated from the free dye by a small gel filtration column (Sephadex
G-50, see materials and methods), purified by immunoprecipitation with an antibody specific
for native coatomer (CM1, (Palmer et al., 1993)) and submitted to SDS gel electrophoresis.
The labeling of individual subunits was detected by scanning of the gel using a confocal
setup with the corresponding lasers focused into the gel (see materials & methods). Figure 7
shows these fluorescence intensity traces of coatomer labeled with Cy3, purified by
immunoprecipitaion and submitted to SDS gel electrophoresis (6% gels).
The specifity of the reaction was found to be dependent on the concentration of dye reagent
used for the reaction. With a high concentration of labeling reagent, all large subunits of
coatomer were found to be labeled (Figure 8 D). The bands showing significant fluoresence
intensity correspond to α-, β-, β'- and γ-COP, with the relative degree of labeling dependent
on the concentration of labeling reagent used (Figure 8B-D).
2. Results
29
Figure 8: Analysis of coatomer after labeling with different concentrations of amine-reactive
derivatives of the fluorescent dye Cy3. The samples were purified by immunoprecipitation,
and submitted to SDS-PAGE (6%), and the fluorescence intensity traces were recorded upon
excitation at 532nm. A: 600pmol Cy3-NHS-ester. The single fluorescence peak coincides
with the position of α-COP. B: 1.5nmol Cy3-NHS-ester. Besides the prominent peak for αCOP, additional peaks in the fluorescence intensity trace are visible, running with the 100kDa
family of coatomer subunits (β-, β' and γ-COP). C: 8nmol Cy3-NHS-ester. Almost equel
relative labeling of α-COP and the 100kDa-family coatomer subunits. D: 1µmol Cy3-NHSester. The labeling of the 100kDa-family subunits exceeds α-COP.
Using the conditions that were used to label coatomer scanned in Figure 8A, specific labeling
of α-COP was achieved. Figure 9 shows a coatomer sample after labeling with Cy3 and Cy5,
purification by immunoprecipitation with CM1 (Palmer et al., 1993), subsequent separation of
the subunits by SDS gel electrophoresis (6-15% gradient gel), and staining with Coomassie.
Bands representing COPs are indicated with the corresponding greek letters. Additional
bands are impurities, mainly introduced by the antibodies used for immunoprecipitation.
2. Results
30
Fluorescence intensity traces of the lanes corresponding to the Cy3- and Cy5-emission
channels, detected by scanning with a confocal setup (see Materials and Methods), are
shown in Figure 9 C and D, respectively. The fluorescence intensity shows a prominent peak
in both channels at a molecular mass corresponding to alpha-COP.
Figure 9: Analysis of coatomer after labeling with amine-reactive derivatives of the
fluorescent dyes Cy3 and Cy5. A: Coomassie stained SDS-PAGE gel of Cy3-Cy5-labeled
coatomer purified by immunoprecipitation with CM1. B: Coomassie intensity trace, individual
subunits are inidcated. C: Cy3 fluorescence (excitation 532nm) and (D) Cy5-fluorescence
(excitation at 635nm) intensity of labeled coatomer separated by SDS-PAGE and purified by
immunoprecipitation with CM1. (The unlabeled bands at ~50kDa and ~25kDa represent IgG
heavy and light chains, respectively).
Total fluorescence emission of the protein bands was calculated for the alpha-subunit and
the beta, beta' and gamma-subunits. An average of 96% of the emission in the Cy3 channel
and 90% of the emission in the Cy5 channel results from the protein band corresponding to
the α-subunit. Thus, almost exclusive labeling of one subunit within coatomer, namely alpha-
2. Results
31
COP, was achieved. The labeling efficiency was estimated as described in Materials and
Methods, and found to be about one Cy3 and one Cy5 dye per coatomer.
2.1.5.
Functionality of labeled coatomer
As described above, coatomer was specifically labeled on its α-subunit. Next, we
investigated whether labeled coatomer retains its functionality, i.e. its capability to bind to
ligands and membranes, and to form COPI vesicles in vitro.
First, binding of cytoplasmic tail domains of membrane proteins to the labeled complex was
investigated in pull-down experiments. To investigate if both the γ-COP binding sites for p24family members and the α- / β'-COP cargo binding sites retained their capability to bind to
their binding partners after labeling, labeled and unlabeled coatomer was incubated with
immobilized fusion proteins harbouring the cytoplasmic domains of OST48 or dimeric p23.
Bound coatomer was recovered, submitted to SDS gel electrophoresis and analyzed by
Western blotting with antibodies directed against α-COP and δ-COP. As shown in Figure 10,
both labeled and unlabeled coatomer show similar binding to these domains (compare lanes
5 and 6, and lanes 7 and 8), and no unspecific binding of labeled or unlabeled coatomer was
observed (lanes 3 and 4).
In order to analyze whether bound coatomer is in fact labeled, the blot was analyzed with
antibodies directed against the dyes Cy3 and Cy5. Coatomer is labeled with the dyes
exclusively on the alpha-subunit as shown in Figure 10, lanes 6 and 8. The remarkable
specificity of the reaction is visualized by comparison of the input crude coatomer preparation
after labeling (Figure 10, lane 2) with the bound samples (Figure 10, lanes 6 and 8), where a
striking loss of non-coatomer protein is observed.
2. Results
32
Figure 10: Functionality of labeled coatomer: Binding to the cytoplasmic domains of p23 and
OST48. His6-constructs of the cytoplasmic tails of p23d and OST48, or a His6-construct as a
control, were immobilized on Ni-Sepharose and incubated with identical amounts of labeled
or unlabeled coatomer, and then washed. Subsequently, bound coatomer was eluted with
SDS sample buffer, subjected to SDS-PAGE and detected by Western blotting (antibodies
directed against α-COP, γ-COP and Cy3/Cy5).
Second, functionality of labeled coatomer was evaluated by analysing its ability to bind to
Golgi membranes. This process depends on prior binding of the small GTP binding protein
Arf1 in its active GTP-form to these membranes, and thus is GTP-specific. Partially purified,
labeled or unlabeled complex was incubated with Golgi-enriched membranes and Arf1 in the
presence or absence of GTPγS (a non-hydrolyzable analogue of GTP). Thereafter, the
membranes were recovered by centrifugation, and the amount of coatomer bound was
estimated using SDS-PAGE and Western blotting with antibodies directed against alphaCOP. The amount of membranes loaded was controlled by the presence of the membrane
protein p24. Figure 11 shows that similar amounts of labeled and unlabeled coatomer are
recruited to Golgi membranes (comparing alpha-COP in lanes 6 and 7), and that this
recruitment is dependent on the presence of Arf1 in its active, GTP-bound form (compare
lanes 4 and 5 with 6 and 7). No labeled or unlabeled coatomer was found in the pellet in the
absence of Golgi membranes (Figure 11, lanes 8 and 9).
2. Results
33
Figure 11: Functionality of labeled coatomer: Membrane binding assay. Identical amounts of
labeled or unlabeled coatomer were incubated with purified Golgi membranes and Arf1 in the
presence or absence of GTPγS. Subsequently, Golgi membranes were centrifuged and
subjected to SDS-PAGE and immunoblotting (with antibodies directed against α-COP, p24 or
the dyes Cy3/Cy5, as indicated).
The samples were redecorated with an antibody directed against the fluorescent dyes, to test
whether coatomer bound to Golgi membranes (shown in Figure 11) is also labeled. Labeling
was detected, and found to be exclusively on the α-subunit of coatomer, as shown in Figure
11, lane 7. The multiple bands visible in the crude input (Figure 11, lane 3;) are degradation
products and other, contaminating proteins that are also labeled with Cy3 and Cy5.
Third, the capability of the labeled coatomer complex to support COPI vesicle formation was
investigated. To this end, Cy3/Cy5-labeled coatomer was added to an in vitro COPI budding
assay with purified Golgi membranes, Arf1 and GTPγS. A vesicle fraction was purified by
differential centrifugation as described in Materials and Methods, and its protein content
analyzed by SDS-PAGE and Western blotting. As shown in Figure 12, the coat protein
pattern typical of COPI vesicles is obtained in the sample incubated with GTPγS, harvested
from a 47% sucrose cushion. Decoration with anti-Cy3/Cy5 antibodies reveals a band that
migrates with α-COP (lane 2). No detectable amounts of coatomer are found in a control
fraction from a sample incubated in the absence of GTPγS.
2. Results
34
Figure 12: Functionality of labeled coatomer: In vitro formation of COP vesicles. Golgi
membranes were incubated with Arf1 and labeled coatomer in the presence and absence of
GTPγS. After centrifugation, the fractions containing purified vesicles were submitted to SDSPAGE and analyzed by Western blotting with antibodies directed against α-COP, δ-COP,
Arf1 and Cy3/Cy5, as indicated.
From these analyses we conclude that the functionality of coatomer is not compromised by
the labeling process, and that functional coatomer bears the dyes on its α-subunit.
2.1.6.
Specific immobilization of labeled coatomer
Initial FRET experiments involving ensemble measurements showed that, in order to
investigate the conformational dynamics of coatomer, we had to design a single-moleculesensitive assay. This new assay was designed to overcome the following three issues:
1. Coatomer precipitates upon interaction with dimeric p23 p24. As shown previously,
the conformational change induced in γ-COP by dimeric p23 and p24 leads to
precipitation of coatomer. This effectively excludes measurements in solution, as the
complex both aggregates and is removed from the measurable pool of soluble
proteins.
2. The preparation of coatomer from rabbit liver contains 10-30% of other contaminating
proteins, that may also be tagged with a dye as described in the labeling procedures.
These contaminations have to be removed prior to measurements.
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35
3. The labeling approach described above generates a heterogenous population of
labeled coatomer, with populations of coatomer carrying a varying number of dyes.
The new assay must be able to specifically select singly donor- and acceptor-labeled
coatomer.
Thus, a surface-based, single-molecule-sensitive assay using a scanning confocal
microscope was developed.
A surface-based assay also allows monitoring of the conformational dynamics of individual,
tethered coatomer complexes, as immobilization of coatomer effectively prevents
precipitation and aggregation of the complex.
The surface-based assay also allows screening of individual labeled complexes, and
selection of singly-donor and –acceptor labeled complexes by evaluating the number of
attached dyes by specifically illuminating individual spots and assessing their fluorescene
intensity trace over time. Such a selection is stringently required since the statistical labeling
process used here generates coatomer molecules labeled at a 1:1 stoichiometry for Cy3 and
Cy5, without excluding complexes labeled multiple times, as mentioned before.
Subsequently, the efficiency of the FRET signal can be calculated for each, singly-donor and
singly-acceptor labeled coatomer complex. After sorting into a histogram, this allows
monitoring of individual sub-populations of conformationally changed and unchanged
complexes. In ensemble measurements, the read-out is always the averaged value for all
different populations present in the measured sample. In contrast, in the setup described
here, we can identify individual coatomer populations that have or have not undergone a
conformational change.
Thus, as a first step, specific recruitment of coatomer from solution to a glass surface had to
be established.
To ensure efficiency and specificity of this immobilization, we chose an antibody that binds
native coatomer, CM1 (Palmer et al., 1993). This antibody is a monoclonal antibody available
from hybridoma cell line supernatant, thus being first choice in both specificity and
availability.
As a bulk protein to quench unspecific adsorption of coatomer and other contaminating
proteins to the surface, we chose bovine serum albumin (BSA). Previous studies in our lab
showed that surfaces incubated with specific coatomer binding partners and passivated with
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BSA can be used to specifically recruit coatomer in an ELISA-like assay (Bethune et al.,
2006).
The following criteria have to be addressed for the single-molecule-sensitive FRET studies
we aimed at:
-
The individual, labeled coatomer complexes must be spaced sufficiently far apart, so
that their emission spots (point spread functions, PSF) do not overlap, to allow
selection of individual complexes for subsequent aquisition of their fluorescence
intensity traces over time for FRET efficiency calculation.
-
A sufficient number of labeled coatomer complexes must be aquired in a typical
scanning area of 20µm x 20µm for efficient data aquisition.
In the present study, two approaches were investigated to accomplish efficient and specific
recruitment of coatomer to a glass surface, and retain functionality of the immobilised
complex.
Figure 13: Immobilisation strategies to specifically tether labeled coatomer to a glass
surface. BSA was chosen as a bulk protein to prevent unspecific adsorption to the surface.
Both approaches are based on the monoclonal antibody CM1 that recognizes assembled
coatomer only.
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37
A. Biotinylated biotin-based immobilisation of coatomer using CM1
The first approach was based on a sandwich-concept (see figure 13A), with biotinylated BSA
as an anchor function on the surface. The advantage of this method is that, with a mixture of
biotinylated BSA and normal BSA forming a relatively homogenous layer on the glass
surface, unspecific adsorption to the surface itself should be minimized.
To immobilize the CM1 antibody on this surface, recombinant streptavidin and biotinylated
Protein A was applied to the surface, followed by a solution of CM1 antibody purified from the
hybridoma cell line supernatant by Protein A affinity purification.
Subsequently, titration experiments were performed to find conditions to specifically recruit
labeled coatomer. Here, however, increasing unspecific adsorption to the sandwich
construction was detected upon additon of each, further component. After addition of
relatively small concentrations of biotinylated Protein A, adsorption of labeled protein was
detected even in the absence of CM1. Also, using different buffers (with higher salt
concentrations to quench electrostatic interactions), detergents (up to 1% NP-40) and
lowering of the density of the anchor functions did not quench unspecific adsorption.
Thus, the second approach was tested.
B. Direct, antibody-based immibilization using CM1
The second approach utilizes the same central anchor function to specifically immobilize
coatomer, the antibody CM1. The antibody was purified by Protein A affinity purification and
subsequently directly applied to the activated glass surface. To quench unspecific adsorption
to the surface, the purified antibody was administered in a solution containing excess bulk
protein, BSA.
To find conditions suitable for single-molecule-sensitive FRET experiments as described
above, various concentrations of antibody and of coatomer administered to the surface were
screened. As a starting value for the anchor functions, a concentration of antibody was
chosen that had shown good results for biotinylated BSA (data not shown).
To control for unspecific adsorption to an antibody-BSA surface, an antibody directed against
an unrelated protein, the Okt8 antibody directed against the CD8 receptor, was applied in a
parallel set of experiments. To increase the stringency of the system, high-salt phosphatebased buffer and washing steps with buffers containing 0.5% NP-40 were used (see
Materials and Methods). As previous studies showed that the ζ-subunit of coatomer partially
dissociates from the coatomer complex if exposed to 1% of NP-40 for more than 30 minutes,
only a short incubation (10min) with buffer containing 0.5% NP-40 was chosen.
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Figure 14: Low intensity surface scans (20µm x 20µm) of glass surfaces coated with a BSAantibody solution. A: Scanned area of a BSA-CM1 surface incubated with 3ng of Cy3-labeled
coatomer. B: BSA-Okt8 surface after administration of 3ng of Cy3-labeled coatomer. C: BSACM1 surface incubated with a sample depleted of coatomer. D: Quantification of surface
densities for the described conditions (n=10).
Figure 14 shows a set of experiments with identical amounts of labeled coatomer applied to
a BSA – CM1 surface. Unspecific adsorption was investigated by applying labeled coatomer
to a BSA – Okt8 surface in identical conditions.
Labeled coatomer was detected by low intensity fluorescence scans of 20µmx20µm areas on
the antibody-BSA surfaces as shown in Figure 14A-C. With the CM1 antibody, 86±13 (SE,
n=10) spots can be observed in one scanning area (Figure 14A).
Unspecific adsorption was quantified by adding an identical concentration of Cy3-labeled
coatomer to a surface incubated with an unrelated antibody (Okt8, specific for the protein
CD8; 21±8 spots (SE, n=10), Figure 14B).
In order to test for unspecific adsorption of contaminating proteins in the coatomer
preparations to the CM1-antibody surface, a sample of labeled coatomer was depleted of
coatomer by three successive immunoprecipitations with an antibody directed against the αand γ-subunits according to an established protocol (Wegmann et al., 2004). The coatomerdepleted supernatant, representing contaminating, labeled proteins in the coatomer
2. Results
39
preparation, was administered to a CM1 surface in conditions identical to those used above.
In a 20µmx20µm area, 19±8 (SE, n=10) spots were detected (Figure 14C).
These results are quantified in Figure 14D, and show that coatomer can be specifically
immobilized on a glass surface covered with an antibody-BSA mixture using the antibody
CM1.
The conditions depicted in figure 14 were chosen for subsequent experiments, as both a
sufficient number of spots was detected per scanning area, and the spots were separated far
enough to allow selection of individual, labeled complexes.
2.1.7.
Surfaces with Cy3-Cy5-labeled coatomer
Next, coatomer labeled with both Cy3 and Cy5 was immobilised on CM1-BSA surfaces
prepared as described above. Figure 15 shows low-intensity surface scans of 20µmx20µm
areas, excited with a 532nm laser (donor only).
Figure 15: Low intensity surface scans (20µm x 20µm) of glass surfaces coated with a BSAantibody solution and incubated with Cy3-Cy5-labeled coatomer. Samples were excited with
a 532nm Nd:YAG laser, and FRET detected by acceptor emission (donor emission imaged in
green, acceptor emission imaged in red).
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40
About 30% of the spots show FRET, i.e. emission in the acceptor channel upon donor
excitation, as depicted in Figure 15 (donor channel emission: green; acceptor channel
emission: red). Different populations are clearly visible, with spots showing lower (yellow)
and higher energy transfer efficiency (red).
2.1.8.
Inter- or intramolecular FRET?
The detected FRET signal could derive from either intramolecular FRET or intermolecular
FRET between two adjacent, labeled coatomer molecules. In order to discern intra- from
intermolecular FRET, two separate labeling reactions were performed: One sample of
coatomer was labeled with Cy3 only, and another sample was labeled with Cy5 only.
Subsequently, a BSA-CM1 surface was incubated with a 1:1 mixture of these labeled
coatomer preparations, and examined by low-intensity surface scans. As shown in Figure
16A, no emission was observed in the acceptor channel upon donor excitation. The
presence of Cy5-labeled coatomer on these surfaces was visualized by direct acceptor
excitation (Figure 16B).
In addition, no intermolecular FRET by aggregation was detected after addition of dimeric
p23 (Figure 16C, acceptor presence was confirmed by direct excitation, shown in Figure
16D).
Figure 16: Low intensity surface scans (20µm x 20µm) of glass surfaces coated with a BSAantibody solution and incubated with a 1:1 mixture of Cy3-labeled coatomer and Cy5-labeled
coatomer. Samples were illuminated with 532nm for Cy3 excitation, and with 635nm for Cy5
excitation. No acceptor emission was detectable upon donor excitation both prior to and after
addition of dimeric p23d (A and C), and the presence of Cy5-labeled coatomer was controlled
by direct acceptor excitation (B and D).
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From these data, we conclude that the acceptor emission observed for Cy3-Cy5-labeled
coatomer is due to intramolecular energy transfer.
2.1.9.
Assessing the number of attached dyes
To assess the number of attached dyes, the spots showing emission in the acceptor channel
upon donor excitation were further analyzed by monitoring their fluorescence intensity traces
over time. To this end, these spots were selected and photo-bleached by continuous
illumination at 532nm, and their bleaching pattern, i.e. their fluorescence intensity trace over
time, was recorded. For this analysis, only spots showing acceptor emission upon donor
excitation were picked, as shown by the selected spots in Figure 17A and B (for details, see
Materials and Methods).
Figure 17: Single-pair FRET in coatomer. A: Low intensity surface scan (20µm x 20µm) of a
glass surface coated with a BSA-antibody solution and incubated with Cy3-Cy5-labeled
coatomer. B: Selection of spots (numbered circles) picked for subsequent recording of
fluorescence intensity trace (see Materials and Methods for selection criteria). C and D:
Representative fluorescence intensity traces of spots, as used for data acquisition and
evaluation. Intensity traces showing a clear bleaching pattern, indicating singly donor- and
acceptor-labeled molecules, with a simultaneous increase in donor emission upon acceptor
bleaching, and single bleaching events in both the donor and the acceptor channel, as in B,
2. Results
42
were further analyzed. The trace depicted in D shows multiple intensity changes after 1.5, 3,
and 4.8 seconds indicating labeling with more than one Cy3 fluorophor.
Figure 17 shows typical fluorescence intensity traces recorded in these experiments. Traces
were used for subsequent energy transfer efficiency calculation only if they fulfilled the
following criteria:
1. A single bleaching event in both the donor and the acceptor channel, leading to a
drop to background levels in single step.
2. An increase in donor emission upon acceptor photobleaching.
Figure 17C shows a single bleaching event in the acceptor channel, with a simultaneous
increase in donor emission upon acceptor photobleaching. Subsequently, donor emission
drops to background level in a single step, thus indicating single donor and acceptor labeling.
Figure 17D displays a fluorescence intensity trace with multiple bleaching steps after 1.5, 3
and 4.8 seconds, indicating multiple labeling of the complex. These traces were discarded.
Thus, the fraction of coatomer labeled with multiple donor and acceptor dye molecules is
minimized in our FRET statistics.
2.1.10. Approximating the FRET efficiency Eapp.
The FRET efficiency was approximated by evaluating the relative emission intensities in the
donor and acceptor channel, using a ratiometric approach:
Eapp = EA / EA + ED
(see Materials and Methods, and (Deniz et al., 2001; Ha, 2001)). To this end, the following
data were extracted from the fluorescence intensity traces: The total emission in the donor
channel, the total emission in the acceptor channel, and background emission. Only traces
with more than 200 photons in both channels were kept for subsequent analysis.
After correction for crosstalk, the FRET efficiency was calculated for each fluorescence
intensity trace as described in Materials and Methods. Each complex was subsequently
sorted into a normalized histogram.
The calculated FRET efficiencies for CM1-immobilized, Cy3-Cy5-labeled coatomer are
summarized in Figure 18 (n=216). A major population with an energy transfer efficiency of
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Eapp = 55-70% can be clearly discerned, with its center at 63%, as determined by fitting a
Gaussian function to the data (see Materials and Methods).
Figure 18: Normalized histogram for Cy3-Cy5-labeled coatomer, no peptide added (n= 214,
class size: 5%). A major population can be identified, centered at a FRET efficiency of 63%.
2.1.11. Interaction of coatomer with cytoplasmic tail domains of ligand
proteins
In order to investigate the interaction of coatomer with p23 and OST48, the FRET efficiencies
of the labeled complex were analyzed after addition of synthetic peptides resembling the
cytoplasmic tail domains of these proteins. Both ligands were used at a concentration of
20µM, i.e. well above the KD and below saturation (Bethune et al., 2006).
First, the interaction of coatomer with OST48 was investigated. OST48 is a cargo protein that
binds as a monomer to the predicted β-propeller domains on the α- and β'-subunits of
coatomer (Eugster et al., 2004). The corresponding peptide was added and the FRET
efficiencies of individual complexes were analyzed. As depicted in Figure 19, a major
population is found that displays an energy transfer efficiency centred at 67%, with a
broadened peak. Thus, after addition of OST48, the centre of the population has shifted
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44
slightly from 63% to 67%, as seen in the overlay shown in Figure 19B. This shift is within the
resolution limits of the method used.
Figure 19, A: Normalized histogram for Cy3-Cy5-labeled coatomer, after addition of 20µM
OST48 (n= 138, class size: 5%). The major population is centered at a FRET efficiency of
67%. B: Overlay of normalised histograms prior to and after addition of OST48.
Next, the interaction of coatomer with p23 was analyzed by adding a dimeric peptide
representing the cytoplasmic tail of p23 to the surface.
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45
Figure 20, A: Normalized histogram for Cy3-Cy5-labeled coatomer, after addition of 20µM
p23d (n= 176, class size: 5%). The major population is now centered at a FRET efficiency of
84%, a subpopulation centered at an efficiency of 59% was also detected. B: Overlay of
normalised histograms prior to and after addition of dimeric p23.
A major population with a FRET efficiency peak at 84% was found, as depicted in Figure
20A. A subpopulation was also detected, with a peak centred at 59%, which may represent
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46
coatomer with no p23 bound to the complex and an unchanged conformation (overlay in
Figure 20B).
Hence, a striking shift of the FRET efficiency of the major population of single-pair FRET
labeled coatomer molecules from 63% to 84% is observed upon binding of coatomer to
dimeric p23, demonstrating that a conformational change induced in the trunk domain of γCOP is transmitted to the α-subunit of the complex. This is of special interest, as no
functional interaction between α-COP and γ-COP has been described to date.
The observed conformational change takes place in a subunit of coatomer that is competent
to bind KKxx- and KxKxx-based ER retrieval signals. In the Clathrin system, cargo binding of
the Adaptor complexes is tightly controlled by a series of conformational changes induced by
phosphorylation and cargo binding (reviewed by (Langer et al., 2007)).
Thus, we investigated if binding of coatomer to proteins bearing the ER retrieval signal
sequences is modulated by the conformational change induced by dimeric p23. To this end,
we made use of an ELISA-like binding assay, to determine if both the KD of coatomer for its
protein binding partners and/or the number of binding sites changes during the
conformational change (Bethune et al., 2006).
2.1.12. ELISA-like binding assay
In this assay, coatomer is immobilised on Protein A ocated 96-well plates using an antibody
(CM1), and incubated with different concentrations of S-tagged fusion proteins of its binding
partner. The amount of protein bound to coatomer is determined using an S-tag-specific
detection system.
The construct used as a representative cargo protein here is comprised of the cytoplasmic
domain of OST48 containing a KxKxx ER retrieval signal (see also table 2), of a His6-tag for
purification, and of an S-tag for detection.
First, we determined the KD of coatomer for this protein by a series of independent
experiments (KD=0.35µM, SD=0.1µM). This is in excellent agreement with published data
(0.9µM, (Bethune et al., 2006)).
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Next, we perfomed a series of independent experiments to determine the KD of this fusion
protein in the absence and presence of dimeric p23. The binding curves are shown in Figure
21.
The KDs were calculated to be 0.34 µM and 1.2 µM. So, surprisingly, a slight increase in the
KD was observed upon addition of dimeric p23. Also, the total number of binding sites did not
change significantly.
Figure 21: ELISA binding curves to monitor the KD and the total number of available binding
sites of coatomer for cargo proteins in the presence and absence of dimeric p23. The tables
contain data of two sets (+/- p23d) of 4 independent experiments.
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2.1.13. Rare events
Some fluorescence intensity traces showed striking dynamics in their intensity profile. In rare
events, a fluctuation between two discrete emission states was observed. Figure 22 shows
examples of these fluorescence intensity traces. The two states displayed in Figure 22
correspond to FRET effiencies of 84% and 55%, thus representing the efficiencies found for
conformationally changed and unchanged coatomer.
Figure 22: Fluorescence intensity trace of a Cy3-Cy5-labeled coatomer that oscillates
between two FRET states. The calculated FRET efficiencies are listed in the bottom.
This fluctuation may indicate that during aquisition of the fluorescence intensity trace, the
complex underwent the conformational change (until 2.1s), reverted back to the unchanged
conformation (2.1s – 3.2s), and changed again (3.2s – 5.0s).
However, this live monitoring of the conformational dynamics was only rarely observed
(1/100 spots).
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2.2.
49
Electron Microscopic investigation of COPI vesicles
The remarkable functional dynamics of coatomer observed in the experiments described
raised the question as to the structure(s) of the heptameric complex after polymerization.
Therefore, in a second project, we investigated the structure of COPI vesicles prepared in
vitro using cryo-electron tomography. In contrast to the other vesiculation systems, where the
cage structures have been characterised at the molecular level (Cheng et al., 2007; Collins
et al., 2002; Fath et al., 2007; Stagg et al., 2006), only limited data is availbale on the
structure of the COPI coat. Here we tested different experimental procedures that allow
generation and purification of large amounts of COPI vesicles in vitro, and submitted them to
cryo-electron tomography experiments.
2.2.1.
Preparation of COPI vesicles in vitro
For initial screening experiments, the yield and purity of different COPI vesicle preperation
assays were investigated. Three different methods to prepare COPI vesicles in vitro were
tested:
The first assay follows the protocol described by Pavel et al. (Pavel et al., 1998). Using the
non-hydrolysable GTP-analog GTPγS, purified Golgi membranes were incubated with cytosol
as a source for coatomer and Arf1. After release of the vesicles using a high-salt buffer, the
samples were applied on top of a sucrose gradient ranging from 50% to 20%. After
centrifugation to equilibrium for 17h at 100000g, the fractions containing the COPI vesicles
were harvested. Typically, this fraction displayed a buoyant density of 38-42% sucrose.
Figure 23 shows electron microscopic images of samples harvested from these gradients,
and submitted to negative staining using 1% uranyl acetate (images taken on a FEI CM120
electron microscope, 120kV, defocus 2µm). The vesicles display a diameter between 60nm
and 80nm, which is in excellent agreement with data published previously (Orci et al., 1998).
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Figure 23: Images of GTPγS-COPI vesicles, purified by sucrose gradient and submitted to
negative stain. Magnification: 16kx, scalebar: 500nm.
To increase both the stability of the coat and the amount of coated vesicles that can be
purified, different chemical crosslinkers were tested. Addition of both formaldehyde and
glutaraldehyde didnt increase the yield of vesicles from a COPI budding assay, as found in
both negative stain electron microscopy and Western Blotting (data not shown).
In parallel, a second experimental approach was tested. Here, the individual components
required for COPI vesicle budding were added separately, and as recombinant or purified
proteins. This approach offers the advantages that less contaminating proteins (as opposed
to using cytosol as a source) are present, and that stable GTP-coated vesicles can be
isolated, because no ArfGAP proteins are present (see introduction; soluble ArfGAP proteins
present in the cytosol mediate uncoating of COPI vesicles). In addition, the final purification
was achieved using a short centrifugation of 1h onto a sucrose cushion with a concentration
of 47% sucrose, with an upper cushion of 37%, significantly lowering the preparation time
(see Materials and Methods, and (Beck et al., 2008)).
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Figure 24: Images of GTPγS-COPI vesicles purified either via a 17h-gradient (A-C) or
centrifugation on a sucrose cushion (D-F). Magnification 16kx, scalebar 500nm.
In initial screening experiments, to screen if the purity and yield of vesicles from this assay
was comparable to the Pavel method using a 17h centrifugation, GTPγS was employed.
Figure 24 shows images of vesicles prepared in both methods (figure 24A-C: 17h gradient
centrifugaiton, figure 24D-F: 1h cushion centrifugation). The vesicle preparation shown in
figure 24D-F contains minor membrane fragments, but a homogenous pool of COPI vesicles
can be clearly identified.
The samples in Figure 24 were prepared using 0.25mg Golgi. To increase the overall yield of
vesicles, the amount of all components was increased by a factor of 10. The fractions
obtained in these experiments were submitted to negative staining experiments. Figure 25
shows images taken of samples diluted 1:1 prior to application to the grid. Clearly, a
homogenous population of vesicles with a diameter of 60-80nm can be discerned.
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Figure 25: Images of COPI vesicles (upscaled sample, 2.5mg of Golgi) prepared with
GTPγS, purified by centrifugation on a sucrose cushion, and submitted to negative stain.
Magnification 19kx, scalebar 500nm.
Subsequently, COPI vesicles were prepared using GTP instead of GTPγS. Identical
conditions as described above were used, and the fraction containing vesicles were applied
to grids, submitted to negative stain and imaged. Again, COPI vesicles were detected,
although the yield is lower by a factor of 5 as compared to GTPγS preparation. These
fractions were taken for subsequent EPON embedding and for cryo electron microscopic
experiments.
2.2.2.
Embedding of chemically fixed vesicles into a matrix
Subsequently, we decided to screen and investigate the morphology of COPI vesicles
prepared in vitro in a system that does not involve adsorption to a carbon backing. To this
end, samples prepared as described above were subsequently embedded in a polymer
matrix. The fraction containing vesicles was fixed with 1% glutaraldehyde and stained with
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53
Ruthenium-Red. After sedimentation of the vesicles by a 100000g centrifugation, the pellet
was stained with OsO4 and embedded in EPON.
Figure 26A and 26B show images of thin sections of these samples. Vesicles were found
with diameters of 60nm to 80nm, which is in excellent agreement with the values estimated
from the negative sain samples and data published previously (Orci et al., 1998). In these
samples, donor membranes that sedimented in the centrifugation with the vesicles are also
present (Figure 26). Clearly, the vesicles do not exhibit a round shape, but exhibit "edges"
and "corners", i.e. the rim of the coat displays short spans with high curvature (marked with
an asterisk in Figure 26B, and long spans with low or without curvature (marked by bars in
Figure 26B).
Figure 26: Images of COPI vesicles prepared with GTP in vitro, embedded in a polymer
matrix and stained with Ruthenium Red and OsO4. Magnification 50k, scalebars 100nm.
2.2.3.
Cryo-electron microscopy of COPI vesicles generated in vitro
Samples of these fractions were mixed with gold particles as fiducials (10nm diameter),
applied to Quantifoil grids, and snap-frozen in liquid ethane. Figure 27 shows images of
these grids, taken at 300kV in a FEI Polara electron microscope with a Gatan stage. The
samples exhibited extremely low contrast, as well as only a very limited number of particles
in the vitreous ice. The low contrast does not allow clear identification of any vesicles,
although particles with the right diameter may be present in solution (figure 27, marked with
arrows). In addition, the vitreous ice displayed only a very limited stability in the beam.
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Figure 27: Images of COPI vesicles (upscaled sample, 2.5mg of Golgi) prepared with
GTPγS, purified by centrifugation on a sucrose cushion, and submitted cryo electron
microscopy. Magnification 16kx, scalebar 500nm.
Both issues may be primarily caused by the high sucrose concentration of the sample (ca.
42%). If the concentration of sucrose was lowered by dilution (1:8) to obtain a sucrose
concentration of ca. 5%, no more particles were detected in the ice. However, the ice
displayed better stability in the beam, and boiling occured only after application of a much
higher electron dose.
Also, after dialysis of the sample against buffer without sucrose, no particles could be
detected in the sample after application to a Quantifoil grid and cryo-EM imaging. To test the
concentration of vesicles, the dialysed samples were applied to carbon-backed grids and
submitted to negative staining. Here, particles could be detected, although the number of
vesicles was decreased by a factor of ~10 (data not shown)
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2.2.4.
55
Using "backed"-Quantifoil
In the experiments described previously, the COPI vesicles showed significant adhesion to
the carbon film in negative stain preparation. The density of vesicles found in these
experiments indicated that a large quantity of vesicles is recruited to the carbon backing.
Thus we investigated if we could make use of this behaviour for the cryo-EM experiments.
To this end, we applied thin films of two backing materials to standard quantifoil grids: First,
we used thin carbon films (see Materials and Methods) with a thickness of ~40A. Second, we
employed thin TiSi films (see Materials and Methods, and (Rhinow and Kuhlbrandt, 2008)).
The advantage of these films is higher stability, and a lower refractive index as compared to
standard carbon films. In the following experiments, however, we used carbon-backed
Quantifoil, as this backing showed higher adsoprtion of COPI vesicles (data not shown).
In addition, using backed Quantioil, the sample can be repeatedly applied to the grid, leading
to an increase in vesicle density on the backing. Furthermore, the grids were washed with
buffer to reduce the sucrose concentration.
Figure 28: Low (A) and high (B) magnification Images of COPI vesicles, deposited on a
carbon-backed Quantifoil grid, embedded in vitreous ice. Scalebars as indicated.
Figure 28 shows 2 images of carbon-backed Quantifoil, with identical samples applied three
times and subsequent washing with buffer (2x). As seen in the low-magnification image, a
number of particles with the dimensions of COPI vesicles can be identified in each Quantifoil-
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56
hole (figure 28A, indicated with arrows). However, contrast was limited again, as shown in
the high-magnification images (figure 28B).
Figure 29: Low (A) and high magnification (B) Images of COPI vesicles with 6nm gold dots
as ficucials deposited on a carbon-backed Quantifoil grid, and embedded in vitroues ice.
Scalebars as indicated.
In Figure 29, gold particles with a diameter of 6nm were added to the sample prior to
application to the grid as high contrast markers for alignment of images in a tilt series. In
these conditions, the fiducials cluster around the vesicles (see Figure 29A and 29B). This
greatly faciliated detection of the vesicles in the ice, but also blocked the protein coat from
view.
To prevent fiducial clustering, the carbon-backed Quantifoil grids were pre-incubated with the
fiducials, and the fiducials left to dry on the film. Subsequently, the samples were applied to
the grid, and treated as described above.
Tomograms were recorded of the particles immobilised on these carbon films. Single-axis tilt
series, ranging from angles of -60° to +60°, were recorded, and reconstructed using the
IMOD software package (Kremer et al., 1996). Figure 30 shows a slice through a tomogram,
with an enlarged view of a particle immobilised on the carbon film.
A particle with the right dimensions can be identified (Figure 30). The vesicle seems to be
comprised of an inner and an outer layer. However, contrast is very limited, and no structural
details can be discerned.
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Figure 30: Section of a tomogram of a sample of purified GTPγS-COPI vesicles deposited on
a thin carbon film, embedded in vitreous ice (scale bars: 100nm). A particle with a diameter of
approximately 60nm can be discerned (marked with an arrow). The particle seems to be
composed of an outer and an inner layer Magnification 26kx.
In addition, the particles did not exhibit a uniform, symmetrical form. This may be due to
distortion caused by adhesion to the carbon film, or osmotic stress during the dilution and
washing steps.
2.2.5.
Direct preparation and imaging of COPI vesicles.
In the previous experiments, the detection and imaging of the COPI vesicles was severly
impeded by the high sucrose concentration in the product fraction: The samples exhibited
low contrast, and only limited stability in the electron beam. Thus, in a parallel approach, a
protocol without a sucrose gradient-based purification was tested.
To this end, the components required for COPI vesicle biogenesis required in vitro (saltwashed Golgi membranes, Arf1, ARNO and coatomer) were incubated with GTP at 37°C.
After 10 minutes, the samples were incubated with colloidal gold as fiducials (6nm), and
directly applied to Quantifoil holey carbon grids.
2. Results
58
Figure 31: TEM image of GTP-COPI vesicles embedded in vitreous ice. As no sucrose
gradient was used to purify the vesicles, the donor membranes are also present.
Magnification: 26kx, scalebar: 100nm.
Figure 31 shows an EFTEM image of one of these samples recorded at 300kV on an FEI
Polara. Clearly, a large excess of donor membranes is present. Some of these membranes
seem to bear a proteinous coat (marked with thin arrows). In addition, a protein-coated
vesicle with a diameter of 60-70nm can be identified (marked with thick arrow). In other
areas, protein-coated buds were identified (see below).
These samples were subsequently used to record single-axis tilt series. Standard low-dose
settings were strictly followed, with a magnification of 25000x at a defocus of -8 µm for initial
experiments. The tomograms were reconstructed using the IMOD software package (Kremer
et al., 1996).
Figure 32 shows slice through the tomogram of the area shown in Figure 31. A coated
vesicle with a diameter of ca. 65nm (marked with thick arrow) can be identified, as well as
coated membranes (indicated with thin arrows). A non-continous protein coat can be
discerned.
2. Results
Figure 32: Section of a tomogram recorded of COPI samples containing COPI vesicles
directly applied to a Quantifoil grid after preparation, without sucrose gradient or cushion
purification. Slice thickness: 4.5nm. Scalebar: 100nm.
Figure 33 shows tomogram slices through a different area, of another sample containing
GTP COPI vesicles directly applied to a Quantifoil grid. The boxed area is magnified in
the series of z-slices depicted in the bottom, with each slice having a thickness of 2.9nm
and a spacing of 5.8nm. Again, a non-continous proteinous coat can be clearly identified
on the membrane.
59
2. Results
60
Figure 33: Sections of a tomogram recorded of COPI samples containing COPI vesicles
directly applied to a Quantifoil grid after preparation, without sucrose gradient or cushion
purification. Slice thickness: 2.8nm, slice spacing 5.9nm. Scalebar 100nm.
The area was extracted using the IMOD package again, and the membrane and the coat
visualized using the software Amira 4.1.1 as described in Materials and Methods.
2. Results
61
Figure 34: Rendering of coated vesicle boxed in Figure 33 using the Amira software
package. Segmentation was done partly by hand and by thresholding. Membranes are
imaged in yellow, and coating protein in green.
In this segmentation, the membrane was imaged in yellow, and the coating protein imaged in
green. The coat seemed to be non-continuous and spiky (two of these "spikes" highlighted in
red).
Figure 35 shows slices of tomograms of a different area. A protrusion can be identified on the
donor membrane, forming a coated bud (boxed area). The diameter of the coated bud is
approximately 60nm. A non-continous, proteinous coat can clearly be identified. Z-slices of
the boxed area containing the coated bud are shown in the image sequence, with a slice
thickness of 2.5nm and a spacing of 5nm. Slice numbers in the tomogram are indicated.
2. Results
62
Figure 35: Sections of a tomogram of a sample directly applied to a Quantifoil grid after
preparation, without sucrose gradient or cushion purification. In this area, a coated bud that is
in the process of forming a COPI vesicle can be identified (boxed area). Scalebar 50nm.
Slices of the boxed area are displayed on the right. Each slice has a thickness of 2.5nm, and
the spacing between slices is 5nm (slice numbers indicated each).
The area was subsequently extracted using the trimvol command. Using the Amira 4.1.1
software, the membrane was marked manually, and the proteinous coat visualized by
thresholding (see Materials and Methods). In Figure 36, the membrane is labeled in yellow,
and the proteinous coat imaged in green. The coat seems to be non-continuous, but
structural features or coat symmetry cannot be discerned at this resolution and contrast yet.
2. Results
Figure 36: Rendering of the coated bud boxed in Figure 35 using the Amira software
package. For this segmentation, selection of the membranes and the coat was done partly by
thresholding. Membranes are imaged in yellow, and coating protein in green.
When this thesis was written, data acquisition on these samples was still in progress.
63
3. Discussion
3.
64
Discussion
Coated vesicles mediate intracellular transport of proteins and membranes. In the
vesiculation process, specificity of coat recruitment and cargo capture must be tightly
controlled. In both the AP-2-, AP-1- and GGA-based Clathrin system, the coating proteins
undergo phosphorylation- and ligand-induced conformational changes that transform the
soluble complex into a membrane-bound form and activate it for cargo binding.
Structural similarities exist between the different coating systems: All coating complexes
contain both α-solenoid and β-propeller domains, and structural similiarities exist between
the coatomer subunits α, β' and ε and the clathrin proteins, and between β, γ, δ and ζ-COPs
and the 4 subunits of the adaptor complexes AP-1 to AP-4 of clathrin-coated vesicles
(Eugster et al., 2000; Gurkan et al., 2006; McMahon and Mills, 2004; Schledzewski et al.,
1999). However, in each system, the shared motifs are put to different uses, as described
below.
3.1.
Conformational dynamics of coatomer
3.1.1.
Data presented in this work
Coatomer is a heptameric protein complex that, in its polymerized form, constitutes the coat
of COPI vesicles. We have analyzed the conformational dynamics of individual coatomer
molecules using single-pair, single-molecule FRET. A surface-based, single-moleculesensitive setup allowed observation of conformational changes induced by ligand binding
(Langer et al., 2008). These changes cannot be followed in solution when a conformational
change induced in a complex causes aggregation and subsequent precipitation, as shown
for coatomer (Reinhard et al., 1999). We are currently working on the automation of data
acquisition that in the future will allow screening of multiple binding partners at varying
concentrations.
Coatomer possesses different binding sites for different ligands. Cargo for retrieval to the ER
binds to α- or β’-COP in its monomeric form via dibasic motif-containing signatures (Eugster
et al., 2004). Dimeric cytoplasmic domains of cycling proteins bind to γ-COP at both its trunk
and appendage domains (Bethune et al., 2006). Synthetic COPI vesicles can be generated
from liposomes with a certain lipid composition using coatomer and Arf1 alone (Spang et al.,
1998); however, when lipopeptides representing cytoplasmic domains of p24-family proteins
were incorporated into liposomes, COPI vesicle budding was independent of the lipid
3. Discussion
65
composition of the donor membranes (Bremser et al., 1999), characterizing these
cytoplasmic domains as part of the minimal membrane machinery for COPI vesicles.
Upon binding of dimeric p23, the trunk domain undergoes a conformational change, leading
to aggregation of coatomer (Bethune et al., 2006; Reinhard et al., 1999). Monomeric
peptides of the cytoplasmic tails of p24 family proteins do not bind to coatomer (Bethune et
al., 2006), and therefore were not investigated in this study.
As all p24-family members in their dimeric form bind to γ-COP, we investigated if all of them
could also induce a conformational change to polymerize the complex. To this end, we
utilised a centrifugation assay that allows separation of the conformationally changed,
aggregated complex from the unchanged, soluble form. We found that, of all the p24-family
members and cargo proteins tested, only p23 and p24 in their dimeric form are able to
precipitate and polymerize coatomer.
To further test our findings, we performed a set of limited proteolysis experiments on
coatomer incubated with either the monomeric or the dimeric forms of p23 and p24, and the
other p24-family members and cargo proteins. As a result, again, only dimeric p23 and p24
were able to induce a conformational change in α-COP. Thus, of the p24-family members,
only dimeric p23 and p24 serve as machinery to both recruit and conformationally change
coatomer. This is of special interest, as these two proteins are the only members of the p24family that are competent to recruit Arf1-GDP to the membrane, forming the initial priming
complex on the donor membrane.
It will be of interest in the future to learn about the functions exerted by the other p24-family
members binding to coatomer.
In order to monitor the conformational dynamics of an individual subunit, we took an
independent, biophysical approach: single-molecule, single-pair fluorescence resonance
energy transfer (FRET).
To this end, we have established labeling conditions that allow predominant labeling of the αsubunit of coatomer. Thus, changes in FRET efficiency indicate a conformational change of
this subunit. The strong preference of the activated dyes to react with α-COP is not
completely surprising, because α-COP is the largest subunit of coatomer and, with its
analogy to the clathrin system, is expected to be exposed on the surface of the complex. In
addition, differential reactivity of coatomer subunits with amino-reactive reagents has been
observed previously (Pavel et al., 1998). Labeled coatomer may also be of significant use in
future experiments that address live imaging of COPI biogenesis on supported bilayers using
fluorescence spectroscopy (in a TIRF setup). This assay is currently established in a
collaboration with the lab of Jim Rothman (Columbia University, New York, USA).
3. Discussion
66
In our single-molecule sensitive assay, we found that labeling in these conditions afforded
Cy3-Cy5-labeled coatomer that contained a major population with a FRET efficiency of 63%.
This population could be monitored in subsequent binding experiments with its interaction
partners.
After addition of OST48, a monomeric, cargo-type ligand binding to α-COP and β'-COP, a
slight shift in FRET efficiency from 63% to 67% was observed. This shift is within the
resolution limits of the method used.
Strikingly, binding of dimeric p23 to coatomer via γ-COP leads to a significant shift of the
FRET efficiency of the major population from 63% to 84%. Thus, a conformational change
induced in the trunk domain of γ-COP (Bethune et al., 2006) is transmitted to the α-subunit of
the complex.
The observed transduction of the conformational change induced in γ-COP within coatomer
to α-COP is of particular mechanistic interest, as no direct interaction between γ- and α-COP
has been described so far. Therefore, at least one coatomer subunit is likely to serve as a
“link” for transmitting the conformational information from γ-COP to α-COP. A candidate for
this function is ε-COP, which has been reported to interact with both α- and γ-COP (Eugster
et al., 2000).
3.1.2.
Membrane protein capture and coat lattice formation
In light of the structural analogies of the tetrameric coatomer subcomplex β-γ-δ-ζ-COPs with
the adaptor complexes, and similarity of α, β’, and ε-COPs with the clathrin proteins ((Duden
et al., 1991; Schledzewski et al., 1999); reviewed by (McMahon and Mills, 2004)), it is of
interest to compare the functions of these complexes.
Adaptor complexes capture membrane proteins, and in a second step recruit clathrin, which
forms the vesicular cage. In contrast, in the COPI system, coatomer is recruited to the
membrane en bloc (Hara-Kuge et al., 1994), and thus the clathrin-related subcomplex (αβ'ε)
is already present during the capture of membrane proteins, a process that is mediated by
the adaptor-like subcomplex (βγδζ) by its γ-subunit. If the clathrin-related subcomplex has a
function similar to clathrin's in forming a shell, then its capability to assemble a higher
aggregate must come into effect. However, this must not occur before coatomer has
captured membrane proteins, because otherwise coatomer could not exist as a soluble
complex. In fact, when soluble coatomer binds to dimeric peptides that mimic the cytoplasmic
tail of the membrane protein p23, the complex aggregates and precipitates (Reinhard et al.,
1999). Therfore, our observation of the transmission of a conformational change through
3. Discussion
67
coatomer from the trunk subcomplex to the clathrin-related outer subcomplex points to a link
in the COPI system between membrane protein capture and cage formation. Accordingly,
dimeric p23 or p24 protein captured by the trunk domain of γ-COP would represent
membrane machinery for the formation of COPI vesicles, as opposed to cargo proteins that
bind α/β'-COP and do not alter coatomer's conformation. Thus, the conformational change
observed in α-COP could represent an initial step of coat lattice formation, rearranging αCOP to form a Clathrin-like shell or lattice on the donor membrane (Figure 37, A). This would
be in good agreement with the findings that only p23 and p24 serve as primary Arf1
receptors on the donor membrane, and form the priming complex during initiation of the
budding process.
Figure 37: Model for putative role of the conformational change in α-COP during COPI
vesicle formation.
In the light of additional mechanisms known from the clathrin system, it is tempting to
speculate on further functions of the observed conformational change of coatomer.
As mentioned, clathrin-coated vesicles form upon sequential recruitment of adaptor protein
complexes and clathrin to the respective donor membrane. During this process, the cargo
binding affinity of the adaptor complexes is tightly regulated. Among other mechanisms, e.g.
phosphoinositide binding (Rapoport et al., 1997) and phosphorylation (Collins et al., 2002;
Fingerhut et al., 2001; Haucke and De Camilli, 1999; Ricotta et al., 2002), conformational
changes induced by the binding of the cytoplasmic domains of transmembrane proteins
3. Discussion
68
specifically activate binding of other cargo to the coating machinery (Boll et al., 2002; Haucke
and De Camilli, 1999). In clathrin-coated vesicles, transmembrane proteins serve as cargo
and membrane machinery of a vesicle at the same time, and it was proposed that endocytic
coated pits may only proceed to form clathrin-coated vesicles if cargo is present (Ehrlich et
al., 2004).
In the formation of COPI vesicles, the observed conformational change within α-COP might
serve a similar purpose (Figure 37, B). As p24-family proteins are stoichiometric components
of COPI vesicles (Malsam et al., 2005; Sohn et al., 1996), the conformational change
induced by p23 upon binding to coatomer might represent a key step for recruitment of cargo
during vesicle formation. The observed transduction of a conformational change from γ-COP
to α-COP might modify the affinity of the cargo binding site in α-COP (Eugster et al., 2004)
by rearranging this subunit upon recruitment of coatomer to the donor membrane via γ-COP.
To investigate this, we made use of an ELISA-like binding assay previously established in
out lab (Bethune et al., 2006). This assay allows determination of the KD of coatomer for its
ligands, and to monitor if the number of available binding pockets changes if the complex
alters its conformation. Here, we immobilised coatomer on 96-well Protein A plates, and
monitored the affinity of S-tagged binding partners by S-tag specific detection.
Presently, we did not see a significant change in both the KD and the number of binding sites
available for cargo proteins. Thus, the conformational change in α-COP does not open a
binding pocket for cargo, as it does in the µ2-subunit of AP-2. However, the structural
rearrangement may serve in a more indirect way: it may bring the β-propeller domains closer
to the membrane during coatomer recruitment, thus faciliating interaction of coatomer's cargo
binding sites with its target cargo transmembrane proteins. Then this rearrangement would
not have any impact on the values obtained for both the KD and the number of available
binding sites, as the ELISA-like binding assay is not a membrane-based system.
Future efforts will focus on investigating such functional consequences of the conformational
dynamics of coatomer.
3. Discussion
3.2.
Electron microscopy and tomography of COPI vesicles
As described above, the different coating proteins share certain structural homologies. In
fact, all coating complexes contain both α-solenoid and β-propeller domains. However,
these re-appearing motifs seem to be put to different uses in the different coating
systems: In the Clathrin system, the β-propeller domains mediate interaction of Clathrin
with the adaptor proteins and accessory proteins (Fotin et al., 2004b). In the COPII
system, these domains (β-propeller) form the vertices of the assembled cages of
Sec13/31 (Fath et al., 2007). In coatomer, the β-proeller domains have been
characterised as the binding sites for KKxx- and KxKxx-based sorting signals (Eugster et
al., 2004).
Whereas both the COPII (Fath et al., 2007; Stagg et al., 2006) and the Clathrin system
have been described in detail at the molecular level (Cheng et al., 2007), little is known
about the structure of coatomer and the COPI vesicle lattice yet. Recent electron
microscopic studies showed "spiky" or "fluffy" coats on Golgi membranes and plant
COPI vesicles (Bouchet-Marquis et al., 2008; Donohoe et al., 2007). Structural
information is available on one subdomain of one subunit, the γ-COP appendage
domain, only, with the overall fold and structure of this subdomain showing striking
analogies to the fold of α-subunit of AP-2, (Watson et al., 2004).
In the present work, we tested various different preparation and purification protocolls to
obtain samples of coated COPI vesicles generated in GTP-conditions. As samples
containing sucrose showed only limited contrast and stability in the electron beam, a
sucrose-free sample preparation was used. These samples contained a large number of
both coated membranes and coated vesicles.
At the time when this thesis was written, data aquistion was still in progress. The first
data sets shown in this work are promising in terms of both contrast and resolution, but
the conditions for aquisition of high-resolution and high-contrast tomograms need to be
improved, especially by imaging vesicles in thinner ice.
In the future, using the protocoll described in this work, we may be able to obtain
tomograms of individual COPI-coated, Golgi-membrane-derived vesicles and resolve the
structure of the coat lattice.
69
4. Materials and Methods
4.
Materials and Methods
4.1.
Materials
4.1.1.
Reagents
70
Chemicals were bought from the companies BioRad (Munich, Germany), Roche (Mannheim,
Germany), Sigma (Deishofen, Germany) and Merck (Karlsruhe, Germany) unless stated
otherwise.
4.1.2.
Peptides
Peptides were synthesized by the company Peptide Speciality Labs (PSL, Heidelberg,
Germany) in a standard solid-phase-coupled procedure. Raw peptides were purified using
RP-HPLC, and tested by MALDI-MS.
Dimeric peptides were generated by addition of a cysteine-residue at the C-terminus, as
indicated in below.
4. Materials and Methods
4.1.3.
71
Beads
For immunoprecipitation a 1:3 mixture of Protein A Sepharose and CL4B sepharose was
used (both bought from BioRad, Munich, Germany) und stored at 4°C in an aqueous 20%
EtOH solution
4.1.4.
Molecular weight standards for SDS – PAGE
The following marker mixes were used:
•
BMW-Marker (broad molecular weight) manufactured by BioRad (Munich, Germany)
and a marker cocktail by PEQLab (Erlangen, Germany):
•
Myosine
200kD
β-Galactosidase
116kD
β-Galactosidase
116kD
Bovine serum albumine
66kD
Phosphorylase B
97kD
Ovalbumine
45kD
Bovine serum albumine
68kD
Laktat-Dehydrogenase
35kD
Ovalbumine
45kD
Carboanhydrase
30kD
RE Bsp 98I
25kD
Trypsine-Inhibitor
20kD
β-Laktoglobuline
18.4kD
Lysozyme
14.3kD
Lysozyme
14.3kD
Aprotinine
6.5kD
A prestained Marker-Cocktail with recombinant protein standards bought from BioRad
(Munich, Germany):
250kD
150kD
100kD
75kD
56kD
37kD
25kD
4. Materials and Methods
4.1.5.
72
Protease – Inhibitors
To inhibit proteases in the Golgi and coatomer preparation, and upon purification of
recombinant
proteins,
protease-inhibitor-cocktail
tablets
("complete,
EDTA-free")
manufactured by Roche Diagnostics (Mannheim, Germany) were used.
4.1.6.
Antibodies
4.1.6.1.
Primary antibodies
Name
Epitope / directed against:
Dilution for WB / remarks
C1PL
β'-COP
1:3000 serum
CM1
native Coatomer
for immunoprec. and surfaces
883
α-COP
ε-r
ε-COP
CY-96
Cy3 / Cy5
1409B
α-COP
1:1000, rabbit
Elfriede
p24
1:5000, rabbit
δ-R
δ-COP
1:1000, rabbit
Okt8
CD8-receptor
Arf1
Arf1
Arf1-C1 (Reinhard et al., 2003)
γ-R
γ-COP
1:1000, rabbit
1:1000, cross-reactivity with
N-terminus of γ-COP
1:2000, rabbit
1:1000,
monoclonal,
mouse,
bought
from Sigma
for surface preps., bought from LGC
Promochem, Wesel, Germany
Table 3: Antibodies used in this study.
4. Materials and Methods
4.1.6.2.
73
Secondary antibodies
Two systems were used for detection in Western blotting: Horseradish peroxidase-coupled
goat-anti-rabbit and goat-anti-mouse IgGs manufactured by BioRad (dilution 1:5000) for
detection using films, and fluorophore-conjugated secondary antibodies (goat-anti-rabbit,
and goat-anti-mouse) for detection in the LI-COR Odyssey System (LI-COR BioSciences,
Lincoln, USA).
4.1.7.
Activated fluorophores
Activated fluorophores were bought from Amersham Biosciences (Freiburg, Germany). For
primary amine-specific coupling to coatomer, the N-hydroxyl-succinimide ester derivatives of
the dyes Cy3 and Cy5 were employed.
Figure 38: Structure of NHS-ester-derivative of the fluorophores Cy3 (n=1) und Cy5 (n=2).
4. Materials and Methods
4.2.
Equipment
4.2.1.
FPLC-Anlagen
74
In the preparation of coatomer from rabbit liver cytosol, we employed an FPLC-unit by the
company BioRad (Munich, Germany) and an Ettan FPLC unit manufactured by Amersham
Biosciences. The columns used are indicated in the corresponding protocols.
4.2.2.
SMART
Purification of labeled coatomer was performed in a SMART system manufacted by
Amersham BioSciences was used. Columns and conditions indicated in the protocols.
4.2.3.
Spectrophotometer
The concentrations of labeled coatomer were measured in a UltroSpec 2000 UV/Vis
Spectrophotometer by Amersham Pharmacia BioTech (Freiburg, Germany) in a 100µl-quartz
glass cuvette.
The Bradfort assays were performed in a Rosys 2001 microplate photometer (Rosys, Wals,
Austria).
4.2.4.
Single molecule-sensitive confocal setup
For detection of coatomer labeling in the gels, and for the single-pair, single-molecule FRET
experiments, a custom-built confocal microscope was employed. Figure 1 shows a scheme
of the setup.
For excitation, two lasers were used: A cw-Nd:YAG laser (Lasergate, Kleinostheim,
Germany) emitting at 532nm was employed for Cy3 excitation, and a red diode laser
(PicoQuant, Berlin) pulsed by a PDL-808-Sepia control unit (80MHz, pulse width 100ps;
PicoQuant, Berlin) and emitting at 635nm was used for Cy5 excitation.
4. Materials and Methods
75
Figure 39: Scheme of the custom-built confocal setup used in this study. DC0, DC1, DC2:
Dichroic mirrors; APD: Avalanche PhotoDiode. Filter 1: 575 DF 60, Filter 2: 675 DF 60.
Both laser beams were superimposed by a dichroic mirror (650DLRP; Chroma Technologies,
Rockingham, USA; see figure 39) and directed in an inverted microscope (Axiovert 100TV;
Zeiss, Jena, Germany). Another dichroic mirror in the microscope (DualBand Beamsplitter
532/633; Chroma Technologies, Rockingham, USA; see figure 39) directed the beam into a
100x oil immersion microscope lens with a numerical aperture of 1.4 (Zeiss, Jena, Germany),
4. Materials and Methods
76
and focused it on the glass surface of the chambered cover slide for single-molecule surface
scanning, or in the gel for fluorescence scanning.
650 DLRP (DC0)
Dualband Beamsplitter (DC1)
100
100
80
Transmission [%]
Transmission [%]
80
60
40
60
40
20
20
0
200
575 DF 60
675 DF 50
640 DLRP (DC2)
300
400
500
600
700
800
Wellenlänge [nm]
0
200
300
400
500
600
700
800
Wellenlänge [nm]
Figure 40: Properties of filters and dichroic mirrors employed in the single-molecule sensitive
setup (DC0, DC1, DC2 and filters).
For surface scanning, a piezo stage (PI Analogue, Physik Instrumente) was used. For gel
fluorescence scanning, a step-motor-driven x,y-scanning stage (Scan 100 x 100, MC2000,
Märzhäuser, Wetzlar, Germany) was employed. Both setups were controlled by the same
software (DAQLineScan, D.P. Herten and C.M. Roth, Heidelberg University). The average
laser power was 25- 150µW for gel fluorescence scanning (dwell times: 1-5ms/pixel; spatial
resolution: 10-100µm/pixel) and 3-5µW for single-molecule surface scanning (dwell time:
1ms/pixel, spatial resolution: 50nm/pixel, lower threshold: 10 photons, upper threshold: 30
photons).
Fluorescence emission was collected by the same microscope lens used for excitation,
passed through the dichroic mirror, and focused on a pinhole (100µm). Prior to detection, the
beam was spectrally separated by a nonpolarizing, dichroic mirror (DC2, 640DLRP; Chroma
Techn.,Rockingham, USA; see figure 40). Reflected light was filtered by a 575DF60Bandpass-Filter (Chroma Techn., Rockingham, USA), and recorded on the first avalanchephoto-diode (SPCM AQR-13; EG&G, Kanada; "donor channel”). Transmitted radiation
passed through a 675DF50- Bandpass-Filter (Chroma Techn., Rockingham, USA), and was
detected by a second avalanche-photo-diode (SPCM AQR-13; EG&G, Kanada; “acceptor
channel”). The signals from both detectors were bundled by a router and collected by a PCcard (SPC-630, Becker&Hickl, Berlin,Germany).
4. Materials and Methods
4.2.5.
77
Electron microscopes
The following electron microscopes were used at the Max-Planck-Institute of Biophysics,
Frankfurt/Main, Germany:
A Philips CM120 transmission electron microscope, operated at 120kV (FEI Philips,
Hillsboro, USA), a FEI Tecnai G2 Spirit transmission electron microscope, operated at 120kV
(FEI, Hillsboro, USA) and a FEI Polara G2 Tecnai field emission transmission electron
microscope (FEI, Hillsboro, USA).
4.3.
Methods
4.3.1.
SDS – PAGE
SDS-treated proteins were analysed using a modified, discontinous gel system (Laemmli,
1970). Both in the stacking and in the separation gel the voltage was kept constant (120V in
the stacking gel, 180V in the separation gel).
4.3.1.1.
Stock solutions for SDS - PAGE
Acrylamide-solution 1:
Acrylamid-solution 2:
4x separation gel buffer:
4x stacking gel buffer:
Elektrophoresis buffer
30%
Acrylamide (w/v)
0.8%
N,N'-Methylenbisacrylamide (w/v)
30%
Acrylamide (w/v)
0.3%
N,N'-Methylenbisacrylamide (w/v)
1.5M
Tris-HCl, pH 8.8
0.4%
SDS (w/v)
0.5M
Tris-HCl, pH 6.8
0.4%
SDS (w/v)
25mM
Tris
192mM
Glycine
0.4%
SDS (w/v)
4. Materials and Methods
4.3.1.2.
78
Separation gels
If not stated otherwise, a custom-built gel system allowing a long separation gel (15cm) and
simultaneous loading of 22 samples, or the Protean III gel system by BioRad were used.
Homogenous separation gels contained 6%, 7.5%, 10% and 12% acrylamide and gradient
separation gels contained a gradient of 7.5% - 16.5% acrylamide, as stated in the
corresponding protocols. For the custom built gel system, the following composition was
used
Gel:
7.5%
10%
12%
7.5% - 16.5%
Acrylamide-solution 2
2.5ml
3.33 ml
4.0 ml
1.3ml
3.05ml
4x separation gel buffer
2.75ml 2.5 ml
2.5 ml
1.3ml
1.3ml
H20
5.5ml
4.16 ml
3.5ml
2.9ml
1.15ml
TEMED
10µl
10µl
10µl
5.2µl
5.2µl
APS (10%)
40µl
40µl
40µl
20.8µl
20.8µl
The gel cartridge contain a separation gel volume of appr. 10ml (15x7x0.1cm). The gel
solutions were added either directly or via a gradient mixer, and covered with isopropanol.
The comb was already inserted during separation gel polymerization. For the Protean III
system, the compositions listed in table 4 were used:
Table 4: Gel compositions used for the Protean III system.
4. Materials and Methods
4.3.1.3.
79
Stacking gels
After one hour polymerization at room temperature, the isopropanol was removed and
washed with H2O. Residual H2O was removed using filter paper, and ca. 5ml of stacking gel
solution with the composition described in Figure X were applied on top of the separation gel
already in the cartridge. To from pockets, combs with 15 and 22 teeth were used. After 1h of
polymerization at room temperature, the gels were packed in wet paper towels, sealed in
plastic wrapping and stored at 4°C.
4.3.2.
Sample preparations
4.3.2.1.
Sample preparation for SDS – PAGE
The protein solution were diluted with 4x sample buffer in a ratio of 3:4, and heated either to
70° for 10 minutes, or to 95°C for 5 minutes, as stated in the protocol. After a short
centrifugation, the samples were injected into the gel pockets.
4x SDS –sample buffer (reducing):
200mM
TRIS-HCl, pH 6
12%
β-mercaptoethanol
8%
SDS (w/v)
20%
Glycerine (w/v)
Bromphenolblau was added to the sample buffer to obtain a blue-colored sample buffer, to
faciliate loading of the samples.
4.3.2.2.
Tri-Chloro-acetic acid (TCA) – precipitation
Desoxycholate (2% stock solution) were added to the protein sample to a final concentration
of 0.2% and incubated at room temperature for 15 minutes. The sample was the adjusted to
a final concentration of 10% TCA by addition of cold 50% TCA solution (stored in ice-water),
and incubated on ice for 30 minutes. The sample containing the precipitated proteins was
then centrifuged (Heraeus-Biofuge, 13000 g, 4°C, 15min), and the supernatant discarded.
The precipitate was washed with cold aceton (stored at -20°C) and centrifuged again (see
above). After removing the supernatant, the precipitate was dried at room temperature for 5
minutes, and the precipitates proteins resuspended in an appropriate volume of 1x SDS
sample buffer.
4. Materials and Methods
4.3.2.3.
80
Immunoprecipitation (IP)
Coatomer was purified or depleted using immunoprecipitation. The IP-γ buffer was used
either with or without detergent, as stated in the corresponding protocol.
25
mM
TRIS-HCl, pH7.4
150
mM
KCl
1
mM
EDTA
0.5
%
NP-40
To bind ca. 30µg of coatomer, 200µl CM1 solution (hybridoma cell culture supernatant) and
120µl Protein A sepharose beads were used. For the incutation times stated below, the
samples were agitated in an overhead shaker at room temperature, or overnight as stated in
the protocol.
Step
Buffer / Volume
Time
Wash
IP-γ with Det. / 1ml
3x 10min
Addition / coupling of antibody
IP-γ with Det. / 500µl
1h
Wash
IP-γ with Det. / 1ml
3x 10min
Addition of coatomer .
IP-γ with Det. / 400µl
1h
Wash
IP-γ with Det.
3x 10min
Wash
IP-γ without Det.
10min
Elution with sample buffer
4x SDS sample buffer /
--
30µl
Subsequently, samples were submitted to SDS PAGE as described above.
4.3.3.
Staining of proteins in SDS – gels
4.3.3.1.
Coomassie-Staining
The gels were incubated in Coomassie staining solution for 1h at room temperature, which
led to a complete staining of the gel. Using destaining solution, the gels were subsequently
washed to give the appropriate staining levels.
4. Materials and Methods
81
Coomassie stock solution:
Coomassie staining solution:
Destaining solution:
4.3.3.2.
1g
Serva-Blue G
4g
Serva-Blue R
500ml
H2O
25%
(v/v)
Isopropanol
10%
(v/v)
HOAc
5%
(v/v)
Coomassie stock solution
10%
(v/v)
Isopropanol
10%
(v/v)
HOAc
Silver stain
Gels were treated according to the following protocol. During all steps, samples were
agitated at room temperature.
Step
Solution
Time
Fixation
50% MeOH, 10% HOAc
>1h
Wash
50% EtOH
2x 15min
Incubation with Natriumthiosulfate
10mg / 50ml Na2S2O3 x 5H2O
1min
Wash
H2O
2 x 20s
Incubation with AgNO3-solution.
100mg / ml AgNO3, 50µl Formalin
15min
Wash
H2O
2 x 20s
Develop
(3g Na2CO3, Na2S2O3 x 5H20) in
<10min
50ml H2O; 50µl Formalin
Stop
12% HOAc
>5min
Upon reaching the desired staining levels, the reaction was stopped by addition of 12%
HOAc.
4.3.4.
Western blot analysis
4.3.4.1.
Transfer of proteins separated by SDS-PAGE onto PVDF-membranes
Proteins were transferred from SDS-PAGE onto PVDF-membranes (Immobilon-P, Millipore,
Eschborn) using a "semi-dry" procedure in Trans-Blot SD cells manufactured by BioRad
(Munich, Germany). Per gel, 7 filter papers (Whatman 3mm) and a membrane were cut in
4. Materials and Methods
82
the size of the gel to be blotted. Two papers each were soaked in Anode buffers 1 and 2, and
deposited on the anode board (anode 1 soaked paper on bottom). After application of the
PVDF-membrane, the gel was deposited on top, and subsequently covered with the 3
remaining paper slips soaked in cathode buffer. After careful removal of air bubbles using a
round object, the cathode plate was set on top. Proteins were then transferred onto the
membrane at 24V (constant voltage) for 90 minutes.
Anode buffer 1:
Anode-buffer 2:
Cathoden buffer:
4.3.4.2.
300
mM
20
%
25
mM
20
%
25
mM
40
%
Tris
(v/v)
Methanol
Tris
(v/v)
Methanol
Tris
(v/v)
Methanol
Ponceau-Staining of proteins on PVDF-membranes
This method was used for staining of proteins separated by SDS-PAGE and transferred onto
PVDF-membranes. The blots were inbutated for 5 minutes with Lyon-Ponceau-staining
solution, and subsequently washed with dest. water until the desired destaining levels were
reached.
Lyon-Ponceau-staining solution:
4.3.4.3.
0.8
%
4
%
(w/v)
Ponceau
TCA
Immunochemical detection of proteins on PVDF-membranes
Membranes with transferred proteins were either incubated 1h at room temperature or over
night at 4°C in PBS-T containing 5% (w/v) milk powder. Subsequently, membranes were
washed 2x10 Minuten in PBS-T. The blots were then incubated with the corresponding
primary antibodies in PBS-T containing 1% BSA for 30 minutes at room temperature. The
blots were washed 3x10 Minuten with PBS-T, and subsequently incubated with secondary
antibody in PBS-T containing 3% milk powder for 30 minutes at room temperature. The blot
was then developed using either one of the two following systems: 1. The ECL system by the
company Amersham Pharmacia Biotech (Freiburg, Germany) for detection of peroxidasecoupled secondary antibodies. Here, the protocol of the manufacturer was used. For
detection fo the chemoluminescence, CEA RP New films (Strangnas, Sweden) were used. 2.
4. Materials and Methods
83
The LI-COR Odissey system manufactured by LI-COR Biosciences (Lincoln, USA). For
detection, the protocol of the manufacturer was followed.
4.3.5.
Bradford assay
Protein concentrations were determined using staining solutions bought from BioRad
(Munich, Germany). An appropriate volume of protein solution (about 10 – 20 µg) was diluted
to a final volume of 800µl in mp water (triplicate measurements with different dilutions), with
an 8-point calibration curve in a range of 2.5 – 20µg BSA (0, 2.5, 5, 10, 15, 20 and 25µg/ml
final concentration, in 800µl final volume). Then, 200µl of Bradford reagent were added to all
samples and incubated at room temperature for 10 minutes. 200µl of each sample were
transferred into a microtiterplate and the optical density at 620nm measured in the microplate
reader.
4.3.6.
Isolation of coatomer from rabbit liver cytosol
To achieve a higher yield and purity of the coatomer complex, the original protocol by J.
Pavel (Pavel et al., 1998) was slightly modified.
4.3.6.1.
Isolation of rabbit liver cytosol
For one preparation, 3 livers (ca. 160g) from freshly killed rabbits (starved for 12h to reduce
glycogen in the livers) were used. The livers were transferred into ice/water right after
extraction, cut into thin slices (>1cm) and placed into homogenisation buffer (200ml).
Homogenisation of the tissue was performed in a Warring blender with pulses of 2x 30s and
a pause for cooling (2min) on ice. The homogenate was centrifuged at 15000xg at 4°C for 1h
(Heraeus-Suprafuge 22, Rotor: HFA 12.500). The supernatant was filtered through 4 layers
of mull and centrifuged at 100000 xg, 4°C (Ultracentrifuge, Rotor: TFT 50.38). The
supernatant ("rabbit liver cytosol") was filtered through 4 layers of mull again.
4.3.6.2.
Ammoniumsulfate precipitation of rabbit liver cytosol
Rabbit liver cytosol was diluted 1:2 with cold homogenisation buffer 2 and transferred into a
glass beaker. Slowly, freshly pulverized Ammoniumsulfate was added to a final concentration
of 35% under constant stirring on ice, and after completed addition stirred for another 45
minutes. The precipitate was centrifuged for 30 minutes at 10.000 xg, 4°C (HeraeusSuprafuge 22, Rotor: HFA 12.500), and the supernatant was discarded. The precipitate was
resuspended in 100ml of resuspension buffer, homogenized in a Douncer and dialysed 2x 1h
and 1x overnight against 5l IEX-1 (Ion-EXchange chromatography buffer 1) each. Unsoluble
4. Materials and Methods
84
proteins were precipitated by a centrifugation at 100000 xg (Ultracentrifuge, Rotor: TFT
50.38) for 30 minutes at 4°C. This centrifugation was repeated until the supernatant was a
clear solution. This fraction was then filtered through a membrane with a pore size of 0,45µm
("ammoniumsulfate precipitate, ASP").
4.3.6.3.
DEAE anion exchange chromatography of ASP
A 300ml DEAD-FF column (Amersham Pharmacia Biotech, Freiburg, Germany) was
equilibrated with 1l of IEX-1. Subsequently, the ammoniumsulfate precipitate was applied to
the column at a flow rate of 10ml/min and washed with IEX-1 until baseline absorption was
reached again. Bound protein was then eluted with mixture of 50% IEX-1 and 50% IEX-2 at a
flow rate of 10ml/min. The whole protein peak was collected and the conductivity set to the
value of IEX-1 ("DEAE-Pool").
4.3.6.4.
SourceQ anionic exchange chromatography of the DEAE pool
A SourceQ column with a bed volume of 15ml (Amersham Pharmacia Biotech, Freiburg,
Germany) was equilibrated with 50ml of IEX-1 (flow rate 2ml/min), and the DEAE-pool
applied to the column over night (max. flow rate 1mg/ml). Subsequently, the column was
washed with IEX-1 until baseline absorption was reached again. Bound protein was eluted in
a linear gradient from IEX-1 to IEX-2 over 200ml. The eluate was collected in fractions of
3ml, and 5µl of each fraction submitted to SDS PAGE and Coomassie staining and Western
blotting. The fractions containing coatomer were pooled and adjusted with conductivity buffer
to the conductivity of IEX-1 ("SourceQ-Pool").
4.3.6.5.
Concentration of the SourceQ-Pools
A SourceQ column with a bed volume of 1ml (Amersham Pharmacia Biotech, Freiburg,
Germany) was equilibrated with 5ml IEX-1 (flow rate 250µl/min). The SourceQ pool was
loaded onto the column with the same flow rate, and the column re-equilibrated to IEX-3.
Coatomer was eluted in a linear gradient of IEX-3 to IEX-4 (0%-60% IEX-4) in a volume of
6.5ml. Coatomer was collected in fractions of 300µl and checked in Coomassie stained gels
and by Western blotting against coatomer subunits. According to purity, different fractions
were pooled. Protein concentration was adjusted to 5mg/ml, aliquoted into 10µl aliquots,
snap-frozen in liquid nitrogen and stored at -80°C.
4. Materials and Methods
4.3.6.6.
85
Buffers for coatomer preparation
Homogenisation buffer 1:
25
mM
Tris-HCl, pH 7.4
500
mM
KCl
250
mM
Saccharose
2
mM
EDTA-KOH
1
mM
DTT
2 Tabl./100ml Protease-Inhibitor, complete
Homogenisation buffer 2:
Resuspension buffer:
25
mM
Tris-HCl, pH 7.4
2
mM
EGTA-KOH
1
mM
DTT
25
mM
Tris-HCl, pH 7.4
200
mM
KCl
1
mM
DTT
2 Tabl./100ml Protease-Inhibitor, complete
IEX-1:
IEX-2:
Conductivity buffer:
IEX-3:
25
mM
Tris-HCl, pH 7.4
200
mM
KCl
1
mM
DTT
10
%
Glycerine (w/v)
25
mM
Tris-HCl, pH 7.4
1
M
KCl
1
mM
DTT
10
%
Glycerine (w/v)
25
mM
Tris-HCl, pH 7.4
1
mM
DTT
10
%
Glycerine (w/v)
25
mM
HEPES-KOH, pH 7.4
100
mM
KCl
10
%
Glycerine (w/v)
4. Materials and Methods
IEX-4:
86
25
mM
HEPES-KOH, pH 7.4
1
M
KCl
10
%
Glycerine (w/v)
Table 5: Buffers used for the preparation of coatomer from rabbit liver cytosol.
4.3.7.
Labeling coatomer
A sample of purified coatomer (50µg protein) was incubated with dye solution (600pmol of
Cy3-NHS-ester and Cy5-NHS-ester (in 2µl ddH2O each) for the single-molecule
experiments) in a total volume of 20µl. The pH was set to 8.0 by addition of 1.5µl of
previously adjusted pH-shift reagent (25mM HEPES KOH, 100mM KCl, pH 8.2). The sample
was incubated for 3h on ice. Precipitated protein was removed by centrifugation at 100,000g
for 15 min at 4°C. Unbound dye was removed by gel filtration on a Sephadex G-50 column
with a bed volume of 1.5ml mounted in a SMART FPLC system (GE Healthcare; PBS buffer:
see above). Labeled coatomer eluted in the void volume and these fractions were pooled
and stabilized by addition of BSA to a final concentration of 1mg/ml.
Prior to addition of BSA, the labeling efficiency was estimated by relating the absorption at
280nm (protein; corrected for crosstalk by the dyes, 8% for Cy3 and 5% for Cy5) and at
552nm (Cy3, ε(552)=150000 and at 649nm (Cy5, ε(649)=250000). Using these conditions,
we found about one donor (Cy3) and one acceptor (Cy5) dye per coatomer (0.943 ± 0.09
(SD,n=6)).
4.3.8.
Analysis of labeled coatomer subunits
Coatomer was purified by immuno precipitation with the antibodies CM1 (monoclonal, anti-β’COP, mouse) and 883 (polyclonal, anti-α-COP, rabbit), as described previously (Wegmann
et al., 2004), and its subunits were separated by SDS-PAGE under reducing conditions on
12% gels. Gels were prepared for fluorescence scanning by fixation in 30% (v/v) methanol.
Thereafter, they were transferred into a chambered cover slide (Lab-Tek, Nunc GmbH,
Wiesbaden) and scanned at excitation wavelengths of 532nm and 635nm in a confocal laser
setup, as described above.
4. Materials and Methods
4.3.9.
87
Isolation of rat liver Golgi
For one preparation, 11 male Wistar rats were killed and left to bleed out briefly. The livers
were instantly upon excision transferred into precooled homogenisation buffer and cut into
small pieces (<1cm). After weighing, 3 parts (v/w) of homogenisation buffer per liver mass
were added.
The liver were then homogenized using an Ultrathorax for 30s at highest possible speed. The
homogenate was transferred into 50ml Falcon tubes and centrifuged for 10min at 2200rpm at
4°C. The supernatant was filtered through 4 layers of mull to obtain Perinuclear supernatant
(PNS). This supernatant was applied onto 20ml of a 1.25M Sucrose/Tris solution (ca. 16ml of
PNS in SW-28 tube, appr. 10 tubes).
The samples were centrifuged for 1:30h at 25000rpm at 4°C. The interphase (IPH) was
harvested with a plastic pipette (appr. 35ml). The sucrose concentration of this fraction was
adjusted to 1.255M by adding 2M sucrose.
The IPH was then applied to a second gradient in SW-28 tubes, as described below (pour
from bottom):
+ 10ml 0.5M Sucrose
top
+ 10ml 1.0M Sucrose
+ 10ml 1.1M Sucrose
+ 6ml IPH
bottom
The samples were then centrifuged for 2:30h at 25000rpm (4°C). The interphase between
0.5M and 1.0M sucrose was harvested, containing the Golgi (recognizable by white flakes on
white layer). The purified Golgi membranes were then snap-frozen in liquid nitrogen and
stored at -80°C.
4.3.10. Pull down experiments
The recombinant proteins used (Trx-His6, Trx-His6-p23d, Trx-His6-OST48) were described
previously (Bethune et al., 2006). Constructs (200pmol) with the cytoplasmic domains of
OST48 or dimeric p23 and a Thioredoxin- (Trx) and His6-Tag were immobilized on NiSepharose (Amersham Biosciences) in PBS buffer (see above) by incubation for 1h at RT.
After three wash steps, the beads were blocked with 0.1% BSA (w/v). 2pmol of coatomer
were added and incubated for 30 min at RT. After three wash steps with imidazole (30mM)
and detergent (Triton X-100, 0.5%), bound coatomer was eluted from the Sepharose beads
by incubation with SDS sample buffer at 95°C, separated by SDS-PAGE and visualized by
immunoblotting.
4. Materials and Methods
88
4.3.11. Membrane binding assay
Rat liver Golgi membranes and Arf1 were prepared as previously described (Franco et al.,
1995). 10µg of enriched rat liver Golgi membranes were incubated with Arf1 (1µg), GTPγS
(100µM) and labeled or unlabeled coatomer (1µg) in a total volume of 50µl (25mM
HEPES/KOH, 20mM KCl, 2.5mM Mg(Ac)2, 0.2M saccharose, 1mM DTT, 1mg/ml ovalbumin,
40µM ATP, pH 7.2).
After incubation for 15 min at 37°C, the samples were loaded on top of a sucrose cushion
(15% (w/v), 165µl per 50µl sample volume) and membranes and bound coatomer were
precipitated by centrifugation at 20,800g for 30 min. The pellet was dissolved in SDS sample
buffer, subjected to SDS-PAGE and analyzed by immunoblotting, as described above.
4.3.12. In vitro COPI vesicle budding assay
4.3.12.1. 17h gradient purification
For a typical COPI budding assay, the following components were mixed (in the order they
are listed) for a "10ml-preparation":
AP Buffer:
25mM Hepes-KOH, pH 7.0, 2mM Mg(OAc)2
1ml
DTT:
1M
2.5µl
Sucrose:
2M
516µl
KCl:
3M
45.9µl
Rat liver cytosol:
38.2mg/ml
1.254ml
The sample was incubated for 5 minutes at 37°C, and then 1.550ml of rat liver Golgi
membranes (prepared as described above, typically 0.5mg/ml) and 10µl of a 100mM GTPγS
solution were added.
The sample was incubated for 20 minutes at 37°C and then put on ice for 10 minutes.
Subsequently, the sample was centrifuged for 20minutes at 10500rpm in an SS-34 rotor (in a
Sorvall RC-6 centrifuge) at 4°C. The pellet was resuspended in 600µl of LSSB buffer,
transferred into 1.5ml Eppendorff cups and incubated for 10minutes on ice. The samples
were centrifuged again for 15minutes at 13000 rpm (Eppendorff centrifuge 5415 R) at 4°C.
The pellet was resuspended in 600µl HSSB and incubated for 10 minutes on ice, followed by
a centrifugation at 13000rpm for 10 minutes at 4°C. The supernatant was again centrifuged
for 15 minutes at 13000rpm at 4°C to remove large membrane fragments.
4. Materials and Methods
89
The sucrose concentration of this supernatant was adjusted to ~20%. Then the sample was
applied on top of the following gradient comprised of the following layers in SW-55 tubes
(poured from bottom):
sample containing COPI vesicles
top
+ 715µl 25% Sucrose
+ 715µl 30% Sucrose
+ 715µl 35% Sucrose
+ 715µl 40% Sucrose
+ 715µl 45% Sucrose
+ 715µl 50% Sucrose
bottom
The gradients were centrifuged in a SW-55Ti rotor at 33000rpm for 18h at 4°C, and the
gradient fractionated in 200µl fractions by incising the bottom of the tubes in a custom-built
setup. The sucrose concentrations were determined and then aliquots of each fraction
submitted to Electron microscopy or to SDS PAGE and Western blotting. Typically, the
fractions with a sucrose concentration of 39%-42% contained a homogenous population of
COPI vesicles.
4.3.12.2. 1h cushion centrifugation
Rat liver Golgi membranes were submitted to a high-salt wash (1M KCl final concentration, in
AP buffer: 25mM Hepes-KOH, pH 7.0, 2mM Mg(OAc)2), diluted to a sucrose concentration of
<10% and incubated on ice for 10minutes.
After placing sucrose cushions (17%) at the bottom of the tubes, the membranes were
pelleted by centrifugation at 12000rpm for 15min in a TLS-55 rotor. The supernatant was
discarded, and the membranes resuspended in assay buffer containing 23% sucrose (125µl
per 1mg of Golgi membranes). For a typical COPI vesicle budding experiment, 200µg of
purified, salt-washed rat liver Golgi membranes were incubated with 40µg of labeled and
unlabeled coatomer, 5µg purified Arf1, 10mM GTPγS, 1mM DTT, 85mM Sucrose, 90mM KCl,
50µM GTPγS in a total volume of 500µl assay buffer (25mM HEPES-KOH, pH 7.0; 2mM
Mg(OAc)2). The samples were incubated at 37°C for 10 minutes in order to obtain primed
Golgi membranes. Then the samples were transferred on ice, and the vesicles released by
incubation for 10 minutes on ice after addition of 20µl 3M KCl (final conc.: 200µM). The
membranes were then precipitated by centrifugation at 16100 xg for 5 minutes at 4°C. The
supernatant was transferred into SW-55Ti tubes with split adaptors, and underlayered with a
cushion of 50µl 37% sucrose and 5µl 47% sucrose. After centrifugation at 32000rpm at 4°C,
the pellets were harvested from the bottom in a volume of 17µl, mixed with SDS sample
buffer, subjected to SDS PAGE and analyzed by immunoblotting.
4. Materials and Methods
90
4.3.12.3. Sucrose-free preparation of crude vesicles
For sucrose-free COPI vesicle preparation, rat liver Golgi membranes were submitted to a
high-salt wash (1M KCl final concentration, in AP buffer (see above), diluted to a sucrose
concentration <10% and incubated on ice for 10 minutes. The Golgi membranes were then
spun down by centrifuging at 12000 xg for 5 minutes.
The components were added as described for the 1h cushion centrifugation, and incubated
on ice for 5 minutes, and subsequently at 37°C for 10 minutes. The samples were then
mixed in a 1:1 rationwith a solution of colloidal gold as high contrast markers for tomography,
and directly applied to the Quantifoil grids.
4.3.13. Grid preparation for negative staining
Thin (~10nm) carbon-coated grids were generated by evaporating carbon by resistive
heating onto mice and floating onto standard 400 mesh copper electron microscopy grids.
Before using these carbon-coated grids in negative staining, the grids were freshly glow
discharged (30s). 3µl of sample were applied onto each grids, and allowed to adsorb to the
carbon backing for 2 minutes. Using filter paper (Whatman Nr. 4, Whatman Corporation,
Maidstone, UK), the sample was removed by blotting and 3µl of a solution of 1% uranyl
acetate were added. The uranyl acetate was immediately blotted again with filter paper (see
above), another drop of 3µl of 1% uranyl acetate added onto the grid and left to stain for 2
minutes. The grid was then blotted again with filter paper,and left to air dry (5 minutes) before
inserting into the electron microscopy.
4.3.14. Grid preparation for cryo electron tomography
After glow discharging at 1000V for 30 seconds, the quantifoil grids (R2/2 on copper 300
mesh grids, Quantifoil Micro Tools GmbH, Jena, Germany) were examined in the light
microscope for tears and holes in the carbon film.
Colloidal gold particles with diameters of 6,10 and 25nm (Aurion, Wageningen, The
Netherlands) as fiducial markers were mixed with the sample to be frozen in a 1:1 ratio. The
grids were secured in forceps, and a drop of 3µl of sample was applied to the carbon
surface. The forceps were then fixed into a custom-built, gravity driven freezeplunging device
(Deryck Mills, Max-Planck Institute for Biophysics, Frankfurt/Main, Germany). After blotting
for 3 -10 seconds (depending on the viscosity of the sample), the grids were plunged into a
reservoir of liquid ethane and transfered into a liquid nitrogen cooled aluminium sample
4. Materials and Methods
91
chamber, and kept in liquid nitrogen until the grids were transfered into the electron
microscope (never warming above 130K).
4.3.15. Precipitation assay
Rabbit liver coatomer (0.1µM) was incubated with the indicated concentrations of peptides
(10, 20 and 50µM for dimeric p23/24, and 50µM for the other peptides) in a total volume of
20µl of buffer (25mM HEPES-KOH, pH 7.4; 100mM KCl) for 30min at room temperature. The
samples were then centrifuged for 15min at 40000g, and the supernatant discarded. The
pellets containing precipitated protein were resuspended by addition of 5µl of 4% SDS buffer
and 5µl of 4x SDS sample buffer. The samples were then submitted to SDS-PAGE and
stained with Coomassie.
4.3.16. Limited proteolysis
Rabbit liver coatomer (0.1µM) was incubated in presence and absence of peptides in the
corresponding concentrations (monomeric peptides: 50µM, dimeric peptides 25µM) for 1h at
room temperature in 25mM Hepes-KOH pH7.4, 100mM KCl. Subsequently, Thermolysine
(0.008µM) was added for the indicated periods of time, and proteolysis was stopped by
addition of EDTA (final concentration: 40mM). After addition of SDS sample buffer, the
samples were subjected to SDS-PAGE and analyzed by immunoblotting.
4.3.17. Antibody purification
50ml of CM1 or Okt8 hybridoma cell line supernatant were incubated with 500µl Protein A
sepharose beads (Amersham Biosciences) at 4°C overnight after addition of an EDTA-free
protease inhibitor tablet (Roche) and 5ml 10x PBS. Subsequently, beads were washed with
2x 15ml PBS and then resuspended in 1ml PBS and transferred to a disposable column
(BioRad), The antibody was eluted with 3x 1ml of elution buffer (0.1M glycin, pH 2.8) into
tubes already containing 175µl of neutralization buffer to immediately neutralize the solution
(pretested on elution buffer not containing any sample).
The protein concentration was determined, aliquots (10µl) snap-frozen in liquid nitrogen and
stored at -80°C.
4. Materials and Methods
92
4.3.18. Surface preparation
Antibodies were purified by Protein A affinity chromatography (as described above). 8-well
chambered cover slides (Lab-Tek, Nunc, Wiesbaden, Germany) were cleaned and activated
with oxygen plasma (30s, Tepla 100-E, Asslar, Germany) and incubated overnight at 4°C
with PBS containing 10mg/ml BSA (Sigma-Aldrich, München, Germany) and 2µg/ml of
purified antibody (CM1 or Okt8). All subsequent steps were carried out using PBS with
1mg/ml BSA. After 5 washing steps (800µl, 5min each), labeled coatomer (3ng) was added
and incubated for 20 min at 4°C. After two wash steps with detergent (NP-40, 0.5%), the
surfaces were washed five times without detergent (5min each). The surfaces were then
directly scanned and the sp-FRET measured in the custom-built confocal setup described
above.
4.3.19. Calculation of FRET efficiency Eappr
For FRET experiments, the surfaces were illuminated with the Nd:YAG laser with emission at
532nm. Only spots showing detectable intensity in the acceptor channel (>10
photons/pixel*ms) during low intensity surface scanning were selected for subsequent timeresolved fluorescence intensity recording, and each spot was irradiated until complete
photobleaching occurred.
Only spots showing a single-step photo-bleaching event in the acceptor channel, a
simultaneous increase in the donor channel, and a subsequent single-step photo-bleaching
event in the donor channel were kept for subsequent evaluation (<5% of picked spots).
The time until acceptor photo-bleaching (Tpb), as well as the total number of photons in the
donor (ID,T) and acceptor channels (IA,T) during this period, were extracted manually from the
recorded data. In addition, the individual background of each spot was recorded after
complete photo-bleaching for at least 30s (IA,B and ID,B) as shown in figure 41.
With the parameters employed for imaging (dwell time: 1ms/pixel, spatial resolution:
50nm/pixel, lower threshold: 10 photons, upper threshold: 30 photons), the lowest detectable
FRET efficiency is 25%. Thus coatomer complexes with a FRET efficiency below 25% are
omitted from the histograms.
4. Materials and Methods
Figure 41:
93
Fluorescence intensity trace of a typical singly donor- and acceptor-labeled
coatomer complex, and scheme of data extraction that was used for subsequent calculation
of EApp. Cross-talk was measured in independent experiments.
The FRET efficiency Eapp of an individual complex was approximated from the fluorescence
intensity trace using the formula: Eapp = IA / (IA + ID) (Ha, 2001), where IA = (IA,T - IA,B) – β * (ID,T
- ID,B) and ID = (ID,T – ID,B) + β * (ID,T -ID,B). The term (Ix,T - Ix,B) subtracts background noise and
the term β * (Ix,T – Ix,B) corrects for spectral cross-talk. After subtraction of background and
correction for crosstalk, a minimal threshold was applied, and spots with n<200 photons in
either channel were discarded.
For the above calculations, the coefficient β was calculated using a surface labeled with only
Cy3-labeled coatomer (acceptor only). 20 spots were picked, photobleached and the emitted
photons were recorded. The coefficient β (β = 0.1153) was estimated from the ratio of
emitted photons in the donor and the acceptor channel. The correction factors for quantum
yield and overall detection efficiencies in the donor and acceptor channels were omitted for
the FRET pair Cy3 and Cy5, as proposed by (Ha, 2001). A scheme of the calculation of EApp
is shown in Figure 42.
4. Materials and Methods
94
Figure 42: Schematics of steps involved in calculation of EApp. The coefficient for correction
for cross-talk was measured in an independent set of experiments using only Cy3-labeled
coatomer.
Each coatomer molecule evaluated was subsequently sorted according to its energy transfer
efficiency Eapp in a histogram with a class size of 5%. The major populations present in the
histogram were assumed to be normally distributed and their mean values and standard
deviations were determined by fitting one or two Gaussian functions with a chi-squared
ranging between 0.90 and 1.22 using Microcal Origin 7.5. The discrepancy between fitted
and measured values is attributed to the assumption that random labeling generates
homogenous background. However, this deviation has no effect on the determination of the
centre of the major peaks.
4.3.20. ELISA-like binding assay
The protocol this assay was described previously (Bethune et al., 2006). Coatomer was
immobilized on 96-well microtiter plates pre-coated with protein A (Perbio science, Bonn).
First, the plates were incubated with 300 µL of PBS-T containing 5% BSA overnight at 4°C
each. Then, the wells were washed twice with PBS-T and incubated with 100 µL per well of
CM1 antibody (from a hybridoma cells supernatant) for 30 min. Subsequently, the plates
were washed with binding buffer, and then incubated for 1h at room temperature with a
solution of 0.1 µM coatomer in PBS-T containing 1% BSA (100 µL per well). Subsequently,
the plates were incubated with different dilutions of fusion proteins in binding buffer (50 mM
Hepes, 90 mM KCl, 300 mM NaCl, 1 mM EDTA, 1% BSA, 0.5% Triton X-100) for 1h at RT at
concentrations of 0 nM, 1 nM, 10 nM, 30 nM, 100 nM, 300 nM, 1 µM, 3 µM, 10 µM and 20
4. Materials and Methods
95
µM. Then, the plates were washed 5 times with binding buffer followed by incubation with
100 µl of a 1:4000 dilution of S-protein HRP Conjugate (Novagen, Bad Soden) in binding
buffer for 30 min at room temperature. The plates were then washed 3 times with binding
buffer and once with binding buffer containing no BSA and Triton X-100. Fusion proteins
were detected using a solution of tetramethylbenzidin (5 mg in 100 µL of DMSO added to 50
mL of 0.1M sodium carbonate buffer at pH 6 and containing 0.02% HO). The coloration was
stopped by adding 50 µL of a 0.5 M sulfuric acid solution in each well. For quantification, the
absorbance at 450 nm was measured with a microplate reader (Rosys Anthos)
.
4.3.21. Electron microscopy and tomography
The following microscopes were used to screen and image both negative stain grids and
cryo specimens: A Philips CM120 transmission electron microscope (Philips, FEI Philips,
Hillsboro, USA) operated at 120kV, a Tecnai G2 Spirit (FEI, Hillsboro, USA) transmission
electron microscope operated at 120kV and a Polara G2 Tecnai field emission transmission
electron microscope (FEI, Hillsboro, USA) operated at 300kV. All electron microscopes were
cooled to liquid nitrogen temperature. For cryo samples, low dose conditions were strictly
followed. Screening and imaging of selected areas was performed using Digital Micrograph
(Gatan Inc., Pleasanton, USA) coupled to a 1k x 1k CCD (on the CM120), a 1k x 1k CCD (on
the Spirit) and a Tridiem XXX 30 Gatan Imaging Filter with a 2k x 2k CCD sensor (on the
Polara; Gatan Inc., Pleasanton, USA).
4.3.21.1. Aquisition of tilt series
Tilt series were recorded using FEI tomography software (FEI Company, Hillsboro, USA).
Typically, samples were tilted from -60° to +60° in steps of 1 – 2°. Higher tilting angles were
achieved occasionally, depending on the of the imaged area in the grid square and on the
grid.
4.3.21.2. Tomographic reconstruction
The IMOD software package was employed for tomographic reconstructions (Kremer et al.,
1996). Using digital image processing steps, spurious bright pixels caused by x-rays
detected by the CCD were removed, the individual images of a tilt series were aligned using
high-contrast fiducial markers to generate a volume using weighted back-projection.
4. Materials and Methods
96
4.3.21.3. Segmentation of tomograms
To visualize information contained in the tomograms, volumes of interest were segmented
using Amira 4.1.1 (Visage Imaging GmbH, Fürth, Germany). Features of interest were
labeled with different colors, with the segmentation done partly by hand and partly by
thresholding.
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Acknowledgments
First, I want to thank Professor Felix Wieland for giving me the opportunity to work in his lab,
and for offering me these projects. I want to express my gratitude for his supervision and
patience, constant support and inspiring discussions. He opened up the fascinating and
challenging field of biochemistry for me.
Second, I would like to thank Professor Irmgard Sinning for being my second advisor.
I also want to thank Dr. Viktor Sourjik and Dr. Hans-Michael Müller for being in my
examination committee.
My thanks go to Dr. Dirk-Peter Herten for introducing me to single-molecule spectroscopy,
and allowing me to use the facilites in his lab.
Furthermore, I would like to thank Professor Werner Kühlbrandt for giving me the opportunity
to use his electron microscope facilites at the Max-Planck-Institute of Biophysics, Frankfurt.
I am obliged to all members of the Wieland group in Heidelberg, especially Julien Bethune,
Jörg Moelleken, Rainer Beck, Stefanie Hubich, Carolin Weimer, Priska Eckert, Michael
Geiling, Emily Stoops and Britta Brügger. Only rarely could one hope to find a group of
people that are scientifically as critical and encouraging, and form a community that reached
far beyond the lab.
I would also like to thank the members of the Herten group for introducing me to singlemolecule spectroscopy, especially Christian Roth, Thomas Heinlein and Alexander Kiel.
In my second project, I spent a lot of time at the Max-Planck-Institute in Frankfurt. Here I
want to thank Deryck Mills, Janet Vonck and Mike Strauss for teaching me electron
microscopy, for their patience and encouragement even when my samples proved to be
challenging.
Finally, and most of all, I want to thank my parents.
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