Submitted by Anny Fis Submitted at Institute of Biophysics Supervisor and First Examiner Peter Hinterdorfer Second Examiner Peter Pohl January, 2018 JOHANNES KEPLER UNIVERSITY LINZ Altenbergerstraße 69 4040 Linz, ¨Osterreich www.jku.at DVR 0093696
Forces and Dynamics in
Protein
Translocation
through
the
Bacterial
Translocon
Doctoral Thesis
to obtain the academic degree of
Doktorin der technischen Wissenschaften
in the Doctoral ProgramF O R C E S A N D D Y N A M I C S O F P R O T E I N T R A N S L O C AT I O N T H R O U G H T H E B A C T E R I A L T R A N S L O C O N
S TAT U T O R Y D E C L A R AT I O N
I hereby declare that the thesis submitted is my own unaided work, that I have not used other than the sources indicated, and that all direct and indirect sources are acknowledged as references. This printed thesis is identical with the electronic version submitted.
Linz, January 2018
A C K N O W L E D G M E N T S
The three years of my PhD studies have been a life-changing experience for me and it would not have been possible to succeed without the support and guidance that I received from many people. I would like to express my sin-cere gratitude to:
• my supervisor Prof. Hinterdorfer, not only for offering me the great op-portunity to become a member of the interdisciplinary graduate college NanoCell, but also for his continuous support, motivation and immense knowledge.
• Prof. Pohl, the second member of my thesis committee, for his insightful comments, support, and for sharing his broad knowledge on the Sec machinery.
• the third member of my thesis committee, Prof. Gruber, for his willing-ness and enthusiasm to provide this study with new inputs as well as for the stimulating discussions.
• Mirjam Zimmermann and Roland Kuttner for the fruitful collaboration that we have established the last three years.
• the AFM group for the pleasant working environment and especially Lukas, Melli, Jürgen, Yoo Jin and Andi. It was great sharing the lab facilities with you all during the last three years.
• the NanoCell consortium for the fruitful discussions and the great mo-ments in - and especially - out of the labs.
• the Austrian Science Fund (FWF) for providing the financial support enabling us to conduct this study.
• my helpful, interested, encouraging and always supportive life partner. • my parents and my sister for the emotional support, the wise counsel
and for everything that cannot be described in words.
A B S T R A C T
Many membrane and secretory proteins are translocated across the endo-plasmic reticulum membrane or the bacterial/archaeal plasma membrane through a conserved channel, the Sec61 or the SecYEG complex, respectively. In post-translational translocation the SecYEG channel associates with the cy-toplasmic motor protein SecA to deliver secretory proteins to the periplasm. Subsequently, in an ATP-dependent way, the polypeptide chain is trasported to the periplasm. Less is known about the exact binding mechanism behind the initial interaction between SecA and SecYEG, but also the translocation model that describes the movement of the polypeptide chain through the channel has been discussed controversially. In this thesis we are suggesting a binding model based on a single molecular approach. Performing experi-ments with the fully assembled translocon allowed us to imitate the entire translocation process. For these purposes, a newly developed mode capable of detecting functional SecYEG channels and providing quantitative informa-tion on different interacinforma-tions of interest was employed. This mode is a combi-nation of high resolution simultaneous topographical and recognition (TREC) imaging with AFM single molecule force spectroscopy. Moreover, appropri-ate tip and surface chemistry had to be used in order to establish a system that could simulate the physiological conditions while at the same time, it can be studied by AFM. Deeper understanding on kinetic parameters, bond lifetimes and interaction energy landscapes was gained by a dynamic inves-tigation of the entire system by means of AFM-based dynamic force spec-troscopy. A critical discussion of the obtained data in the light of the existing literature, provided new insights into the initial binding of SecA to SecYEG as well as into the role of the pure lipid membrane. Along with these findings, experiments carried out in the presence of the preprotein ProOmpA and ATP consist the initial steps towards differentiation between the proposed translo-cation mechanisms.
Z U S A M M E N FA S S U N G
Viele Membranproteine und sekretorische Proteine werden durch die Mem-bran des endoplasmatischen Retikulums oder durch die PlasmamemMem-bran von Bakterien und Archaeen transloziert, wobei die jeweiligen Transportsys-teme entweder als Sec61 oder SecYEG bezeichnet werden. In der posttrans-lationalen Translokation bindet das zytoplasmatische Motor-Protein SecA an das Kanalprotein SecYEG, um sekretorische Proteine auf die periplasmatis-che Seite zu transportieren. Der zugrunde liegende Bindungsmechanismus zwischen SecA und SecYEG ist noch unklar, ebenso wird das Translokations-modell, welches die Bewegung der Polypeptid-Kette durch den Kanal be-schreibt, kontroversiell in der Literatur diskutiert. In dieser Arbeit schlagen wir ein Bindungsmodell vor, welches auf einzelmolekularen Kraftspektros-kopie-Messungen basiert. Durch Experimente, in welchen alle Komponenten des Translokons inkludiert wurden, konnten wir den gesamten Translokations-prozess imitieren. Hierzu nutzten wir eine kürzlich entwickelte Analyse-Me-thode, mit der wir einzelne funktionelle SecYEG Kanalproteine sowohl loka-lisieren als auch deren Interaktion mit unterschiedlichen Bindungspartnern untersuchen konnten. Diese Methode stellt eine Kombination aus hochau-flösender Rastersondenmikroskopie und einzelmolekularer Erkennungs-Kraft-spektroskopie dar, wofür eine geeignete chemische Modifizierung der Mess-nadel und der Oberfläche vorgenommen werden musste um eine möglichst nahe Rekonstruktion der nativen Bedingungen zu erreichen. Der Einsatz von dynamischer Kraftspektroskopie ermöglichte die genaue Bestimmung der kinetischen Parameter und der zugrunde liegenden Energielandschaft. Eine kritische Erörterung der erhaltenen Daten mit Berücksichtigung der ex-istierenden Literatur erlaubte neue Einblicke hinsichtlich der Bindung von SecA an SecYEG, als auch neue Erkenntnisse über die Rolle der Lipidmem-bran. Zusätzliche Experimente in Gegenwart des Präkursor-Proteins ProOmpA sowie ATP ermöglichten zudem eine genauere Differenzierung zwischen den beiden vorgeschlagenen Translokationsmechanismen.
P R E FA C E
The dissertation “Forces and Dynamics in Protein Translocation through the Bacterial Translocon” describes the results of my PhD study, initiated in Oc-tober 2014 and completed in December 2017. The study was performed to its full extent at the Institute of Biophysics, Department of Applied Experimental Biophysics, JKU Linz under the supervision of Prof. Hinterdorfer. The project was well embedded within the framework of the interdisciplinary graduade college NanoCell, established among Johannes Kepler University Linz, the Technical University Vienna and the Institute of Science and Technology Aus-tria. This study is a result of a close collaboration with the Pohl group (JKU, Linz) and the Gruber group (JKU, Linz). The following thesis has been writ-ten in order to fulfill the graduation requirements of the doctoral program in Biophysics at JKU Linz.
The Austrian Science Fund (FWF) provided the financial support, and is gratefully acknowledged.
C O N T E N T S
i i n t r o d u c t i o n 1
1 m o t i vat i o n a n d a i m s 3
2 t h e b i o l o g i c a l s y s t e m 5
2.1 Membrane proteins 5
2.2 The translocation machinery 5
2.3 The bacterial translocation channel SecYEG 6
2.4 The motor-protein SecA 8
2.4.1 Translocation mechanism 9
3 at o m i c f o r c e m i c r o s c o p y (afm) 11
3.1 AFM imaging 11
3.2 Simultaneous topography and recognition (TREC) imaging 14
3.3 AFM-based single molecule force spectroscopy 15
3.4 Dynamic Force Spectroscopy 19
3.5 Combined TREC imaging with single molecule force spectroscopy 24
3.6 Tip chemistry 25
ii e x p e r i m e n ta l s e c t i o n 29
4 e x p e r i m e n ta l s e c t i o n 31
4.1 Materials 31
4.1.1 Buffers and Reagent Solutions 31
4.1.2 Proteins 32
4.2 Methods 34
4.2.1 Tip and surface chemistry 34
4.2.2 Atomic Force Microscopy 36 iii r e s u lt s a n d d i s c u s s i o n 43
5 r e s u lt s 45
5.1 Binding mechanism behind the bacterial translocation process 45
5.1.1 Interaction between the motor protein SecA and SecYEG reconstituted into a lipid bilayer 46
5.1.2 The role of ATP in the reconstituted configuration 52
5.1.3 Detergent-immobilized SecYEG 52
5.1.4 Altering the lateral pressure profile of the bilayer 55
5.2 Mimicking the translocation process by AFM 58
5.2.1 The role of the preprotein’s signal sequence during the binding process 58
5.2.2 Forces in protein translocation through the bacterial translo-con 59
5.2.3 The translocation mechanism 63
6 d i s c u s s i o n a n d c o n c l u s i o n s 67
iv a p p e n d i x 71
a a p p e n d i x 73
L I S T O F F I G U R E S
Figure 2.1 Association of membrane proteins with the lipid bilayer. 5
Figure 2.2 The structure of SecYEG. 7
Figure 2.3 The crystal structure of SecA from B. subtilis. 9
Figure 2.4 The “push and slide” mechanism. 10
Figure 2.5 The Brownian ratchet mechanism. 10
Figure 3.1 The Atomic Force Microscope. 12
Figure 3.2 Force regimes for the tip-sample interactions. 12
Figure 3.3 The principle of TREC imaging. 14
Figure 3.4 The importance of the oscillation amplitude for TREC imaging. 15
Figure 3.5 Comparison of full and half amplitude TREC. 16
Figure 3.6 The force-distance cycle. 18
Figure 3.7 Examples of force-distance cycle shapes. 20
Figure 3.8 Energy profile of the dissociation process. 22
Figure 3.9 Energy landscapes of the dissociation process along a reaction coordinate with the corresponding loading rate dependence according to the Bell-Evans model. 24
Figure 3.10 The three steps of AFM tip functionalization. 27
Figure 4.1 AFM tip functionalization with SecA and proOmpA pro-teins. 35
Figure 4.2 Immobilization of SecY(A420C)EG mutant detergent-micelles on a silicon surface. 36
Figure 4.3 Typical example of the overview kspec19p plot. 40
Figure 4.4 Exemplary graphs of the two different loading rate de-pendence evaluations. 41
Figure 5.1 Schematic representation of the used periodically sus-pended membranes. 46
Figure 5.2 Experimental configuration of combined TREC with sin-gle molecule force spectroscopy. 47
Figure 5.3 Structure of cocrystallized SecA-SecYEG complex. 48
Figure 5.4 Binding probability of the interactions between SecA and lipid membrane (PLE) and SecA and SecYEG re-constituted into lipid bilayer SecYEG. 49
Figure 5.5 Bell-Evans fits of the loading rate dependence scatter
plots of SecA interacting with SecYEG and the PLE lipids. 50
Figure 5.6 SecA(WT) and SecA∆N20 interaction with SecYEG re-constituted into a lipid bilayer measured by TREC. 51
Figure 5.7 The role of ATP in the reconstituted system. 52
Figure 5.8 Detergent-immobilized SecY(A204C)EG. 53
Figure 5.9 Average rupture forces for SecA(WT) and SecA ∆N20 interacting with detergent-immobilized SecY(A204C). 54
gent immobilized SecY(A204C)EG.
Figure 5.11 Tip- or surface-immobilized SecYEG proteoliposomes. 56
Figure 5.12 Comparison of binding probabilities between untreated and TFE-treated lipid membranes. 57
Figure 5.13 Interaction forces between SecA(WT) and SecYEG in the TFE treated membrane. 57
Figure 5.14 Overview of the determined bond lifetimes of the in-teraction SecA(WT or ∆N20 )-SecYEG under different conditions. 58
Figure 5.15 Schematic comparison of the translocation process in vivo and the mimicked translocation by AFM. 59
Figure 5.16 TREC imaging with two different ProOmpA constructs. 60
Figure 5.17 Binding probability plot of the fully assembled translo-con. 61
Figure 5.18 Most probable unbinding force of the fully assembled translocon. 62
Figure 5.19 Loading rate dependency plot of the fully assembled translocon. 62
Figure 5.20 Model of SecA translocating a polypeptide through the SecYEG channel. 63
Figure 5.21 Comparison of binding probabilities of the fully assem-bled translocon in the presence of ATP or AMP-PNP. 64
Figure 5.22 Interaction forces of the ProOmpA with fully assembled translocation in the presence of ATP or AMP-PNP. 65
Figure 6.1 Proposed binding model for the SecA-SecYEG interac-tion. 68
Figure A.1 SDS-PAGE of the SecYEG complex. 74
Figure A.2 SDS-PAGE of SecA. 75
Figure A.3 SDS-PAGE of ProOmpA. 76
L I S T O F TA B L E S
Table 5.1 Sequences of ProOmpA. 60
A B B R E V I AT I O N S
ADP adenosine diphosphate
AC alternating current
AFM atomic force microscopy
AMP-PNP adenylyl-imidodiphosphate
APTES (3-aminopropyl)triethoxysilane
ATP adenosine trisphosphate
BP binding probability
CB crystallization buffer
DC direct current
DDM n-Dodecylb-D-maltoside
DFS dynamic force spectroscopy
DHFR dihydrofolate reductase DMSO dimethylsulfoxide DOPC 1,2-dioleoyl-sn-glycero-3-phosphocholine DOPE 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine DOPS 1,2-dioleoyl-sn-glycero-3-phospho-L-serine DTT dithiothreitol
EDTA ethylenediaminetetraacetic acid
EGTA ethylene glycol-bis(2-aminoethylether)-N,N,N´, N´-tetraacetic acid
ER endoplasmic reticulum
FDC force-distance cycle
FPLC fast protein liquid chromatography FRET Förster resonance energy transfer
FWHM full width at half maximum
HBS HEPES-buffered saline
HEPES 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid
HSD helical scaffold domain
HWD helical wing domain
HS-AFM high speed atomic force microscopy
LRD loading rate dependence
MAC magnetic alternating current
MD molecular dynamics
NBF nucleotide binding fold
NHS N-hydroxysuccinimide
NTA nitrilotriacetic acid
OBD optical beam deflection
OD600 optical density (at wavelength of 600 nm)
PBS phosphate buffered saline
PDB protein data bank
PDF probability density function
PEG polyethylene glycol
PM plasma membrane
ProOmpA precursor form of outer membrane protein A
PLE polar lipid extract from E.Coli
PPXD preprotein cross-linking domain
PSM periodically suspended membrane
SA streptavidin
Sec61 eukaryotic translocation channel
SecA ATP motor-protein
SecB cytosolic chaperone
SecYEG bacterial translocation channel
SDS sodium dodecyl sulfate
SMFS single molecule force spectroscopy
SPR surface plasmon resonance
SRP signal recognition particle
STM scanning tunneling microscope
TB translocation buffer
TCEP tris(carboxyethyl)phosphine
TFE trifluoroethanol
TM transmembrane
S Y M B O L S
s y m b o l d e s c r i p t i o n u n i t
A surface m2
A0 zero frequency amplitude m2s
Awhite white noise amplitude m2s
c (molar) concentration moll−1, M
F force N
F∗ most probable unbinding force N
G∗ activation free energy J
k(F) dissociation rate under force s−1
kB Boltzmann constant 1.38·10−23 JK−1
kc cantilever spring constant Nm−1
ko f f dissociation rate constant s−1
Kd equilibrium dissociation constant moll−1, M
ke f f effective spring constant Nm−1
kl spring constant of the linker molecule Nm−1 N number of unbinding events
P area under power spectrum curve m2
p(F) probability distribution of forces at rupture Q quality factor
r loading rate Ns−1
S(t) time-dependent bond survival probability
σ standard deviation
T temperature K
t time s
τ lifetime s
v (pulling) velocity ms−1
ω radial frequency rads−1
ω0 radial resonance frequency rads−1
xβ width of energy barrier m
z cantilever deflection m
Part I
I N T R O D U C T I O N
This part presents an overview about the investigated biological system with emphasis on the two major key players SecYEG and the ATP motor-protein SecA. Additionally, a description of the principle of function of the atomic force microscope is given, as it is the main technique used for the purposes of this study.
1
M O T I VAT I O N A N D A I M S
A percentage of 25-30% of proteins are embedded in membranes or are re-quired to cross biological membranes in order to carry out their distinct func-tions. Transport of proteins during or after their biosynthesis across the brane of the endoplasmic reticulum (ER) or across the bacterial plasma mem-brane is mediated through an evolutionary conserved protein-conducting channel. It consists of a heterotrimeric membrane protein complex, called Sec61p in eukaryotes and SecYEG complex in bacteria and archaea, respec-tively. Two different translocation modes are existing through the Sec com-plex. First, the co-translational translocation that is most commonly preferred for eukaryotic protein secretion and membrane protein insertion in all do-mains of life. The second mode, which is the mode that we were interested in within this study, is the post-translational translocation and is used primar-ily for protein secretion in bacteria and archaea. During post-translational translocation the ATP motor protein receives the unfolded preprotein from the cytosolic chaperone and subsequently associates with the SecYEG. In an ATP-dependent way, the polypeptide chain gets transported from the cyto-plasm to the pericyto-plasm. Despite the fact that this translocation process is a topic of discussion among different research groups, the exact binding mech-anism is poorly understood.
In the last decade, big steps towards understanding the protein transloca-tion were made, especially after the determinatransloca-tion of SecA’s and SecYEG’s crystal structures.[112, 117] However, the dynamic mechanism underlying
the transportation of polypeptide sequences across the bacterial plasma mem-brane remains puzzling until today. What gives the direction to the substrates to move towards the periplasm and what is the exact function of ATP and the protein motive force during the protein conducting process are still not elucidated. A model that could describe this mechanism remains a matter of contention as two assumptions diametrically opposed to each other have been proposed until today. In 2014, the Rapoport group proposed an actively translocated substrate through a power stroke mechanism initiated by SecA’s two helix finger,[10] whereas two years later Collinson et al. with their
Brown-ian ratchet model suggested that no power stroke mechanism is necessary for an efficient preprotein translocation.[5] According to the latter model, the
pre-proteins can freely diffuse with Brownian motion within the SecYEG channel until they get transported to the periplasm.
By screening the existing literature we have observed, that despite the fact that the atomic force microscope (AFM) is a powerful and versatile tool, it has been barely used for SecYEG studies. To our knowledge, four studies applied the AFM or the high-speed AFM to investigate the translocation system.[74,109,81,22] In those studies only the imaging mode was used for
topographical characterization of the SecYEG molecules or for estimation of
the density of the channels on the sample surface. We are assuming that the reason that no interaction studies were performed by AFM on the translocon is the complex design of those experiments. Parameters such as orientation and functionality of the SecYEG molecules, efficient AFM tip functionaliza-tion and the establishment of a robust experimental approach have to be considered.
To address the above mentioned open questions we used a combination of simultaneous topography and recognition (TREC) imaging with single molecule force spectroscopy (SMFS) in order to obtain information about the topography of reconstituted SecYEG into suspended lipid bilayers together with interaction forces between the two key players SecA and SecYEG, but also with the pure lipid membrane for the characterization of the initial SecA-SecYEG binding. Upon establishment of a functional system, we added the preprotein ProOmpA (precursor form of outer membrane protein A) and ATP, aiming to gain a deeper insight into the mechanism that drives the sub-strates through the SecYEG channel.
2
T H E B I O L O G I C A L S Y S T E M
2.1 m e m b r a n e p r o t e i n s
Lipid bilayers are the principal structural elements of cell membranes but the membrane proteins are the elements that provide their specific charac-teristics. Lipid bilayers are tightly associated with different types of proteins that are either permanently embedded in the lipid membrane either they get temporarily attached onto them. These two categories are referring to the so-called integral membrane proteins that in most of the cases are spanning the lipid bilayer and the peripheral proteins that are coming in intermittent con-tact via non-covalent interactions with the lipid membrane. Integral proteins can be grouped in three major categories (fig. 2.1): (i) α-helices that mainly
Figure 2.1: Association of membrane proteins with the lipid bilayer.[4] function as receptors, (ii) helical bundles that provide the cell either with their enzymatic activity as transporters or as receptors, and, (iii) β-barrels that play a role in channel proteins.
2.2 t h e t r a n s l o c at i o n m a c h i n e r y
Numerous proteins need to get inserted or transported through biological membranes in order to reach their area of function. The Sec channel is one of the major pathways for soluble polypeptide membrane intergration and for translocation of hydrophobic segments to the periplasm. The protein-conducting channel is highly conserved through evolution in all domains of life. The translocation channel is an integral membrane heterotrimeric protein complex, termed Sec61 complex in eukaryotes and SecYEG in archaea and bacteria. The translocation channels Sec61/SecYEG are passive pores allow-ing polypeptide chains to move back and forth. Two major Sec pathways exist: the translational and the post-translational translocation pathway. The co-translational system is the primary pathway for protein secretion in
otes, and for protein insertion in all domains of life. The post-translational pathway is used primarily by prokaryotes for protein secretion, although an increasing number of eukaryotic proteins have been reported to follow this pathway as well.[26] During the the co-translational translocation mode, the
ribosome binds to the protein-conducting channel and the unfolded polypep-tide chain is transferred directly from the ribosome’s exit through or inside the Sec complex. The targeting of the nascent chain to the membrane occurs by the signal recognition particle (SRP) and the corresponding membrane re-ceptor (SRP rere-ceptor).[61][90] It has been suggested that for co-translational
translocation, four copies of the Sec61 complex are needed for connecting to the ribosome via four to seven connection points. Taking into account the asymmetric shape of the ribosome, the four copies of Sec61 have to connect differently to it.[94]
The second translocation mode, the post-translational translocation, is a pathway used both by eukaryotic and prokaryotic proteins. In eukaryotes, proteins that use this pathway carry less hydrophobic signal sequences com-pared to the proteins that get co-translationally translocated. The transloca-tion channel in this case is associated with another membrane complex, the Sec62/63 which is a tetrameric complex that together with Sec61 forms a multicomponent Sec channel.[34] In prokaryotes, the post-translational
path-way is followed by the vast majority of preproteins. In this configuration, the heterotrimeric translocation channel is termed SecYEG. The most important partner of SecYEG is the ATP-motor protein SecA. Numerous studies have re-ported that SecA undergoes certain conformational changes upon ATP bind-ing and transports preproteins step-wise into the SecY complex.[47] In the
following sections, the SecYEG channel and the motor protein SecA will be discussed in further detail, as they consist the major key players of the bac-terial post-translational translocation and are of utmost significance in this study.
2.3 t h e b a c t e r i a l t r a n s l o c at i o n c h a n n e l s e c y e g
The SecYEG channel is a heterotrimeric transmembrane complex, sitting in the inner bacterial membrane. Detailed information on the structure of Se-cYEG weas extracted in 2004 when Rapoport et. al[95] obtained the first high
resolution crystal structure from Methanocaldococcus jannaschii.
The main and largest subunit of the complex is the SecY that is divided into two halves consisting of five transmembrane segments (TM) each (TM1-5 and TM6-10). The two halves are connected by an external loop between TM5 and TM6, which is responsible for the opening of the lateral gate for preprotein membrane insertion. The two smaller subunits, termed SecE and SecG, respectively, have only one TM each and their N-termini are exposed to the cytosol. It is important to mention, that SecG comes only in limited contact with the main subunit SecY whereas SecE is in contact with four transmembrane segments (TM1, TM5, TM6, TM10) in both SecY halves and clamps them together.[117] Consequently, deletion of the SecE subunit leads
2.3 the bacterial translocation channel secyeg 7
to protein instability and thus is important for the viability of the cell.[77]
Concerning the SecG subunit, it is assumed that its existence is not necessary for protein functionality but it has been reported that it might facilitate the translocation process in vitro. In the present study the SecYEG was extracted from Escherichia Coli, whose structure is highly similar to Methanocaldococcus jannaschii.
Figure 2.2: The structure of SecYEG. (a) Side view of the crystal structure of SecYEG from M. Jannaschii (PDB 1RHZ). TM1-5 is shown in blue, forming an hourglass-shaped structure. The TM6-10 are depicted in orange. The plug domain that seals the pore of the channel is highlighted in red. The SecE and the SecG are shown in green and purple, respectively.[117] (b) Top view from the cytoplasmic side. (c) Side view of a complex of SecYEG and SecA from T. Maritima (PDB 3DIN). In this schematic representation the conformational rearrangements of SecYEG upon SecA binding can be seen. The differences are depicted in pink, green and purple colors.[35] A large opening located in the cytoplasmic site of the SecYEG complex of approximately 20-25 Å could function as channel entrance. The size of this funnel-like cavity reduces as we are moving towards the middle of the mem-brane and an hourglass-shaped channel is revealed. The pore ring located in the middle of the channel is consisting of six hydrophobic residues and blocks the channel during preprotein translocation. Of great importance for channel functionality is a segment of TM2 that is commonly termed in the literature as “the plug” that seals the entire protein-conducting channel by separating the cytoplasmic side from the outer aqueous environment. In that way, in the channel’s closed state the permeation of ions or of other smaller molecules is inhibited. In the channel’s active state the plug moves aside and allows the movement of preproteins through the channel.
A matter of controversial discussion is the oligomeric state of the SecY complex during polypeptide chain translocation. Despite the fact that a sin-gle SecYEG copy is enough for efficient preprotein translocation,[75] some
studies support the idea that a dimeric SecYEG state enhances SecA binding and preprotein translocation.[13]
2.4 t h e m o t o r-protein seca
A functional translocation channel consists of the SecYEG complex, that is assumed to be the passive pore in the inner membrane, and the translo-case motor-protein SecA that provides the driving force in post-translational translocation. The first crystal structure of the SecA translocation ATPase was obtained in 2002 by Hunt et al.[71] As seen in fig. 2.3, SecA has two
nucleotide binding folds NBF-I and NBF-II (shown in blue and yellow), a preprotein-crosslinking domain (PPXD) (shown in green), a helical scaffold domain (HSD) and a helical wing domain (HWD) (shown in light and dark orange, respectively). ATP binds with high affinity to a site of SecA that is located between NBF-I and NBF-II. While interacting with the translocation channel SecYEG, SecA forms a contact area of approximately 6800 Å2, as revealed by the crystallized SecA-SecYEG complex. The major interaction be-tween the two key players occurs bebe-tween the PPXD of SecA and the 8-9 and 6-7 loops of SecYEG.[127] Of great importance for SecA’s functionality
are two more structural elements, the N-terminal domain and the so-called “two-helix finger” (fig. 2.3, red). According to the crystal structure, the
N-terminal domain is the only domain that can come in contact with the lipid membrane, therefore it is assumed that the lipophilic behavior of SecA arises only from the positively charged N-terminus. The interaction of SecA with the lipid membrane will be discussed extensively in following sections as it represented a major topic of investigation within our study. The second im-portant structural element consists of the two shorter HSD helices. These two alpha helices can be inserted into the cytoplasmic funnel and facilitate with their movement the transportation of the preprotein substrate through the channel.[49][10] The formed angle between the two fingers is approximately
45º in relation to the membrane’s plane.
Both SecYEG and SecA are assumed to undergo numerous conformational changes upon complex association and during substrate translocation. Crys-tallization of monomeric SecA revealed a large conformational change upon ATP binding, resulting in a groove formation, similar to the peptide binding site of other proteins. This open conformation is assumed to represent SecA’s active state.[95] More than 25 protein structures are available in the protein
data base obtained from different organisms and under different conditions. Substitution of ATP by other ATP analogues such as ADP or non-hydrolizable AMP-PNP may alter SecA’s binding behavior to SecYEG and to the pure lipid membrane as well. Bulk measurements by Surface Plasmon Resonance (SPR) indicated a higher binding affinity for the SecA-SecYEG interaction when ATP was present in the system.[31]
Early studies on SecA’s oligomerization revealed a dimeric state at native concentrations,[2] a topic which remained under discussion until today.
How-ever, what can be said with certainty, is that a number of parameters can influence the oligomeric state of SecA. The presence of negatively charged lipids, detergent, salt, and different signal peptides is known to play a role for oligomerization of SecA.
2.4 the motor-protein seca 9
Figure 2.3: The crystal structure of SecA from B. subtilis. Left structure: Closed con-formation of SecA (PDB 1M6N). In blue and yellow the two nucleotide binding folds are depicted (NBF-I and NBF-II). The HSD (orange) is lo-cated between the NBF-II and the HWD. The PPXD and the two-helix finger are depicted in green and red, respectively.[71] Right structure:
Open structure of SecA (PDB 1TF5). Here, the displacement of the PPXD (green) towards the NBF-II (yellow) can be observed. The color coding remains the same.[95]
2.4.1 Translocation mechanism
Two major mechanisms have been proposed until today concerning the translo-cation process. In 2004, Bauer et al. suggested a combination of power stroke and passive diffusion (described as a “push and slide model”[10]) that is a
more detailed description of previously described translocation mechanisms. [97,89] The main assumption of this model is that upon ATP binding to SecA,
the two helix-finger is interacting with certain amino acids of the polypeptide chain and actively transports them through the SecYEG channel. In case that SecA’s two helix finger does not interact with the amino acid of the polypep-tide chain, it can freely diffuse back and forth. A crucial role in this process plays the hydrolysis of ATP that is assumed to reset the fingertip without needing a movement of the polypeptide chain.[10] Additionally, in the
ADP-bound state the substrate can move back and forth in the SecYEG channel independently of the interacting amino acids (fig.2.4).
The second possible translocation scenario was recently suggested by Allen et al. and supports the idea of a Brownian mechanism.[5] In this model, the
polypeptide chain can diffuse back and forth into the channel until a bulky residue blocks its entrance. This gate blocking triggers the exchange of the energy molecule and sequentially ATP causes a pore widening within the Se-cYEG channel. This pore extension allows the bulky residue to pass through the channel until the point that ATP hydrolysis reinstates the size of the pore as depicted in fig.2.5. The validity of these two models will be discussed
ex-Figure 2.4: The “push and slide” mechanism. The circles symbolize the amino acids of the translocated polypeptide. The two-helix finger (in blue) undergoes a power stroke (curved arrow) in ATP-bound state. If an interaction oc-curs between the two-helix finger and a certain amino acid (green), it pushes it forward (green arrow). Otherwise, the amino acid (in red) can diffuse forward or backward . The two-helix fingers get reset upon nu-cleotide exchange from ATP to ADP (purple, curved arrow). In the ADP-bound state, SecA allows passive sliding of any amino acid.[10]
tensively in a later section as the characterization of the translocation mecha-nism constituted a major objective of this study.
Figure 2.5: The Brownian ratchet mechanism. SecYEG is shown in red (the lateral gate in light red), SecA in blue (substrate channel in light blue and the two-helix finger in cyan), and the substrate in green (with the signal se-quence as a turquoise rectangle). The substrate can diffuse bidirectionally until the point that a bulky amino acid blocks the entrance or the exit of the channel (i-ii). A block at the entrance triggers nucleotide exchange (iii), and consequently the lateral gate opens (iv). With a wider pore the previously blocked amino acid can diffuse within the pore, until ATP hy-drolysis reestablishes the size of the pore, trapping the bulky amino acid either outside (v) or inside (ii) the membrane. This cycle produces a net forwards driving force due to the fact that the chunk at the channel exit does not trigger nucleotide exchange and thus channel opening, therefore the substrate is ratcheted in one direction (vi).[5]
3
AT O M I C F O R C E M I C R O S C O P Y ( A F M )
In early years, the term “microscope” was directly linked to optical or elec-tron microscopes that could provide a magnification of objects from 1000x to 100000x. Despite the technological breakthrough that they offered back at their time, they could only provide 2D information of the surface of the ob-ject, lacking the possibility to characterize the real dimensions of the sample itself or of the sample-support surface. In the early 1980’s, a new generation of microscopes was developed by Binnig and Roher, the scanning tunneling microscope. With this technique, a sharp tip in the range of several nm is scanning over the sample surface resulting in high resolution topographical images. The resolution of an STM microscope is 0.1 nm on the x-y axis and 0.01 nm in the z direction. Additionally, STM can be performed in ultra vac-uum, air, but also in water and in different buffer solutions that might imitate the physiological conditions. A very important technique based on STM and sharing its main functional principle is the atomic force microscope (AFM), which will be extensively discussed in the next sections as it is the main technique that was used within this study.
Principle of function
A nanometric-sized tip located at the very extremity of a cantilever comes into contact and interacts with the sample surface. While scanning the sample surface of interest, developed forces between the AFM tip and the surface are detected (fig.3.1).
Different types of interaction forces can be detected and quantified by AFM, such as electrostatic, adhesion, or Van der Waals forces. These interactions al-ter the motion of the cantilever which behaves according to Hooke’s law of elasticity F = −kcz, where F is the restoring force, kc the spring constant of the cantilever and z the cantilever displacement. A laser beam is constantly focused on the backside of the cantilever that is coated with a reflective ma-terial. Tip-sample interactions are causing a bending movement of the can-tilever and thus changes in the deflection of the laser signal, which is per-manently measured by a four-segment photodiode.[20] These changes are
fed into a feedback loop which keeps constant the applied force during scan-ning in contact mode, and the oscillation amplitude during tapping mode by adjustment of the tip-sample distance (section3.1).
3.1 a f m i m a g i n g
Various AFM imaging modes have been developed for the investigation of dif-ferent types of samples under difdif-ferent measurement conditions. The most commonly used modes are the contact mode, the intermittent contact
Figure 3.1: The Atomic Force Microscope. A laser beam is focused on the back of a reflective cantilever, carrying at its extremity a sharp tip. The motion of the cantilever upon interaction with the sample surface is detected by an optical beam deflection system (OBD; as detector a four-segment photo-diode is used). The detected signal is processed by a feedback loop. With appropriate computer software, the signal is converted into an image, providing the 3D topography of the sample.
ping) mode, the non-contact mode, and the MAC® mode (fig. 3.2). These
imaging modes will be discussed in the following.
Figure 3.2: Force regimes for the tip-sample interactions.[73]
Contact mode
During contact mode the AFM tip is in permanent contact with the sample surface and exerts a constant force on the sample. The developed repulsive forces are causing a bending of the cantilever while the tip senses changes in the sample topology. Contact mode is until today the most broadly used AFM
3.1 afm imaging 13
mode, mainly due to the simplicity of its function as the only parameter that requires a fine adjustment is the applied force to the sample. The feedback loop receives as input the deflection and maintains it constant throughout the acquisition by correcting the cantilever’s position in the z-direction. As the AFM tip is scanned in permanent contact over the sample, the resulting high shear forces can cause damages of the sample surface. For this reason, the contact imaging mode is mostly preferred for durable and robust samples, where sub-nanometer resolution can be achieved.[46] For soft and sensitive
samples such as biological samples, a more dynamic imaging more was de-veloped, the so-called intermittent contact or tapping mode.[76,63]
Tapping mode
During intermittent contact (tapping) mode, the cantilever is acoustically os-cillating close to its resonance frequency (typically ~5-150 kHz for conven-tional AFM cantilevers) and periodically touches the sample surface. Due to the short contact time this approach minimizes the shear forces as well as the risk of specimen damaging. In contrast to the contact mode where the feedback system maintains the deflection constant, in tapping mode the os-cillation amplitude is kept constant during imaging. The force applied to the sample is regulated through the setpoint value. The acoustic excitation arises from the alternating current (AAC) acting on a piezoelectric element placed on the cantilever holder, causing its vibration. Alternatively, the cantilever can be excited magnetically by application of magnetic alternating current. This excitation path led to the development of an alternative intermittent imaging mode, the MAC mode.
Magnetic AC mode (MAC)
For MAC® mode (Keysight, Santa Rosa, CA, USA) imaging the cantilever is magnetically excited by a coil either placed directly in the cantilever holder, either under the sample plate. As a consequence, cantilevers for MAC mode imaging need to be magnetically coated. Generally, MAC mode is considered to be a more handy and sensitive technique as a single prominent resonance peak is detectable during cantilever excitation,[56] whereas during AAC
tun-ing multiple peaks are generated.
Non-contact mode
As its name underlines, during this mode no contact is required between the tip and the sample surface. The major principle is based on the fact that non-contact mode is taking advantage of the weak attractive force interactions as depicted in fig.3.2, that occur at the tip-specimen interface (typically van der
Waals interactions). As these interactions are extremely short-ranged in liquid environment, the non-contact mode is rarely used for biological samples that need to be measured under physiological conditions.
3.2 s i m u lta n e o u s t o p o g r a p h y a n d r e c o g n i t i o n (trec) imaging In 2004, Stroh et al.[114] brought the high resolution AFM imaging one step
further by developing a revolutionary technique that can not only provide information about the topography of the sample surface but also about inter-molecular interactions between tip-anchored ligands and surface-immobilized receptors. For simultaneous topography and recognition (TREC) imaging, the AFM tip needs to be transformed into a biosensor by attaching the molecules of interest on the tip via a flexible PEG linker (detailed protocols will be pre-sented in section3.6). The functionalized cantilever is scanned over the
sam-ple surface in tapping mode (AAC or MAC), while the tip-attached molecule can rotate freely due to the flexibility provided by the PEG linker. Based on a special feedback loop, two images can be generated simultaneously, the clas-sic topographical image and the recognition image which depicts the specific receptor binding sites.
Figure 3.3: The principle of TREC imaging. The special electronics circuit of the Pi-coTREC® box divides the oscillation amplitude into a lower and an upper part. The upper part serves for recognition signal production whereas the lower part corresponds to the topographical image.
In more detail, the TREC mode works as following: The picoTREC® box (Keysight, formerly Agilent, Santa Rosa, CA, USA) receives the signal of the oscillation amplitude and separates it in two parts with respect to the zero position, whereas the upper part produces the recognition signal and the lower part the topographical signal (fig.3.3).[85,24,66] Higher structures on
the sample surface result in a reduction of the amplitude at the lower part whereas holes or surface gaps are increasing it. On the other hand, a specific binding of the tip-anchored ligand to the receptor causes a decrease of the amplitude at the upper part. This decrease can be explained by the fact that upon specific interaction the upward movement of the tip is limited by the length of the linker. The binding sites are depicted in the recognition image
3.3 afm-based single molecule force spectroscopy 15
as black spots at the same coordinates as the interacting receptor molecule in the topographical image. To check the specificity of the interaction, a block experiment should be performed, in which the binding sites are saturated and therefore no recognition signal will be produced.
The most important parameter that needs to be properly adjusted for TREC imaging is the oscillation amplitude, as proven by Preiner et al.[99] As shown
in fig.3.4, if small amplitudes are selected, the PEG linker is not sufficiently
stretched and therefore no reduction in the oscillation amplitude and hence no recognition signal is produced. In the other extreme case of selecting very high oscillation amplitudes, numerous unbinding events occur which lead to a unstable recognition signal production. The oscillation amplitudes should be carefully chosen by taking into account the length of the linker together with the size and position of the attached molecule. The fact that only a short range of oscillation amplitudes are appropriate for production of a TREC signal together with the performed block experiments provide the basis for a reliable specificity control. Attention should be paid to the fact that the ratio between the amplitude setpoint and the free amplitude should remain constant.
Figure 3.4: The importance of the oscillation amplitude for TREC imaging. The ideal oscillation amplitude has to be carefully predefined for TREC signal gen-eration. Image adapted from [99].
According to the study of Preiner et al., TREC imaging should be per-formed with half-amplitude feedback as in this way, the recognition signal is not causing any perturbation of the topography signal which would result in biased height values (fig. 3.5). In the same line, for better and more
ac-curate TREC imaging, driving frequencies below the cantilever’s resonance frequency at the tip-sample surface interface should be selected. This has been proven to create less undesired effects such as “cross-talk” phenomena between the topography and recognition image.
3.3 a f m-based single molecule force spectroscopy
AFM is one of the most versatile microscopic tools due to its capability to provide not only topographical images but the possibility to probe and char-acterize interactions on the single molecular level.[66,14] Information on the
life-Figure 3.5: Comparison of full and half amplitude TREC. Perturbations in the si-nusoidal cantilever movement due to surface features or interactions be-tween the ligand and the receptor can be temporally separated by the use of the lower part of the oscillation amplitude as feedback parameter (HA). Image adapted from [99].
times and energy landscapes can be obtained. The technique of AFM-based single molecule force spectroscopy (SMFS) is based on the following princi-ple: A functionalized AFM cantilever is approached to and retracted from the sample surface at constant velocity whereas the x-y coordinates remain fixed (movement only in the z-direction). The parameter of interest during AFM spectroscopy experiments is the deflection of the cantilever.
The first step for a successful spectroscopic experiment is the function-alization of the AFM tip. Three basic prerequisites should be fulfilled by the tip functionalization process: (i) The attached molecules should be pro-vided with sufficient mobility in order to be able to find and interact with its respective binding partner on the sample surface. For this reason, com-monly polyethyleneglycol (PEG) spacers are preferred that can be synthe-sized and adapted to each experiment’s needs, as functions and lengths can vary.[67,43, 45] (ii) Potential unspecific adhesion arising from the ligand
im-mobilization should be omitted or minimized, and (iii) the forces that keep the sensor molecule attached to the AFM tip have to be significantly higher compared to the interaction forces between the two interacting molecules. Even though the detailed tip-functionalization protocols will be presented in section3.6, it should be mentioned here that statistically only one molecule
is tethered to the very apex of the tip, making the investigation of 1:1 inter-actions possible.[46, 43] AFM-based force spectroscopy is a well established
technique not only on isolated surface-immobilized molecules,[33, 76] but
also on living cells.[126,100]
The combination of the cantilever and the linker molecule can be modeled by a system of two spring in series, with kc being the cantilever’s spring constant and klthe spring constant of the linker. The effective spring constant ke f f of this combination then follows to
1 ke f f = 1 kc + 1 kl (3.1)
3.3 afm-based single molecule force spectroscopy 17
The spring constant of the AFM cantilever might differ in reality from the nominal spring constant provided by the manufacturing company. The de-viations between the actual and the nominal ones might arise either from the manufacturing process itself either from the functionalization process during which different chemical components are acting on the cantilevers. Through years, a number of techniques have been developed in order to de-termine the spring constant of the used cantilever. These techniques can be grouped into two main categories: (i) dynamic methods (e.g., thermal noise method,[72, 19, 18] Sader method[107, 108]) and (ii) force-loading methods
such as piezosensor, nanoindentation calibration,[113, 70] and electrostatic
force balance (EFB) calibration[86]. The most commonly used technique and
the approach that was also used in this study is the thermal noise method as it is a fast, simple and accurate approach. Briefly, at first the ratio be-tween the measured deflection of the cantilever and the output of the optical photo-detector in Volt (V) needs to be quantified. This relation is known as “optical cantilever sensitivity” or simply “sensitivity” (nm/V). As next step, a spectrum of the amplitude versus frequency is created and the noise level can be defined. Subtracting the background noise and fitting the spectrum with the Lorentzian function for a simple harmonic oscillator results in the spring constant of the cantilever kc. The thermal noise method is based on the equipartition theorem according to which the thermal motion is related to the thermal energy by
kc = kBT
hz2i (3.2)
where kB equals 1.38·10−23 J/K (Boltzmann constant), T stands for tem-perature and z2 is the mean square of the cantilever’s displacement. As mentioned above, the thermal spectrum is fitted by a Lorentz fit for a simple harmonic oscillator according to the following equation
A= Awhite+
A0ω40
(ω2−ω20)2+ (ωω0/Q)2
(3.3) with Awhite being the white noise level, A0 the zero frequency amplitude,
ω0the radial resonance frequency, and Q the quality factor.[68] By integrating
without taking Awhite into account, the spring constant can be determined as[72]
kc = kBT
P (3.4)
with P representing the area of the power spectrum of the thermal fluctua-tions alone.[72]
For single molecule force spectroscopy experiments cantilevers with low spring constants (in the range of ~0.01 N/m) are commonly preferred as intermolecular forces are in the range of some tens of pN.
The Force - Distance Cycle (FDC)
The process of approaching and retracting the functionalized cantilever from the sample surface at a fixed lateral position while the cantilever’s deflection is continuously monitored results in a so-called force-distance cycle (FDC). A FDC can be easily separated in four major steps as depicted in fig.3.6. In the
beginning, the cantilever is approaching the sample surface and no contact or interaction between the two molecules occurs, resulting in no change in the cantilever’s deflection which remains constantly zero. The closer the can-tilever gets to the surface, different types of forces are developed. Attractive forces (e.g., van der Waals, electrostatic forces) will bend the cantilever down-wards and repulsive forces (e.g., electrostatic forces) will cause an upward bending of the flexible AFM cantilever. Higher attractive forces can force the cantilever to “jump into contact” even though there is still a certain distance between the tip and the sample. In the contact regime, a specific interac-tion between the tip-anchored ligand and the surface-immobilized receptor or unspecific adhesion of the tip can occur. Once a pre-defined force value (setpoint) is reached, the motion of the piezo changes direction and the can-tilever is retracted from the sample surface. It is generally suggested that the force setpoint should be kept as low as possible in order to prevent damages of the sample.
Figure 3.6: The distance cycle. Shown are the four typical parts of a force-distance cycle: In the beginning, no interaction occurs and the cantilever remains at its resting position. The approach continues until the contact point is reached. The deflection and therefore the force increases until the force setpoint is reached. The retraction begins and in case of a specific interaction the linker is getting stretched until a critical point at which the bond fails (unbinding event). The characteristic shape of the unbinding events arises from the non-linear stretching of the linker. Adapted from [120]
3.4 dynamic force spectroscopy 19
In case of an interaction between the tip-bound ligand and the surface-immobilized receptor, the linker starts to get more and more stretched until a critical force value is reached, at which the formed bonds breaks and the cantilever is returning to its resting position. This unbinding event can be easily discriminated from unspecific tip adhesion due to the parabolic shape of the force profile which arises from the stretching of the PEG linker. The height of the vertical jump directly reflects the unbinding force ( fu), and the distance between contact point and vertical jump yields the unbinding length. The conversion of the deflection voltage (V) to force values is possible via sensitivity determination. For this purpose, FDCs are performed on a stiff surface (usually glass). The slope in the contact regime is inversely propor-tional to the optical cantilever sensitivity (slope = sensitivity1 ). Multiplying the deflection (V) with the calculated sensitivity results in the displacement of the cantilever z (nm). As mentioned above, an AFM cantilever can be consid-ered as a spring, thus Hooke’s law can be used for force calculation as given in eq.3.5, with kcbeing the spring constant of the cantilever.
F=kcz (3.5)
Differently shaped curves might arise with respect to the measured inter-action. An overview is presented in fig. 3.7, whereas (A) shows a typical
unbinding event with a characteristic parabolic force profile. On the contrary, in the case of unspecific tip adhesion, no stretching of the linker molecule is observed and the response is linear (C). Both conditions can co-exist as shown in (D), where the adhesion and the unbinding event are spatially separated due to the linker stretching.
For a reliable and accurate analysis of force-spectroscopy measurements, a control experiment should be performed for validating the specific nature of the interaction. For such control experiments, either the surface receptors need to be inactivated by injection of a blocking agent into the measurement solution, either the ligand on the tip should me blocked by incubation of free receptors. In case that no suitable blocking agents exist, unfunctionalized AFM tips can be used instead. Upon blocking, the vast majority of specific interactions will disappear whereas it is still possible to observe adhesion. 3.4 d y na m i c f o r c e s p e c t r o s c o p y
Affinity between two molecules can lead to complex formation due to the thermal diffusion. As depicted in fig.3.8, the two states (bound and unbound
state) are separated by a sharp and well defined energy barrier. For associa-tion of the two molecules this energy barrier has to be overcome. The sum of the free energies of the two individual molecules is higher than the free energy of the complex as the thermal motion is contributing in finding their preferable energy state (∆G∗). The majority of the interactions developed
be-tween bio-molecules is shifting towards dissociation with time. The lifetime of the formed bond τ is depending on the dissociation rate ko f f via the fol-lowing relation:
Figure 3.7: Examples of force-distance cycle shapes. (a) Unbinding event and no ad-hesion, (b) no unbinding event and no adad-hesion, (c) no unbinding event but only adhesion, and (d) both unbinding event and adhesion. [68]
τ= 1
ko f f
(3.6) Therefore, the probability of the complex to remain in the bound state un-der the influence of a linearly increasing force is termed survival probability S(t), which follows a first order kinetics with a time-dependent rate[68]
dS(t) dt = −k(t)S(t) (3.7) and thus S(t) =exp − t Z 0 k(t0)dt0 (3.8)
The probability distribution(p(F))of the applied force F at rupture is related to S(t)via
p(F)dF= −S(t)dt (3.9) Solving the equation for p(F) results in
p(F) = k(F) dF dt exp − F Z 0 " k(F0) dF dt # dF0 (3.10)
3.4 dynamic force spectroscopy 21
As depicted in fig.3.8, according to Bell,[12] the presence of an external force
will tilt the energy landscape and lower the energy barrier. Hence, the activa-tion free energy under the influence of an external force∆G(F)is related to the initial free energy∆G according to the following equation
∆G(F) =∆G−Fxβ (3.11)
For AFM-based single molecule force spectroscopy a possible rebinding af-ter dissociation under applied force can be neglected, as both the ligand and the receptor are firmly attached either to the tip or the surface, respectively. Hence, the dissociation can be considered as an irreversible transition from the bound to the unbound state. During irreversible molecular transition un-der the influence of a force, it is assumed that the molecule moves along a combined free energy surface E(x) =E0(x) −F(x) in the direction of pulling x. The free energy E0(x) is considered to have a single well at x = 0 and a barrier of height ∆G at x= xβ.[68] The entire dissociation process is
consid-ered as a stochastic process due to the fact that the system is small and the surrounding heat bath is causing energy fluctuations. As can be seen in fig.
3.8, the external force acting on the system lowers the energy barrier, making
the dissociation by energy fluctuations more likely. Based on the theory of a single energy barrier as proposed by Bell[12] and adjusted by Evans,[50] the
increased rate of bond dissociation under external force can be described as
k(F) =ko f fexp(Fxβ/kBT) (3.12)
with ko f f being the kinetic off rate at zero force. In this case, xβ is assumed to
remain unaffected by the presence of external force.
In a single molecule force spectroscopy experiment, the retraction move-ment of the cantilever in z-direction occurs with a constant pulling velocity (v), yielding a linear force ramp
F(t) =kcvt (3.13)
with kc being the spring constant of the cantilever. For providing the ligand with a certain flexibility in order to find its binding partner on the surface, ligands are typically tethered to the AFM tip via a polyethylene glycol (PEG) linker. Therefore, it is not sufficient to consider kc alone but the spring con-stant of the linker molecule kl as well. The effective spring constant is then given as
k−e f f1 =k−c1+k−l 1 (3.14)
It has been shown,[51] that the spring constant of the linker kl is directly
related to the applied force. For this reason, the worm-like chain approxima-tion was applied to the force profile at the very moment of rupture for kl determination, resulting in a linear force ramp[50,55]
Figure 3.8: Energy profile of the dissociation process. The energy barrier without external force is shown by the solid black line. The activation energy∆G∗, the dissociation rate ko f f and the required spatial separation for transition
from the bound to unbound state, xβare depicted as well. An application of external force to the system (like it the case of SMFS) leads to a decrease of the energy barrier height (∆G∗(F)), as indicated by the black dashed
line. The lowering of the barrier makes the dissociation more likely as the separation barrier can be crossed more easily. (∆G∗ and ∆G∗(F) in this
figure correspond to∆G and ∆G(F)in the main text).[14,12]
The loading rate r, meaning the force load applied to the bond, is the product of the effective spring constant ke f f and the pulling velocity v (r=ke f fv).
The analytical expression for the observed rupture force can be obtained by substitution of eq.3.10and eq.3.12 into eq.3.15
p(F) = ke f f r exp Fxβ kBT − ko f fkBT rxβ expFxβ kBT −1 (3.16) The most probable unbinding force F∗ thereby scales linearly with the loga-rithm of the loading rate[50,12]
F∗= kBT xβ ln x βr kBTko f f (3.17) According to Bell and Evans,[50,12] the required force that induces
disso-ciation within a certain time t is critically depending on the time scale the ex-periments are performed. For AFM exex-periments this range is of milliseconds. Faster loading rates are resulting in shorter times meaning that higher forces are needed to disrupt the bond as the thermal activation is too slow to oc-cur in such limited time-frame. In case of slower loading rates, the provided times are longer than the natural dissociation time of the formed complex and consequently lower or zero forces are needed.[8] Reliable and accurate
results can be obtained only if a high amount of data is analyzed as the un-binding process is of stochastic nature. A typical data set (for one pulling
3.4 dynamic force spectroscopy 23
velocity) is comprised of more than 1000 force-distance cycles. The force dis-tributions can be determined either by the classical histogram representation either by calculation of the empirical probability density function (PDF). The advantage of the PDF is that it can be seen as a continuous histogram, tak-ing into account the uncertainty of each data point. In detail, a PDF can be calculated by summing up Gaussian distributions with unitary area that are calculated from the mean and variance of each unbinding event:[11,46]
pd f(F) = 1 N N
∑
i=1 1 q 2πσ2 i exp (F−Fi) 2 2σi2 ! (3.18) where N is the total number of unbinding events, Fi the measured unbind-ing forces and σi the corresponding standard deviations. The most probable unbinding force F∗ per loading rate r is determined by fitting a Gaussian function to the estimated PDF. To deduce information on bond lifetimes, dis-sociation rates and the width of the energy barrier xβ, a scatter plot of allun-binding forces against the logarithm of the loading rate (ln(r)) is created. The data cloud can be fitted with the linear Bell-Evans model,[12,50] whereas the
slope of the fit reveals information on the xβ value according to slope=
kBT
xβ and the dissociation rate ko f f can be extracted by extrapolation to zero force.
Due to the nature of the formed complex that involves different intermolec-ular interactions, it is possible that the dissociation process might require the crossing of intermediate barriers.[93] For more complex energy landscapes,
the Bell-Evans model assumes that multiple energy barriers result in multiple linear regimes in the loading rate dependence (LRD) plot. From the different linear regimes, the dissociation rate ko f f and the width of the energy barrier xβ for each of the crossed energy barriers (fig. 3.9) can be extracted.
Multi-ple energy barriers along the dissociation pathway lead to a comMulti-plex energy landscape with a highly discontinuous, non-linear LRD plot (fig.3.9).
The Bell-Evans model might be considered as a simplified representation of the overall dissociation process, however, it provides a reliable descrip-tion and estimadescrip-tion of the energy landscape and the kinetic parameters of bio-complex uncoupling.[14] Based on the work of Bell and Evans, different
models arose with non-linear LRD spectra prediction for a more detailed characterization of the dissociation process under the influence of an exter-nal force by taking additioexter-nal parameters into consideration. In 2006, Dudko, Hummer and Szabo modified the Bell-Evans model by estimating the effects of the external force on the position of the transition state.[37] In their model,
it has been proposed that the distance between the bound and the transition state is affected by the applied force.[37,38,39,36,40,41] Assuming the shape
of the overall energy landscape, they employed Kramers’ theory[84] for
diffu-sive barrier crossing. The introduced exponential constant v defines the shape of the free energy profile. For v = 1/2 the energy profile is parabolic with a cusp-like transition barrier, in case that v = 2/3 the energy profile can be imagined as linear-cubic. By v = 1, the linear Bell-Evans relation is restored. Similarly as Bell and Evans, also Dudko et al. are considering the dissociation process as irreversible. This assumption was addressed in 2012 by Friddle,
Figure 3.9: Energy landscapes of the dissociation process along a reaction coordinate with the corresponding loading rate dependence according to the Bell-Evans model. (A) A single energy barrier along a reaction coordinate of a formed complex results in a linear LRD plot. (B) The presence of a second energy barrier will create a second slope in the LRD plot. (C) The presence of multiple energy barriers yields a discontinuous LRD plot. Figure adapted from [14].
De Yoreo et al.,[54] who suggested that at low loading rates rebinding of the
two molecules is possible. According to their model, the dissociation of the complex passes through two distinct regimes: At first, at low loading rates a near-equilibrium regime is crossed, which arises either by rebinding of the two interacting molecules either by sequential rupture of individual bonds. At higher loading rates, the complex then dissociates irreversibly.
The selection of the most appropriate model is crucial for a correct char-acterization of the interaction, as parameters such as kinetic rates, bond life-times and information on the energy landscape are directly determined from the LRD fit. The Bell-Evans can provide reliable results on interactions where the unbinding forces are scaling linearly with the logarithm of the loading rate, whereas linear LRD plots should be fitted with an appropriate non-linear model. Useful information for the right selection of dynamic force spec-troscopy models can be found in the following references: [62,14,12, 50,93, 54,37].
3.5 c o m b i n e d t r e c i m a g i n g w i t h s i n g l e m o l e c u l e f o r c e s p e c -t r o s c o p y
High densities and homogeneous distributions of receptor molecules are re-quired for performing a successful force spectroscopy experiment as the tip is “blindly” approached to the sample surface making ligand-receptor interac-tions possible. Such condiinterac-tions are not always easy to obtain, especially when the molecules of interest are membrane proteins. It is often the case that only
3.6 tip chemistry 25
low reconstitution efficiencies can be obtained for protein integration into an artificial lipid membrane. To overcome this issue, an AFM-technique known as “force mapping” or “force volume” was proposed by Cleveland et al.[25]
In this mode, the cantilever is obtaining one or more FDCs at each posi-tion as it moves in the lateral direcposi-tion across the surface. Force mapping is a well established technique and has been already applied from simple biotin-avidin systems to organic materials and cells.[91, 78, 6, 27] The main
drawback of this handy technique is the acquisition time. The recquired time for obtaining such a map is directly linked to the selected parameters (image size, pixel resolution, number of FDC per pixel, FDC sweep time etc) and can range between minutes and several hours if a high resolution map is needed. Being aware of this issue, several groups tried to decrease the acquisition time, but for high resolution force-volume measurements still a few hours are required.[103, 1,7] In 2017, a combination of two techniques, TREC and
SMFS, was suggested by Köhler et al.,[82] which aims at overcoming this
problem. TREC and SMFS are two AFM techniques that ideally complement each other. TREC is an imaging technique that provides information on recep-tor binding sites and SMFS can fully characterize a molecular interaction by quantitatively determining its energetic and kinetic parameters. The advan-tage of TREC-driven force spectroscopy is that a single functional molecule can be selected and targeted for force spectroscopy. This approach was also deployed for many experiments presented in this study.
This combined technique requires the usage of the exact same cantilever for TREC imaging and SMFS. For TREC, stiffer cantilevers are selected with a spring constant in the range of 0.14-0.30 N/m,[114] whereas for SMFS softer
cantilevers are chosen (0.01-0.03 N/m) to be able to resolve forces of some tens of pN.[128] By using combined TREC imaging with SMFS, particular
care has to be taken as the position of the receptor of interest can be lost during force spectroscopy experiments due to drift. Therefore, a closed loop scanner has to be used and the environmental conditions (e.g., ambient tem-perature, temperature of the measurement buffer and the functionalized can-tilever) have to be adjusted and controlled carefully throughout the experi-ment. As an additional control, the FDC acquisition should be interrupted every ~200 curves and the exact position of the receptor on the sample sur-face should be verified again.
3.6 t i p c h e m i s t r y
Performing TREC imaging and AFM force spectroscopy experiments requires the functionalization of the AFM tip. Molecule attachment can be achieved either by absorption to the tip surface,[53] either by chemical
immobiliza-tion. [104] As mentioned in section 3.3, three main conditions should be
satisfied for a successful functionalization: (i) rotational freedom and flex-ibility should be provided to the ligand molecule, (ii) unspecific adhesion due to the immobilization method should be omitted, (iii) the forces that keep the molecules attached to the tip should be significantly higher than
