High-Resolution Imaging and Multiparametric Characterization of

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High-Resolution Imaging and Multiparametric Characterization of Native Membranes by Combining Confocal Microscopy and Atomic Force Microscopy-Based Multifunctional Toolbox Pawel R Laskowski, Moritz Pfreundschuh, Mirko Stauffer, Zöhre Ucurum, Dimitrios Fotiadis, and Daniel J. Müller ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b03456 • Publication Date (Web): 26 Jul 2017 Downloaded from http://pubs.acs.org on July 28, 2017

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High-Resolution Imaging and Multiparametric Characterization of Native Membranes by Combining Confocal Microscopy and Atomic Force MicroscopyBased Multifunctional Toolbox Pawel R. Laskowski,† Moritz Pfreundschuh,† Mirko Stauffer,‡ Zhöre Ucurum,‡ Dimitrios Fotiadis,‡ & Daniel J. Müller*,†

†

Department of Biosystems Science and Engineering, ETH Zurich, 4058 Basel, Switzerland; ‡

Institute of Biochemistry and Molecular Medicine, University of Bern, 3012 Bern, Switzerland

*Correspondence: [email protected]

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ABSTRACT To understand how membrane proteins function requires characterizing their structure, assembly and inter- and intramolecular interactions in physiologically relevant conditions. Conventionally, such multiparametric insight is revealed by applying different biophysical methods. Here we introduce the combination of confocal microscopy, force-distance curvebased (FD-based) atomic force microscopy (AFM) and single-molecule force spectroscopy (SMFS) for the identification of native membranes and the subsequent multiparametric analysis of their membrane proteins. As a well-studied model system, we use native purple membrane from Halobacterium salinarum, which membrane protein bacteriorhodopsin was His-tagged to bind nitrilotriacetate (NTA)-ligands. First, by confocal microscopy we localize the extracellular and cytoplasmic surfaces of purple membrane. Then, we apply AFM to image single bacteriorhodopsins approaching sub-nanometer resolution. Afterwards, the binding of NTA-ligands to bacteriorhodopsins is localized and quantified by FD-based AFM. Finally, we apply AFM-based SMFS to characterize the (un-)folding of the membrane protein and to structurally map inter- and intramolecular interactions. The multi-methodological approach is generally applicable to characterize biological membranes and membrane proteins at physiologically relevant conditions.

KEYWORDS: Chemical recognition imaging, fluorescence microscopy, force spectroscopy, ligand-binding, membrane protein, multiparametric imaging, single-molecule imaging


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Cellular membranes dynamically reassemble membrane proteins and lipids in order to guide their function.1,2 Depending on this heterogeneous assembly, inter- and intramolecular interactions change thereby modulating the functional state of membrane proteins. Hence, high-resolution imaging of membrane proteins in the native membrane and at physiologically relevant conditions along with quantifying their physical and chemical characteristics is necessary for the comprehensive understanding of how these proteins work and are regulated. A growing number of methods suitable for exploring the biophysical and biochemical properties of biological membranes and membrane proteins at highresolution, including optical microscopy and spectroscopy, electron microscopy and atomic force microscopy (AFM), has increased interest in the field.3-9 However, optimized approaches are necessary to mitigate the shortcomings of any individual method and to provide

multiparametric

information

on

biological

membranes

with

increasing

efficiency.3,6,9-11 Confocal microscopy and AFM are complementary. While fluorescence microscopy is routinely applied in life sciences to characterize fluorescently labelled samples, confocal fluorescence microscopy can be used to image complex biological objects in threedimensions. Conventionally, confocal microscopy can approach a vertical resolution of ≈ 500–800 nm and a lateral resolution of ≈ 200–500 nm. Invented more than 30 years ago, AFM enables to contour solid-state samples to atomic resolution.12-14 The combination of high-resolution AFM imaging with the possibility to use the AFM tip to manipulate matter from the microscopic to nanoscopic scale opened the door to the nanoworld and fundamentally changed the way how to approach and understand matter.15 Applied to biointerfaces, this approach to image and manipulate opens exciting possibilities to engineer complex systems and to guide and sense biological processes.16-19 In life sciences, AFM emerged as a powerful tool for imaging biological surfaces at physiologically relevant conditions at sub-nanometer resolution.3,9,20,21 To image biological surfaces and to simultaneously quantify their multiparametric information various different AFM-based imaging and spectroscopy modes have been developed and combied.9 These AFM-based modes include force-distance curve-based AFM (FD-based AFM),22-24 topography and recognition imaging,25,26 contact mode imaging at different membrane potentials,27 holographic imaging using ultrasonic waves28,29 and multifrequency imaging.30,31 So far,


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however,

the

simultaneous

high-resolution

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AFM

imaging

and

multiparametric

characterization of biological membranes has been experimentally rather complicated and thus restricted to experts.3,27,32-35 Recently, FD-based AFM has entered to a stage at which it can be used by non-experts for high-resolution imaging of native biological systems ranging from cells, membranes, proteins and nucleic acids.36 As most other AFMs, FD-based AFM uses a sharp tip at the end of a cantilever to contour the topography of a sample. However, to record a topographic pixel the AFM tip approaches and retracts to and from the sample at least once to record approach and retraction force-distance (FD) curves (Figure S1). Analysis of the FD curves allows extracting the mechanical properties of the biological sample, including stiffness, elastic modulus, adhesion, deformation and energy dissipation.37-40 Having thus analyzed the FD curves recorded for each topographic pixel, the extracted multiparametric properties are mapped to the topography of the sample. To further detect specific ligand-receptor binding events, FD-based AFM uses tips being functionalized with a ligand.3,24,41,42 While contouring membrane receptors, the approaching AFM tip continuously offers the receptor a ligand to bind, while FD curves recorded upon retracting the functionalized tip record the force required to separate the specific ligandreceptor bond.36 Using this approach, FD-based AFM has been applied to image native membrane receptors at high-resolution while structurally mapping their binding to ligands.41-43 So far, fluorescence microscopy and AFM have been frequently combined for the complementary imaging of living cells, cellular structures, membrane patches and of proteins.24,44-49 However, the possibility to optically image and identify proteins from native membranes and to characterize their physical and chemical properties using the nanoscopic AFM toolbox has not yet been demonstrated. Particularly, such multimicroscopic and multiparametric attempts could not yet reach sub-nanometer resolution. Here, we combine confocal microscopy and AFM to optically localize native membranes and thereafter image and analyze the properties of single membrane proteins by AFM-based methods. As a wellstudied model membrane, we used purple membrane from Halobacterium salinarium,50-52 which is naturally formed by lipids and the light-driven proton pump bacteriorhodopsin. Bacteriorhodopsin, which in native purple membrane assembles trigonal two-dimensional lattices, exposes the C-terminal end to the cytoplasmic surface and the N-terminal ends to


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the extracellular surface. After having localized individual purple membrane patches and identified their sidedness by fluorescence confocal microscopy we image both purple membrane surfaces using FD-based AFM. High-resolution AFM topographs approaching subnanometer resolution record typical structural details of individual bacteriorhodopsins. Using AFM tips functionalized with nickel-nitrilotriacetate (Ni2+-NTA) we then quantify and localize specific ligand-binding events to bacteriorhodopsin having His-tags engineered to the cytoplasmic C-terminal end. Finally, we apply AFM-based single-molecule force spectroscopy (SMFS) to characterize the folding of the membrane protein and the interaction forces stabilizing the protein. The multi-instrumental approach is applicable to image biological membranes from the micro- to the nanoscopic range and to characterize the complex relationship of the folding, structure, and function of native membrane proteins in the physiological relevant environment.

RESULTS and DISCUSSION Imaging purple membrane by confocal microscopy and FD-based AFM. For the multiparametric analysis of native purple membrane we mounted an AFM on top of an inverted confocal microscope (Figure 1a). The setup was isolated from noise by an active damped table, which has been placed in a temperature controlled and noise-adsorbing chamber (Methods). In our experiment, we wanted to first image purple membrane using confocal microscopy and to identify purple membranes exposing the cytoplasmic and the extracellular surface. After this, we wanted to image both purple membrane surfaces at subnanometer resolution by FD-based AFM (Figure S1). High-resolution AFM imaging is sensitive to mechanical vibrations of the sample support,12 which particularly occur using thin glass coverslips for fluorescence microscopy.53,54 Thus, to reduce the vibrations and to acquire fluorescence images at high magnification (objective 63x, NA 1.20) we used thin glass coverslips (≈ 0.15 mm), which we stabilized by two 1 mm thick glass slides glued on the sides of the coverslip. Onto the glass coverslips we glued a very thin layer of mica (≈ 0.05 mm), which is a well-established atomically flat sample support for the high-resolution AFM imaging of biomolecules.55-57 To fluorescently label the extracellular purple membrane surface, we engineered a His10-tag to


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the extracellular N-terminal end of bacteriorhodopsin (Methods and Figure S2). To suppress unspecific adsorption of fluorescently labelled ligands supposed to bind purple membrane, the freshly cleaved mica support was first coated with Ni2+-NTA-Atto 647 diluted in adsorption buffer (300 mM KCl, 20 mM HEPES, pH 7.4). This fluorophore, which was not excited by the wavelength used for confocal microscopy, served to block the unspecific adsorption of very similar fluorophores used later on to specifically label purple membrane. Then, purple membrane having His10-tagged extracellular surfaces was adsorbed to the support for ≈ 20 min in adsorption buffer.56 Thereafter, the fluorescently labelled NTAligand (Ni2+-NTA-Atto 488) was added to the adsorption buffer to specifically label His10tagged purple membrane surfaces exposed to the solution (Figure 1b). Then, the sample was washed ten times with imaging buffer (150 mM KCl, 20 mM HEPES, pH 7.4) and imaged by confocal microscopy, whereas the purple membrane patches exposed uniform fluorescence (Figure 1c). Guided by the confocal images the AFM cantilever was positioned above the purple membrane patches and the sample was imaged by FD-based AFM. The AFM topographs recorded from the same area as imaged by confocal microscopy showed purple membrane patches protruding 7.3 ± 0.3 nm (mean ± sd; n = 102) from the supporting mica (Figure 1d). Because the structural thickness of purple membrane corresponds to ≈ 6.0–6.5 nm,58 the higher value measured by AFM likely originates from electrostatic interactions between the charged purple membrane surface and the AFM tip.59,60 Nevertheless, the comparison of confocal image and AFM topograph showed that the fluorescence co-localized with some purple membrane patches imaged by AFM but not with all (Figure 1e). Since after adsorption of purple membrane to mica only membranes exposing the His10-tagged extracellular surface could bind the fluorescent NTA-ligand, we concluded that the fluorescent purple membranes exposed their extracellular surface to the AFM tip whereas the non-fluorescent ones exposed their cytoplasmic surface. Inspection of the AFM topographs showed that purple membrane showed the typical smooth appearance at the extracellular surface and a rougher appearance at the cytoplasmic surface (Figure S3).61,62 As a control we engineered purple membrane exposing His5-tags at the C-terminal end of bacteriorhodopsin and thus at the cytoplasmic surface. After adsorption of the purple membrane to Ni2+-NTA-Atto 647-coated mica, we incubated the sample with Ni2+NTA-Atto 488. The latter fluorophore was excited by the wavelength of ≈ 488 nm used for confocal microscopy. At this wavelength, the Ni2+-NTA-Atto 647 fluorophore, used to block


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the unspecific adsorption of Ni2+-NTA-Atto 488 to mica, was not excited. Confocal microscopy now showed fluorescent purple membrane patches exposing the cytoplasmic surface (Figure S4). Correlation of fluorescence image and AFM topograph revealed that purple membrane patches exposing the extracellular surface to the buffer solution were not fluorescently labelled. Furthermore, control experiments showed that wild-type purple membrane patches not carrying a His-tag were not labelled by the fluorescent Ni2+-NTAligand (NTA-Atto 488) (Figure S5).

High-resolution AFM imaging of purple membrane. In above experiments, we have identified purple membrane patches exposing the extracellular surface and patches exposing the cytoplasmic surface. Next, we wanted to see whether we could use the microscopy setup to image both purple membrane surfaces at high-resolution by FD-based AFM. We hence adsorbed purple membrane to mica, fluorescently labelled the His5-tagged cytoplasmic surface, and imaged the sample by confocal microscopy and AFM (Figure 2a,b). For high-resolution AFM imaging, we zoomed in on either membrane surface and increased the number of pixels recorded per nm2. The maximal force at which the non-functionalized AFM tip (Method) scanned the membrane was limited to ≈ 120 pN to prevent the mechanical deformation of bacteriorhodopsin37,63 and the parameters of the AFM feedback loop were optimized to reduce the error signal40. At this condition, the assembly of the bacteriorhodopsin trimers became visible and the topographs revealed structural details of the extracellular and cytoplasmic purple membrane surfaces (Figure 2c,e). These structural details were similar to those routinely observed by high-resolution contact-mode,20,61 oscillation mode,64,65, multifreqency30 and FD-based37,40 AFM imaging. Correlation averaging of the bacteriorhodopsin trimers further increased the signal-to-noise ratio of the topographs and the structural details of bacteriorhodopsins forming the trimer became visible. Whereas at the extracellular surface the trimers exposed the typical tripartite arrangement of bacteriorhodopsin (Figure 2c,d), they displayed their typical donut-like assembly at the cytoplasmic surface (Figure 2e,f).66 The trimer to trimer distance of 6.3 ± 0.2 nm (n = 34) was in agreement with structural data reporting ≈ 6.2 nm58,67. Bacteriorhodopsin protruded 0.5 ± 0.1 nm (n = 60) from the extracellular surface and


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0.8 ± 0.2 nm (n = 60) from the cytoplasmic surface of the lipid bilayer, which matched values previously measured by contact mode and FD-based AFM.20,37,66

Affinity imaging with functionalized AFM tips. Next, we wanted to use our combinatorial approach to image purple membrane by confocal microscopy and FD-based AFM to detect and morphologically map ligand-binding events. In a first step we adsorbed purple membrane, which bacteriorhodopsins carried a His5-tag at the C-terminal end, to mica. The sample was then fluorescently labelled with Ni2+-NTA-Atto 488 and imaged by confocal microscopy (Figure 3a) and FD-based AFM (Figure 3b). The AFM topograph showed two purple membranes, from which the confocal image showed one membrane labelled. The fluorescently labelled purple membrane thus exposed the cytoplasmic surface, whereas the not labelled purple membrane exposed the extracellular surface (Figure 3c). The binding between the NTA-ligand and His-tag depends on the presence of Ni2+. The His5-tag-Ni2+-NTA bond dissociates rapidly in the presence of competitors (imidazole) and the linkage ceases in the presence of agents chelating divalent ions such as ethylenediaminetetraacetic acid (EDTA).68 Thus, after having imaged the His5-tagged purple membrane labelled with Ni2+-NTA-Atto 488, we dissociated the NTA-ligand and washed it away with imaging buffer supplemented with 500 mM imidazole (150 mM KCl, 500 mM imidazole, 20 mM HEPES, pH 7.4). To functionalize AFM tips, we covalently tethered 18-unit long heterobifunctionalized polyethylene glycol linkers (PEG18) to the amino functionalized AFM tip and attached tris-nitrilotriacetic acid (tris-NTA) functional groups to the free end of the linkers (Figure 3d and Figure S6). Fully stretched, the linker system consisting of PEG18 and NTA-ligand is ≈ 7 nm long (Figure S6b). Using the functionalized AFM tip, we imaged the same purple membrane in affinity buffer (120 mM KCl, 5 mM NiCl2, 20 mM HEPES, pH 7.4) (Figure 3e and Figure S6a) and recorded a FD curve for each topographic pixel. The oscillation frequency was reduced to 0.25 kHz to provide a prolonged interaction time (≈ 1 ms) of the NTA-ligand tethered to the AFM tip with the His5-tag fused to the C-terminal end of bacteriorhodopsin. To display the adhesive interactions recorded in these FD curves in an adhesion map we applied a rupture distance- and adhesion force-filter to the retraction force curves to separate specific from unspecific interactions (Figure S6c). This filter discarded adhesive forces occurring at tip-sample distances < 3 nm, which could


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account for unspecific interactions in the contact region of tip and sample, and counted the specific adhesive events detected within a tip-sample distance of 3 and 10 nm, which describes the mechanical stretching of the linker system tethering the NTA-ligand to the AFM tip.3,36 The adhesion force filter accommodated for selecting rupture force curves that showed a higher adhesion forces than the usually observed thermal noise of the free cantilever. The resulting adhesion map showed a distinct difference between purple membranes exposing the cytoplasmic surface and membranes exposing the extracellular surface (Figure 3f and Figure S7b,c). Whereas the cytoplasmic purple membrane surface, which carried the His5-tag, showed most adhesion events, the extracellular purple membrane surface showed almost four times fewer events (Figure 3f and Figure S7b-e). The mean rupture force measured for the His5-tag-Ni2+-NTA bond formed at the cytoplasmic purple membrane surface was 154 ± 38 pN (n = 196, mean ± sd) at loading rates of 1.8 ± 0.7 × 106 pN s-1 (mean ± sd). This rupture force lied well within the ranges measured earlier using AFM-based force spectroscopy to detect the strength of His-tag-Ni2+-NTA bonds68-73. However, when comparing the rupture forces of His-tag-Ni2+-NTA bonds one must consider that the strength of the bonds depends on the cantilever spring constant and therefore the loading rate applied74,75, on the support to which the His-tag has been tethered,76 and on the number of histidines forming the His-tag77. As a negative control experiment we re-imaged the same purple membrane patches using the functionalized AFM tip but in the presence of EDTA (Figure 3g and Figure S7f-h). Most of the adhesion events (≈ 80%) previously detected on the cytoplasmic surface disappeared indicating that the adhesion was due to the specific His5-tag-Ni2+-NTA interactions.

Characterizing the folding and stability of bacteriorhodopsin. We combined confocal microscopy and FD-based AFM to image and identify native protein membranes, to detect their sidedness, to image membrane proteins at high-resolution and to detect their specific binding to a ligand. Next, we wanted to characterize the folding of the proteins and the inter- and intramolecular interactions stabilizing their structure and functional state using AFM-based SMFS.4,78,79 For this purpose, confocal microscopy was used to identify purple membrane exposing the extracellular surface, which had been fluorescently labelled by Ni2+-


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NTA-Atto 488 binding to N-terminally His10-tagged bacteriorhodopsin. Thereafter, purple membrane was imaged by FD-based AFM in unfolding buffer (300 mM KCl, 10 mM Tris, pH 7.4). To attach the terminal end of single bacteriorhodopsins a non-functionalized AFM tip was brought into mechanical contact with the non-fluorescent cytoplasmic surface of purple membrane by applying a force of 750 pN for 0.5 s (Methods). As reported earlier this mechanical contact favored the nonspecific adhesion of the terminal end to the AFM tip.78,80 Upon retraction of the AFM tip, we recorded a FD curve, which showed a distinct pattern of adhesive force peaks (Figure 4a). Such sawtooth-like force peak patterns are characteristic for the mechanical unfolding of membrane proteins. The force peak pattern extending to a distance of > 60 nm indicated that the bacteriorhodopsin adhering to the AFM tip was fully unfolded and extracted from purple membrane.80 We repeated the single-molecule experiments many hundred times to gain sufficient amount of FD curves. In ≈ 1% of the attempts (n = 45’056) the FD curves showed a characteristic sawtooth-like force peak pattern each recording the mechanical unfolding of a single bacteriorhodopsin from the Cterminal end. The superimposition of 183 FD curves showed a highly reproducible force peak pattern. Each force peak of the pattern signified one unfolding step of bacteriorhodopsin, with all force peaks together resembling the unfolding intermediates along the mechanical unfolding pathway of the membrane protein. Next, we fitted the force peaks using the worm-like chain (WLC) model to determine the contour lengths of the polypeptide stretches unfolded in each step (Figure 4b).78 These contour lengths were used to

localize

the

unfolding

steps

and

thus

the

unfolding

intermediates

of

bacteriorhodopsin.78,81 Altogether we identified five main unfolding peaks and three side peaks corresponding to the stepwise unfolding of individual α-helices of bacteriorhodopsin from purple membrane (Figure 4c), which was consistent with previous studies.78,80 The force measured to mechanically induce the unfolding of the secondary structures of bacteriorhodopsin was 144 ± 54 pN (n = 203) for the second main unfolding peak at a contour length of ≈ 87 amino acid (aa) and 87 ± 27 pN (n = 198) for the last main unfolding peak at a contour length of ≈ 223 aa, which is in agreement with the published results.82 On the example of more than 15 different membrane proteins, it has been shown that the mechanical unfolding force peak pattern is specific to the membrane protein.4,79 However, it has been also shown that the pattern changes if the membrane protein


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destabilizes or misfolds and even if the membrane protein changes its functional state.83-87 Once the mechanical unfolding pattern of a membrane protein has been identified, it can be taken to identify the membrane protein mechanically extracted from the protein membrane and of its functional state.4,79,88 Thus, the consistency between the force peak pattern recorded here for His-tagged bacteriorhodopsin and that recorded previously for native bacteriorhodopsin indicates that the bacteriorhodopsin characterized in our setup shows the native mechanical stability and fold.

CONCLUSIONS The combination of confocal microscopy and of various AFM-based methods introduced here provides a multifunctional toolbox for the optical imaging and identification of native membranes, for the imaging of surface structures of membrane approaching subnanometer resolution, for the localization and quantification of single ligand-receptor binding events, and for the characterization of the fold and interactions stabilizing single membrane proteins. Our instrumental approach can be readily applied to characterize and manipulate a wide range of native membranes and membrane proteins from the micrometer to the (sub-)nanometer scale. However, as we here used purple membrane as a model-system, which contains only the membrane protein bacteriorhodopsin, the challenge will be to identify the several hundreds of different proteins shaping a more complex environment such as described for membranes from plant, bacterial and animal cells. Ultimately, functional FD-based imaging may help to identify some of these proteins and to learn how they dynamically assemble into functional domains. Revealing even deeper insight, AFM-based SMFS may be used to study how this assembly modulates the fold, stability and functional state of membrane proteins.84,89,90


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METHODS Engineering, Homologous Expression, and Isolation of

His10- and His5-tagged

Bacteriorhodopsin from Halobacterium salinarum. A N-terminal His10-tag was engineered between the signal peptide sequence and the bacteriorhodopsin in three steps: First, two polymerase chain reaction (PCR) reactions were performed in parallel using the gene encoding bacterio-opsin (bop) and i) the forward primer 5’-AAA AGG ATC CGA CGT GAA GAT GGG GC-3’ and reverse primer 5’-TGA TGA TGG TGG TGA TGT CCG GTG ATC TGG GCC T-3’, and ii) the forward primer 5’-CAT CAC CAC CAT CAT CAC CAT CAT CAC CAC ACC GGA CGT CCG GAG TGG-3’ and reverse primer 5’-AAA AAA GCT TGA TTC AGT CGC TGG TCG CGG CC3’. Second, products from the two PCR reactions were pooled and submitted to PCR annealing. Third, the product from the previous annealing reaction was amplified by PCR using the forward primer 5’-AAA AGG ATC CGA CGT GAA GAT GGG GC-3’ and reverse primer 5’-AAA AAA GCT TGA TTC AGT CGC TGG TCG CGG CC-3’. The C-terminally His5-tagged bacteriorhodopsin version was engineered and cloned as described.91 Both genetically engineered bacteriorhodopsin versions were cloned into the shuttle plasmid pHS blue using the restriction enzymes BamHI and HindIII, and transformed and expressed homologously in the bacteriorhodopsin-deficient H. salinarium strain L33 according to Gordeliy et al.91 Purple membranes containing N-terminally His10- or C-terminally His5-tagged bacteriorhodopsin were isolated as described43,91,92. Compared to the N- and C-termini of wild-type bacteriorhodopsin,

i.e.,

NH2-MLELLPTAVEGVSQAQITGRPEW...-COOH

and

NH2-

...LRSRAIFGEAEAPEPSAGDGAAATSD-COOH, the termini of the two engineered constructs were NH2-MLELLPTAVEGVSQAQITGHHHHHHHHHHTGRPEW...-COOH (N-terminally His10tagged bacteriorhodopsin) and NH2-...LRSRAIFGEAGHHHHH-COOH (C-terminally His5-tagged bacteriorhodopsin) (fused portions in bold). Purple Membrane Preparation for AFM. Sample support preparation involved punching round mica disks (≈ 0.05 mm thickness, 9.5 mm diameter), punch set ‘punch and die’, Precision Brand Products) and Teflon rings of a slightly bigger radius (inner diameter 11.1 mm, outer diameter 15.9 mm). Teflon rings and mica disc were subsequently glued on top of 0.15 mm thick glass coverslips (24 x 60 mm coverslips, Thermo Scientific, Germany) using a two-component transparent glue (Araldite Crystal, Araldite) such that the mica sheet lay inside the Teflon ring. Next, two pieces of 1 mm thick glass slides (26 x 76 mm2 microscope


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slides, Thermo Scientific) cut to 26 x 26 mm2 squares were glued to the prepared supports, adjacent to the Teflon ring on both sides in order to limit vibrations of the glass coverslip during AFM imaging. The supports were dried for at least 24 h prior to experiments. Freshly cleaved mica surfaces were incubated with 100 µL of Ni2+-NTA-Atto 647 (Sigma Aldrich, USA) (2.5 μg mL–1) diluted in fresh adsorption buffer (300 mM KCl, 20 mM HEPES, pH 7.4) for 30 min and unbound dye was washed away five times with 100 µL adsorption buffer. Purified C-terminal His5-tagged or N-terminal His10-tagged bacteriorhodopsin/purple membrane was diluted to 0.02 mg ml–1 in adsorption buffer. 70 μl of purple membrane solution was adsorbed on mica surfaces for 20 min. Unbound purple membrane was washed away by gently pipetting the droplet up and down five times. Next, adsorbed purple membrane was incubated with 100 μl of 100 ng ml–1 Ni2+-NTA-Atto 488 diluted in adsorption buffer for 1 h in darkness. Unbound dye was washed away ten times using 100 µL adsorption buffer. Before imaging, the sample was washed ten times with 100 µL imaging buffer (150 mM KCl, 20 mM HEPES, pH 7.4). Buffers were prepared using nanopure water (18 MOhm cm–1) and analytical grade chemicals. All preparation steps were performed at room temperature. Before affinity imaging, the sample was washed five times with 100 µL imaging buffer supplemented with 500 mM imidazole (150 mM KCl, 500 mM imidazole, 20 mM HEPES, pH 7.4) and incubated in the same buffer for 30 min. Functional AFM imaging was performed in affinity buffer (120 mM KCl, 5 mM NiCl2, 20 mM HEPES, pH 7.4), while negative controls were carried out in imaging buffer supplemented with 10 mM EDTA (150 mM KCl, 10 mM EDTA, 20 mM HEPES, pH 7.4). AFM Tip Functionalization. To functionalize the Si3N4 cantilever and tip we followed a previously described protocol (Figure S6a).93 Briefly, AFM cantilevers (AC-40, Bruker) were treated with ultraviolet radiation and ozone cleaning (Jetlight, USA) for 10 min. For the amino-functionalization, AFM tips were immersed in an 8.1 M ethanolamine solution for 16 h. Tips were cleaned three times for 1 min in dimethyl sulfoxide (DMSO) and then for 2 min in ethanol, rinsed with ethanol and dried with filtered N2. For the linker attachment, the tips were immersed in a solution of acetal-PEG18-NHS (1 mg dissolved in 0.5 ml chloroform (CHCl3), Gruber lab, JKU Linz, Austria) and 30 µL triethylamine (Et3N) for 2 h. Tips were then cleaned in CHCl3 (3 x 10 min) and dried with filtered N2. The cantilevers were immersed in 1 mL of citric acid (1%, w/v) for 10 min. After washing in ultrapure H2O (3 x 5


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min), cantilevers were dried with filtered N2. A 1 M sodium cyanoborohydride (NaBH3CN) solution was prepared by adding 13 mg of NaCNBH3 to 20 µL of 100 mM NaOH and then mixed with 180 µL of ultrapure water. The cantilevers were immersed in a 100 µL droplet of the 0.5 mM amino-tris-NTA molecule (αS,α’S,α’’S)-11-(6-amino-1-oxohexyl)-α,α’,α’’tris[bis(carboxymethyl)amino]-δ,δ’,δ’’-trioxo-1,4,8,11

tetraazacyclotetradecane-1,4,8-

tripentanoic acid (Toronto Research Chemicals) in phosphate buffered saline solution (PBS). 2 µL of the cyanoborhydride solution was added and mixed carefully. The cantilevers were incubated for 2 h. The reaction was quenched by adding 5 µL of 1 M ethanolamine (pH 8.0) for 10 min. The cantilevers were washed in PBS three times for 5 min and used within 48 h. Tips functionalized with the tris-NTA-ligand were immersed in affinity buffer for 20 min prior to use. Optical Imaging. Samples were imaged by inverted confocal microscopy (LSM 800, Carl Zeiss Germany) using a 10 mW, 488 nm laser at 4–8% power and a 1 airy unit pinhole. Images were acquired with a 63× water immersion lens (421787-9970-799 objective, NA 1.20, Carl Zeiss). All imaging was performed at room temperature. Combined AFM and Confocal Microscopy Setup. An AFM (BioScope Resolve, Bruker) was installed on a stage of an inverted confocal microscope (LSM 800, Carl Zeiss) (Figure 1 and Figure S1) and operated with NanoScope 9.2R1 software (Bruker). FD-based AFM imaging was conducted in the ‚Peak Force QNM’ mode. The AFM and was equipped with a 100 x 100 x 15 µm (x, y, z) piezoelectric scanner. Rectangular (AC40, Bruker) and triangular (ScanAsyst Fluid+, Bruker) AFM cantilevers were used. Cantilevers had nominal spring constants of 0.09 and 0.7 N m–1 and resonance frequencies in water of 25 and 150 kHz, respectively. The AFM cantilevers were calibrated using thermal tuning and by ramping on a solid surface. Images were obtained with AC40 AFM cantilevers applying 200 pN imaging force, 2 kHz oscillation frequency, 1 Hz scanning rate and 60 nm amplitude, unless otherwise stated. For high-resolution imaging ScanAsyst Fluid+ AFM cantilevers were used. Parameters were adjusted to an oscillation frequency of 2 kHz, a contact force of ≈ 120 pN and an amplitude of 16 nm. An area of ≈ 150 x 150 nm was scanned at 512 x 512 pixels and a scan rate of 0.8 Hz. In order to detect His5-tag-Ni2+-NTA interactions, Ni2+-tris-NTA-functionalized tips were used in affinity imaging buffer. Affinity imaging was performed with 0.25 kHz oscillation frequency. Scanning rates and feedback gains were iteratively optimized to


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achieve low contrast of the error signal. Images were recorded at 256 x 256 pixels at a scanning rate of 0.125 Hz and FD curves were saved for subsequent data analysis. We kindly note that the scanning parameters, which we optimized for our experimental system, may be different for other receptor-ligand pairs and experimental setups (i.e., AFM, cantilevers, sample, support, and buffer solution). SMFS. Mechanical unfolding of bacteriorhodopsin was conducted in the ’FAST Force Volume’ mode (Bruker). In this mode, the non-functionalized tip of the AFM cantilever (AC40, Bruker) was used to localize purple membrane patches exposing their cytoplasmic surface. Then, the bare tip was approached onto the cytoplasmic purple membrane surface. After a set trigger force value has been reached (≈ 750 pN) the tip was held in contact with the sample for 0.5 s, which facilitated the bacteriorhodopsin polypeptide to adhere nonspecifically to the AFM tip.78,94 Next, the tip was retracted for a distance of 250 nm at a velocity of 1 µm s–1. After a force curve has been recorded for each approach and retraction cycle of the AFM tip, the tip was moved to an adjacent position of the purple membrane surface to record the next approach and retraction cycle. SMFS was performed in unfolding buffer (300 mM KCl, 10 mM Tris, pH 7.4). FD Curve and Image Analysis. FD curves from affinity imaging were analyzed both with commercial software (NanoScope Analysis v.1.7, Bruker) and with MatLab (MathWorks, USA) scripts developed in-house. Adhesion events were identified as specific when they occurred at distances 3–10 nm from the contact area. The standard deviation (sd) of the baseline of the FD curve was calculated and only adhesion events showing forces higher than five times the sd were taken into account (Figure S6c). FD curves were processed as described78,95 and analyzed using an automated approach96. Briefly, the FD curve with the highest similarity to all other FD curves was identified with the Euclidean distance correlation against all curves and was used to align the rest of the curves. The force peaks in every FD curve were then recognized using noise reduction and fitted with the worm-like chain model. The resulting high density force/contour length value pairs were clustered and identified using the DBSCAN algorithm.97 For every cluster the mean contour length was calculated. High-resolution threefold symmetrized correlation averages were calculated using the SEMPER software.98


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ACKNOWLEDGEMENTS We are very thankful to J. Thoma for contributing to SMFS data analysis, M. Tikhomirova for helping to develop the Matlab script used for analyzing the specific adhesion data recorded by FD-based AFM. We thank E. Mulvihill for assistance in high-resolution FD-based AFM imaging and R. Newton for critically reading of the manuscript. We thank U. Eidhoff for providing the plasmid pHS blue and the BR-deficient H. salinarium strain L33. The work was supported by the Swiss National Science Foundation (Grants 310030B_160225 and 205320_160199 to D.J.M.) and the NCCR Molecular Systems Engineering (to D.J.M. and D.F.).

ASSOCIATED CONTENT Supporting

Information

Available:

Principles

of

FD-based

AFM,

Biochemical

characterization of His5- and His10-tagged bacteriorhodopsin and comparison with wild-type bacteriorhodopsin by SDS-PAGE and Western blot analysis, fluorescence microscopy and topographical differences between cytoplasmic and extracellular surfaces of purple membrane, optical identification of fluorescently labelled purple membrane after adsorption to supporting mica, fluorescence images and AFM topographs of non-tagged wild-type purple membrane adsorbed to mica, strategies of Functionalizing AFM tips to detect specific ligand-binding events and analysis of the force-distance curves, separation of specific and unspecific adhesion events detected while imaging purple membrane, and quantification of mean forces stabilizing structural segments of N-terminally His10-tagged bacteriorhodopsin mechanically unfolded from the C-terminal end. The authors declare no competing financial interest.

AUTHOR INFORMATION Corresponding author *Email: [email protected]


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ORCID Daniel J. Müller: 0000-0003-3075-0665

Author Contributions D.J.M., D.F., P.R.L. and M.P. designed the project and contributed to the experimental design. M.S. and D.F. genetically modified bacteriorhodopsin, and produced and purified His-tagged purple membranes. M.P. and P.R.L. designed and performed the AFM probe functionalization. P.R.L. performed the confocal microscopy and AFM experiments, extracted and analyzed the data. All authors wrote the paper.


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(75) Friddle, R. W.; Noy, A.; De Yoreo, J. J., Interpreting the Widespread Nonlinear Force Spectra of Intermolecular Bonds. Proc. Natl. Acad. Sci. U S A 2012, 109, 13573-13578. (76) Nieba, L.; NiebaAxmann, S. E.; Persson, A.; Hamalainen, M.; Edebratt, F.; Hansson, A.; Lidholm, J.; Magnusson, K.; Karlsson, A. F.; Pluckthun, A., Biacore Analysis of HistidineTagged Proteins Using a Chelating NTA Sensor Chip. Anal. Biochem. 1997, 252, 217-228. (77) Knecht, S.; Ricklin, D.; Eberle, A. N.; Ernst, B., Oligohis-Tags: Mechanisms of Binding to Ni2+-NTA Surfaces. J. Mol. Recognit. 2009, 22, 270-279. (78) Muller, D. J.; Engel, A., Atomic Force Microscopy and Spectroscopy of Native Membrane Proteins. Nat. Protoc. 2007, 2, 2191-2197. (79) Bippes, C. A.; Muller, D. J., High-Resolution Atomic Force Microscopy and Spectroscopy of Native Membrane Proteins. Repo. Progr. Phys. 2011, 74, 086601. (80) Oesterhelt, F.; Oesterhelt, D.; Pfeiffer, M.; Engel, A.; Gaub, H. E.; Muller, D. J., Unfolding Pathways of Individual Bacteriorhodopsins. Science 2000, 288, 143-146. (81) Muller, D. J.; Kessler, M.; Oesterhelt, F.; Moller, C.; Oesterhelt, D.; Gaub, H., Stability of Bacteriorhodopsin Alpha-Helices and Loops Analyzed by Single-Molecule Force Spectroscopy. Biophys. J. 2002, 83, 3578-3588. (82) Janovjak, H.; Kessler, M.; Oesterhelt, D.; Gaub, H.; Muller, D. J., Unfolding Pathways of Native Bacteriorhodopsin Depend on Temperature. EMBO J. 2003, 22, 5220-5229. (83) Kedrov, A.; Krieg, M.; Ziegler, C.; Kuhlbrandt, W.; Muller, D. J., Locating Ligand Binding and Activation of a Single Antiporter. EMBO Rep. 2005, 6, 668-674. (84) Zocher, M.; Fung, J. J.; Kobilka, B. K.; Muller, D. J., Ligand-Specific Interactions Modulate Kinetic, Energetic, and Mechanical Properties of the Human Beta2 Adrenergic Receptor. Structure 2012, 20, 1391-1402. (85) Thoma, J.; Burmann, B. M.; Hiller, S.; Muller, D. J., Impact of Holdase Chaperones Skp and Sura on the Folding of Beta-Barrel Outer-Membrane Proteins. Nat. Struct. Mol. Biol. 2015, 22, 795-802. (86) Serdiuk, T.; Madej, M. G.; Sugihara, J.; Kawamura, S.; Mari, S. A.; Kaback, H. R.; Muller, D. J., Substrate-Induced Changes in the Structural Properties of LacY. Proc. Natl. Acad. Sci. U S A 2014, 111, E1571-1580. (87) Serdiuk, T.; Balasubramaniam, D.; Sugihara, J.; Mari, S. A.; Kaback, H. R.; Muller, D. J., YidC Assists the Stepwise and Stochastic Folding of Membrane Proteins. Nat. Chem. Biol. 2016, 12, 911-917. (88) Kedrov, A.; Janovjak, H.; Sapra, K. T.; Muller, D. J., Deciphering Molecular Interactions of Native Membrane Proteins by Single-Molecule Force Spectroscopy. Annu. Rev. Biophys. Biomo.l Struct. 2007, 36, 233-260. (89) Sapra, K. T.; Besir, H.; Oesterhelt, D.; Muller, D. J., Characterizing Molecular Interactions in Different Bacteriorhodopsin Assemblies by Single-Molecule Force Spectroscopy. J. Mol. Biol. 2006, 355, 640-650. (90) Serdiuk, T.; Sugihara, J.; Mari, S. A.; Kaback, H. R.; Muller, D. J., Observing a LipidDependent Alteration in Single Lactose Permeases. Structure 2015, 23, 754-761. (91) Gordeliy, V. I.; Schlesinger, R.; Efremov, R.; Buldt, G.; Heberle, J., Crystallization in Lipidic Cubic Phases: A Case Study with Bacteriorhodopsin. Meth. Mol. Biol. 2003, 228, 305316. (92) Petrosyan, R.; Bippes, C. A.; Walheim, S.; Harder, D.; Fotiadis, D.; Schimmel, T.; Alsteens, D.; Muller, D. J., Single-Molecule Force Spectroscopy of Membrane Proteins from Membranes Freely Spanning across Nanoscopic Pores. Nano Lett. 2015, 15, 3624-3633.


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FIGURE LEGENDS

Figure 1. Combined confocal fluorescence microscopy and FD-based AFM imaging. (a) Experimental setup. A thin layer of mica (< 0.05 mm) glued on a glass coverslip of the inverted confocal microscope is used as support for native purple membrane. While raster scanning the sample the AFM tip at the free end of the cantilever and the sample are approached and retracted pixel-by-pixel. For each approach and retraction cycle a forcedistance (FD) curve is recorded. The maximal imaging force pressing the tip to the sample is kept at a minimum to prevent sample distortion. (b) After adsorption of His-tagged purple membrane to mica, the sample is incubated with fluorescently labelled NTA-ligands (Ni2+NTA-Atto 488), which had been already complexed with Ni2+. In the presence of the coordinating Ni2+, NTA can bind to the His10-tag engineered to the N-terminal end of bacteriorhodopsin (structure adapted from PDB ID: 1FBB)99 exposed at the extracellular purple membrane surface. (c) Fluorescent images indicate purple membrane exposing the extracellular surface to the buffer solution. (d) AFM topograph of the sample imaged by confocal microscopy showing purple membrane adsorbed to mica. Purple membranes exposing the cytoplasmic surface (not fluorescent) protruded 7.2 ± 0.4 nm (mean ± sd; n = 53) from the mica and membranes exposing the extracellular surface (fluorescent) protruded 7.5 ± 0.2 nm (n = 49). (e) Merged fluorescence image and topograph. The fluorescent signal localizes purple membranes exposing the extracellular surface, while unlabelled purple membranes exposing the cytoplasmic surface are only imaged by AFM. The sample was adsorbed and labelled in adsorption buffer (300 mM KCl, 20 mM HEPES, pH 7.4) and imaged in imaging buffer (150 mM KCl, 20 mM HEPES, pH 7.4) at room temperature as described (Methods).

Figure 2. High-resolution AFM imaging of purple membrane with C-terminally His5-tagged bacteriorhodopsins. (a) Overview topograph showing purple membranes adsorbed to mica. (b) Overlay topograph and confocal image recorded of fluorescently labelled (Ni2+-NTA-Atto 488, green) purple membrane exposing His5-tags at their cytoplasmic surface. Correlation of both microscopy images identifies purple membrane exposing either the cytoplasmic


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(green) or the extracellular (golden/brown) surface. (c) High-resolution AFM topograph of the extracellular surface or purple membrane showing characteristic structural features of bacteriorhodopsin trimers. (d) Correlation averaged topograph calculated from ≈ 150 bacteriorhodopsin trimers. (e) High-resolution topograph of the cytoplasmic surface showing characteristic structural features of bacteriorhodopsin trimers exposing the extracellular surface. (f) Correlation averaged topograph from ≈ 150 bacteriorhodopsin trimers exposing the cytoplasmic surface. The sample was adsorbed and labelled using Ni2+NTA-Atto 488 in adsorption buffer (300 mM KCl, 20 mM HEPES, pH 7.4) and imaged in imaging buffer (150 mM KCl, 20 mM HEPES, pH 7.4) at room temperature as described (Methods).

Figure 3. Mapping specific ligand-receptor binding events between a NTA-functionalized AFM tip and His-tagged bacteriorhodopsin of purple membrane. (a and b) Confocal image and AFM topograph of purple membrane patches adsorbed to mica and incubated with fluorescently labelled Ni2+-NTA-ligands (Ni2+-NTA-Atto 488, green) in adsorption buffer (300 mM KCl, 20 mM HEPES, pH 7.4). Because the His5-tagged C-termini of bacteriorhodopsin localize at the cytoplasmic purple membrane surface, NTA-Atto 488 labels the purple membranes exposing the cytoplasmic surface. The AFM topograph shows two purple membrane patches exposing either their cytoplasmic or extracellular surface. (c) Merging confocal image and AFM topograph co-localizes the fluorescent signal with only one purple membrane exposing the cytoplasmic surface. The other purple membrane exposes the extracellular surface. (d) The AFM tip was chemically functionalized with a heterobifunctionalized 18-unit long PEG-linker to which free end tris-NTA was attached. In the presence of Ni2+, NTA can bind the His5-tagged C-terminus of bacteriorhodopsin. (e) Purple membranes were washed with imaging buffer supplemented with 500 mM imidazole (150 mM KCl, 500 mM imidazole, 20 mM HEPES, pH 7.4) to remove the Ni2+-NTA-Atto 488 and the topograph was recorded with the tris-NTA functionalized tip in affinity buffer (120 mM KCl, 5 mM NiCl2, 20 mM HEPES, pH 7.4). (f) Adhesion map of the purple membranes recorded in the presence of 5 mM Ni2+ to coordinate the formation of His5-tag-NTA bonds with the functionalized AFM tip. Adhesion events occur mostly on purple membrane exposing the cytoplasmic surface. (g) The same AFM tip detects significantly fewer adhesion


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events on purple membrane in imaging buffer supplemented with 10 mM EDTA (150 mM KCl, 10 mM EDTA, 20 mM HEPES, pH 7.4) to chelate the coordinating Ni2+ ions. Pixels indicating adhesion have been enlarged by a factor two to improve visualization.

Figure 4. Mechanical unfolding of single N-terminally His10-tagged bacteriorhodopsin from purple membrane. (a) Single FD curves recorded upon mechanically unfolding and extracting single bacteriorhodopsins from native purple membrane. For unfolding bacteriorhodopsin was mechanically pulled from the C-terminal end. (b) Density plot of 183 superimposed FD curves depicting common unfolding force peaks. Colored lines are wormlike chain (WLC) curves indicating the contour length of the unfolded polypeptide in each unfolding step.78,80 Contour lengths in amino acids (aa) are given at the top of each WLC curve. Dashed lines indicate minor force peaks (light grey) and solid lines indicate major force peaks (black). The scale at the right indicates the density of data points of superimposed FD curves. (c) Schematic representation of the unfolding intermediates described by the major force peaks recorded during the mechanical unfolding of bacteriorhodopsin. Secondary structure elements are colored similar to the WLC curves fitting the of major force peaks describing their unfolding. Contour lengths taken from the WLC curves and used to assign the unfolding structural segments of bacteriorhodopsin are indicated. Purple membrane was adsorbed and labelled with Ni2+-NTA-Atto 488 in adsorption buffer and the sidedness of purple membrane was analyzed with fluorescence microscopy. SMFS was performed in unfolding buffer as described (Methods).


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TOC graphics for manuscript 39x24mm (300 x 300 DPI)


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Figure 1. 113x80mm (300 x 300 DPI)


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Figure 2. 57x18mm (300 x 300 DPI)


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Figure 3. 148x132mm (300 x 300 DPI)


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Figure 4. 136x240mm (300 x 300 DPI)


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