Single-molecule force spectroscopy of membrane proteins from membranes freely spanning across nanoscopic pores

Photosynthetic Protein-Based Retinal Ganglion Cell Receptive Fields for Detecting Edges and Brightness Illusions

Bacteriorhodopsin, isolated from a halophilic bacterium, is a photosynthetic protein with a structure and function similar to those of the visual pigment rhodopsin. A voltaic cell with bacteriorhodopsin sandwiched between two transparent electrodes exhibits a time-differential response akin to that observed in retinal ganglion cells. It is intriguing as a means to emulate excitation and inhibition in the neural response. Here, we present a neuromorphic device emulating the retinal ganglion cell receptive field fabricated by patterning bacteriorhodopsin onto two transparent electrodes and encapsulating them with an electrolyte solution. This protein-based artificial ganglion cell receptive field is characterized as a bandpass filter that simultaneously replicates excitatory and inhibitory responses within a single element, successfully detecting image edges and phenomena of brightness illusions. The device naturally emulates the highly interacting ganglion cell receptive fields by exploiting the inherent properties of proteins without the need for electronic components, bias power supply, or an external operating circuit.

Single-Molecule Force Spectroscopy of Membrane Protein Folding

Single-molecule force spectroscopy is a unique method that can probe the structural changes of single proteins at a high spatiotemporal resolution while mechanically manipulating them over a wide force range. Here, we review the current understanding of membrane protein folding learned by using the force spectroscopy approach. Membrane protein folding in lipid bilayers is one of the most complex biological processes in which diverse lipid molecules and chaperone proteins are intricately involved. The approach of single protein forced unfolding in lipid bilayers has produced important findings and insights into membrane protein folding. This review provides an overview of the forced unfolding approach, including recent achievements and technical advances. Progress in the methods can reveal more interesting cases of membrane protein folding and clarify general mechanisms and principles.

Anisotropic Friction in a Ligand-Protein Complex

The effect of an externally applied directional force on molecular friction is so far poorly understood. Here, we study the force-driven dissociation of the ligand-protein complex biotin-streptavidin and identify anisotropic friction as a not yet described type of molecular friction. Using AFM-based stereographic single molecule force spectroscopy and targeted molecular dynamics simulations, we find that the rupture force and friction for biotin-streptavidin vary with the pulling angle. This observation holds true for friction extracted from Kramers' rate expression and by dissipation-corrected targeted molecular dynamics simulations based on Jarzynski's identity. We rule out ligand solvation and protein-internal friction as sources of the angle-dependent friction. Instead, we observe a heterogeneity in free energy barriers along an experimentally uncontrolled orientation parameter, which increases the rupture force variance and therefore the overall friction. We anticipate that anisotropic friction needs to be accounted for in a complete understanding of friction in biomolecular dynamics and anisotropic mechanical environments.

Histidine-Specific Bioconjugation for Single-Molecule Force Spectroscopy

Atomic force microscopy (AFM) based single-molecule force spectroscopy (SMFS) is a powerful tool to study the mechanical properties of proteins. In these experiments, site-specific immobilization of proteins is critical, as the tether determines the direction and amplitude of forces applied to the protein of interest. However, existing methods are mainly based on thiol chemistry or specific protein tags, which cannot meet the need of many challenging experiments. Here, we developed a histidine-specific phosphorylation strategy to covalently anchor proteins to an AFM cantilever tip or the substrate via their histidine tag or surface-exposed histidine residues. The formed covalent linkage was mechanically stable with rupture forces of over 1.3 nN. This protein immobilization method considerably improved the pickup rate and data quality of SMFS experiments. We further demonstrated the use of this method to explore the pulling-direction-dependent mechanical stability of green fluorescent protein and the unfolding of the membrane protein archaerhodopsin-3.

Single-Molecule Force Spectroscopy of Protein Folding

The use of force probes to induce unfolding and refolding of single molecules through the application of mechanical tension, known as single-molecule force spectroscopy (SMFS), has proven to be a powerful tool for studying the dynamics of protein folding. Here we provide an overview of what has been learned about protein folding using SMFS, from small, single-domain proteins to large, multi-domain proteins. We highlight the ability of SMFS to measure the energy landscapes underlying folding, to map complex pathways for native and non-native folding, to probe the mechanisms of chaperones that assist with native folding, to elucidate the effects of the ribosome on co-translational folding, and to monitor the folding of membrane proteins.

Membrane-Protein Unfolding Intermediates Detected with Enhanced Precision Using a Zigzag Force Ramp

Precise quantification of the energetics and interactions that stabilize membrane proteins in a lipid bilayer is a long-sought goal. Toward this end, atomic force microscopy has been used to unfold individual membrane proteins embedded in their native lipid bilayer, typically by retracting the cantilever at a constant velocity. Recently, unfolding intermediates separated by as few as two amino acids were detected using focused-ion-beam-modified ultrashort cantilevers. However, unambiguously discriminating between such closely spaced states remains challenging, in part because any individual unfolding trajectory only occupies a subset of the total number of intermediates. Moreover, structural assignment of these intermediates via worm-like-chain analysis is hindered by brief dwell times compounded with thermal and instrumental noise. To overcome these issues, we moved the cantilever in a sawtooth pattern of 6-12 nm, offset by 0.25-1 nm per cycle, generating a "zigzag" force ramp of alternating positive and negative loading rates. We applied this protocol to the model membrane protein bacteriorhodopsin (bR). In contrast to conventional studies that extract bR's photoactive retinal along with the first transmembrane helix, we unfolded bR in the presence of its retinal. To do so, we introduced a previously developed enzymatic-cleavage site between helices E and F and pulled from the top of the E helix using a site-specific, covalent attachment. The resulting zigzag unfolding trajectories occupied 40% more states per trajectory and occupied those states for longer times than traditional constant-velocity records. In total, we identified 31 intermediates during the unfolding of five helices of EF-cleaved bR. These included a previously reported, mechanically robust intermediate located between helices C and B that, with our enhanced resolution, is now shown to be two distinct states separated by three amino acids. Interestingly, another intermediate directly interacted with the retinal, an interaction confirmed by removing the retinal.

Atomic Force Microscopy of Proteins

Atomic force microscopy (AFM) enables imaging of surface-deposited proteins and protein structures under physiological conditions, which is a benefit compared to ultra-high vacuum techniques such as electron microscopy. AFM also has the potential to provide more information from the phase in tapping mode or from functional AFM modes. The sample preparation, probe selection, and imaging conditions are crucial for successful imaging of proteins. Here we give a detailed account of the steps toward imaging of soft samples in both air and liquid along with the basic theory underpinning these details.

Single-Molecule Force Spectroscopy of Transmembrane β-Barrel Proteins

Single-molecule force spectroscopy (SMFS) has been widely applied to study the mechanical unfolding and folding of transmembrane proteins. Here, we review the recent progress in characterizing bacterial and human transmembrane β-barrel proteins by SMFS. First, we describe the mechanical unfolding of transmembrane β-barrels, which follows a general mechanism dictated by the sequential unfolding and extraction of individual β-strands and β-hairpins from membranes. Upon force relaxation, the unfolded polypeptide can insert stepwise into the membrane as single β-strands or β-hairpins to fold as the native β-barrel. The refolding can be followed at a high spatial and temporal resolution, showing that small β-barrels are able to fold without assistance, whereas large and complex β-barrels require chaperone cofactors. Applied in the dynamic mode, SMFS can quantify the kinetic and mechanical properties of single β-hairpins and reveal complementary insight into the membrane protein structure and function relationship. We further outline the challenges that SMFS experiments must overcome for a comprehensive understanding of the folding and function of transmembrane β-barrel proteins.

High-Resolution AFM-Based Force Spectroscopy

Atomic force microscopy (AFM)-based force spectroscopy is a powerful technique which has seen significant enhancements in both force and time resolution in recent years. This chapter details two AFM cantilever modification procedures that yield high force precision over different temporal bandwidths. Specifically, it explains a fairly straightforward method to achieve sub-pN force precision and stability at low frequencies (<50 Hz) by removing the metal coatings from a commercially available cantilever. A more involved procedure utilizing a focused ion beam milling machine is required to maintain high force precision at enhanced bandwidths. Both modification methods allow site-specific attachment of biomolecules onto the apex area of the tips for force spectroscopy. The chapter concludes with a comparative demonstration using the two cantilever modification methods to study a lipid-protein interaction.

Nanoscale imaging and force probing of biomolecular systems using atomic force microscopy: from single molecules to living cells

Due to the lack of adequate tools for observation, native molecular behaviors at the nanoscale have been poorly understood. The advent of atomic force microscopy (AFM) provides an exciting instrument for investigating physiological processes on individual living cells with molecular resolution, which attracts the attention of worldwide researchers. In the past few decades, AFM has been widely utilized to investigate molecular activities on diverse biological interfaces, and the performances and functions of AFM have also been continuously improved, greatly improving our understanding of the behaviors of single molecules in action and demonstrating the important role of AFM in addressing biological issues with unprecedented spatiotemporal resolution. In this article, we review the related techniques and recent progress about applying AFM to characterize biomolecular systems in situ from single molecules to living cells. The challenges and future directions are also discussed.

Hidden dynamics in the unfolding of individual bacteriorhodopsin proteins

Protein folding occurs as a set of transitions between structural states within an energy landscape. An oversimplified view of the folding process emerges when transiently populated states are undetected because of limited instrumental resolution. Using force spectroscopy optimized for 1-microsecond resolution, we reexamined the unfolding of individual bacteriorhodopsin molecules in native lipid bilayers. The experimental data reveal the unfolding pathway in unprecedented detail. Numerous newly detected intermediates-many separated by as few as two or three amino acids-exhibited complex dynamics, including frequent refolding and state occupancies of <10 μs. Equilibrium measurements between such states enabled the folding free-energy landscape to be deduced. These results sharpen the picture of the mechanical unfolding of membrane proteins and, more broadly, enable experimental access to previously obscured protein dynamics.

High-Resolution Imaging and Multiparametric Characterization of Native Membranes by Combining Confocal Microscopy and an Atomic Force Microscopy-Based Toolbox

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 curve-based (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, whose 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 multimethodological approach is generally applicable to characterize biological membranes and membrane proteins at physiologically relevant conditions.

Detecting Ligand-Binding Events and Free Energy Landscape while Imaging Membrane Receptors at Subnanometer Resolution

Force-distance curve-based atomic force microscopy has emerged into a sophisticated technique for imaging cellular membranes and for detecting specific ligand-binding events of native membrane receptors. However, so far the resolution achieved has been insufficient to structurally map ligand-binding sites onto membrane proteins. Here, we introduce experimental and theoretical approaches for overcoming this limitation. To establish a structurally and functionally well-defined reference sample, we engineer a ligand-binding site to the light-driven proton pump bacteriorhodopsin of purple membrane. Functionalizing the AFM stylus with an appropriate linker-system tethering the ligand and optimizing the AFM conditions allows for imaging the engineered bacteriorhodopsin at subnanometer resolution while structurally mapping the specific ligand-receptor binding events. Improved data analysis allows reconstructing the ligand-binding free energy landscape from the experimental data, thus providing thermodynamic and kinetic insight into the ligand-binding process. The nanoscopic method introduced is generally applicable for imaging receptors in native membranes at subnanometer resolution and for systematically mapping and quantifying the free energy landscape of ligand binding.

Single-Molecule Peptide-Lipid Affinity Assay Reveals Interplay between Solution Structure and Partitioning

Interactions between short protein segments and phospholipid bilayers dictate fundamental aspects of cellular activity and have important applications in biotechnology. Yet, the lack of a suitable methodology for directly probing these interactions has hindered the mechanistic understanding. We developed a precision atomic force microscopy-based single-molecule force spectroscopy assay and probed partitioning into lipid bilayers by measuring the mechanical force experienced by a peptide. Protein segments were constructed from the peripheral membrane protein SecA, a key ATPase in bacterial secretion. We focused on the first 10 amino-terminal residues of SecA (SecA2-11) that are lipophilic. In addition to the core SecA2-11 sequence, constructs with nearly identical chemical composition but with differing geometry were used: two copies of SecA2-11 linked in series and two copies SecA2-11 linked in parallel. Lipid bilayer partitioning interactions of peptides with differing structures were distinguished. To model the energetic landscape, a theory of diffusive barrier crossing was extended to incorporate a superposition of potential barriers with variable weights. Analysis revealed two dissociation pathways for the core SecA2-11 sequence with well-separated intrinsic dissociation rates. Molecular dynamics simulations showed that the three peptides had significant conformational differences in solution that correlated well with the measured variations in the propensity to partition into the bilayer. The methodology is generalizable and can be applied to other peptide and lipid species.

Stability and dynamics of membrane-spanning DNA nanopores

Recently developed DNA-based analogues of membrane proteins have advanced synthetic biology. A fundamental question is how hydrophilic nanostructures reside in the hydrophobic environment of the membrane. Here, we use multiscale molecular dynamics (MD) simulations to explore the structure, stability and dynamics of an archetypical DNA nanotube inserted via a ring of membrane anchors into a phospholipid bilayer. Coarse-grained MD reveals that the lipids reorganize locally to interact closely with the membrane-spanning section of the DNA tube. Steered simulations along the bilayer normal establish the metastable nature of the inserted pore, yielding a force profile with barriers for membrane exit due to the membrane anchors. Atomistic, equilibrium simulations at two salt concentrations confirm the close packing of lipid around of the stably inserted DNA pore and its cation selectivity, while revealing localized structural fluctuations. The wide-ranging and detailed insight informs the design of next-generation DNA pores for synthetic biology or biomedicine.

Although DNA nanopores are widely explored as synthetic membrane proteins, it is still unclear how the anionic DNA assemblies stably reside within the hydrophobic core of a lipid bilayer. Here, the authors use molecular dynamics simulations to reveal the key dynamic interactions and energetics stabilizing the nanopore-membrane interaction.

The physics of pulling polyproteins: a review of single molecule force spectroscopy using the AFM to study protein unfolding

One of the most exciting developments in the field of biological physics in recent years is the ability to manipulate single molecules and probe their properties and function. Since its emergence over two decades ago, single molecule force spectroscopy has become a powerful tool to explore the response of biological molecules, including proteins, DNA, RNA and their complexes, to the application of an applied force. The force versus extension response of molecules can provide valuable insight into its mechanical stability, as well as details of the underlying energy landscape. In this review we will introduce the technique of single molecule force spectroscopy using the atomic force microscope (AFM), with particular focus on its application to study proteins. We will review the models which have been developed and employed to extract information from single molecule force spectroscopy experiments. Finally, we will end with a discussion of future directions in this field.

Unbinding forces and energies between a siRNA molecule and a dendrimer measured by force spectroscopy

We have measured the intermolecular forces between small interference RNA (siRNA) and polyamidoamine dendrimers at the single molecular level. A single molecule force spectroscopy approach has been developed to measure the unbinding forces and energies between a siRNA molecule and polyamidoamine dendrimers deposited on a mica surface in a buffer solution. We report three types of unbinding events which are characterized by forces and free unbinding energies, respectively, of 28 pN, 0.709 eV; 38 pN, 0.722 eV; and 50 pN, 0.724 eV. These events reflect different possible electrostatic interactions between the positive charges of one or two dendrimers and the negatively charged phosphate groups of a single siRNA. We have evidence of a high binding affinity of siRNA towards polyamidoamine dendrimers that leads to a 45% probability of measuring specific unbinding events.