Simon Scheuring

Imaging and manipulation of biological structures with the AFM

Many biologists have dreamt of physically touching and manipulating the biomolecules they were investigating. With the invention of the atomic force microscope (AFM), this dream has come true. Here, recent applications of the AFM to image and to manipulate biological systems at the nanometer scale are reviewed. Macromolecular biological assemblies as well as individual biomolecules can be subjected to controlled nanomanipulation. Examples of AFM application in imaging and nanomanipulation include the extraction of chromosomal DNA for genetic analysis, the disruption of antibody--antigen bonds, the dissection of biological membranes, the nanodissection of protein complexes, and the controlled modulation of protein conformations. Also reviewed is the novel combination of single molecule imaging and force spectroscopy which allows biomolecules to be imaged, and inter- and intramolecular forces to be measured. Future application of these nanotechniques will reveal new information on the structure, function and assembly of biomolecules.

Electrostatically Balanced Subnanometer Imaging of Biological Specimens by Atomic Force Microscope

To achieve high-resolution topographs of native biological macromolecules in aqueous solution with the atomic force microscope (AFM) interactions between AFM tip and sample need to be considered. Short-range forces produce the submolecular information of high-resolution topographs. In contrast, no significant high-resolution information is provided by the long-range electrostatic double-layer force. However, this force can be adjusted by pH and electrolytes to distribute the force applied to the AFM tip over a large sample area. As demonstrated on fragile biological samples, adjustment of the electrolyte solution results in a local reduction of both vertical and lateral forces between the AFM tip and proteinous substructures. Under such electrostatically balanced conditions, the deformation of the native protein is minimized and the sample surface can be reproducibly contoured at a lateral resolution of 0.6 nm.

Nanodissection and high-resolution imaging of the Rhodopseudomonas viridis photosynthetic core complex in native membranes by AFM

In photosynthesis, highly organized multiprotein assemblies convert sunlight into biochemical energy with high efficiency. A challenge in structural biology is to analyze such supramolecular complexes in native membranes. Atomic force microscopy (AFM) with high lateral resolution, high signal-to-noise ratio, and the possibility to nanodissect biological samples is a unique tool to investigate multiprotein complexes at molecular resolution in situ. Here we present high-resolution AFM of the photosynthetic core complex in native Rhodopseudomonas viridis membranes. Topographs at 10-Å lateral and ≈1-Å vertical resolution reveal a single reaction center (RC) surrounded by a closed ellipsoid of 16 light-harvesting (LH1) subunits. Nanodissection of the tetraheme cytochrome (4Hcyt) subunit from the RC allows demonstration that the L and M subunits exhibit an asymmetric topography intimately associated to the LH1 subunits located at the short ellipsis axis. This architecture implies a distance distribution between the antenna and the RC compared with a centered location of the RC within a circular LH1, which may influence the energy transfer within the core complex. The LH1 subunits rearrange into a circle after removal of the RC from the core complex.

High resolution AFM topographs of the Escherichia coli water channel aquaporin Z.

Aquaporins form a large family of membrane channels involved in osmoregulation. Electron crystallography has shown monomers to consist of six membrane spanning α‐helices confirming sequence based predictions. Surface exposed loops are the least conserved regions, allowing differentiation of aquaporins. Atomic force microscopy was used to image the surface of aquaporin Z, the water channel of Escherichia coli. Recombinant protein with an N‐terminal fragment including 10 histidines was isolated as a tetramer by Ni‐affinity chromatography, and reconstituted into two‐dimensional crystals with p4212 symmetry. Small crystalline areas with p4 symmetry were found as well. Imaging both crystal types before and after cleavage of the N‐termini allowed the cytoplasmic surface to be identified; a drastic change of the cytoplasmic surface accompanied proteolytic cleavage, while the extracellular surface morphology did not change. Flexibility mapping and volume calculations identified the longest loop at the extracellular surface. This loop exhibited a reversible force‐induced conformational change.

Cellular capsules as a tool for multicellular spheroid production and for investigating the mechanics of tumor progression in vitro

Significance Tumor growth intrinsically generates pressure onto the surrounding tissues, which conversely compress the tumor. These mechanical forces have been suggested to contribute to tumor growth regulation. We developed a microfluidic technique to produce 3D cell-based assays and to interrogate the interplay between tumor growth and mechanics in vitro. Multicellular spheroids are grown in permeable elastic capsules. Capsule deformation provides a direct measure of the exerted pressure. By simultaneously imaging the spheroid by confocal microscopy, we show that confinement induces a drastic cellular reorganization, including increased motility of peripheral cells. We propose that compressive stress has a beneficial impact on slowing down tumor evolution but may have a detrimental effect by triggering cell invasion and metastasis. Deciphering the multifactorial determinants of tumor progression requires standardized high-throughput preparation of 3D in vitro cellular assays. We present a simple microfluidic method based on the encapsulation and growth of cells inside permeable, elastic, hollow microspheres. We show that this approach enables mass production of size-controlled multicellular spheroids. Due to their geometry and elasticity, these microcapsules can uniquely serve as quantitative mechanical sensors to measure the pressure exerted by the expanding spheroid. By monitoring the growth of individual encapsulated spheroids after confluence, we dissect the dynamics of pressure buildup toward a steady-state value, consistent with the concept of homeostatic pressure. In turn, these confining conditions are observed to increase the cellular density and affect the cellular organization of the spheroid. Postconfluent spheroids exhibit a necrotic core cemented by a blend of extracellular material and surrounded by a rim of proliferating hypermotile cells. By performing invasion assays in a collagen matrix, we report that peripheral cells readily escape preconfined spheroids and cell–cell cohesivity is maintained for freely growing spheroids, suggesting that mechanical cues from the surrounding microenvironment may trigger cell invasion from a growing tumor. Overall, our technology offers a unique avenue to produce in vitro cell-based assays useful for developing new anticancer therapies and to investigate the interplay between mechanics and growth in tumor evolution.

Variable LH2 stoichiometry and core clustering in native membranes of Rhodospirillum photometricum.

The individual components of the photosynthetic unit (PSU), the light‐harvesting complexes (LH2 and LH1) and the reaction center (RC), are structurally and functionally known in great detail. An important current challenge is the study of their assembly within native membranes. Here, we present AFM topographs at 12 Å resolution of native membranes containing all constituents of the PSU from Rhodospirillum photometricum. Besides the major technical advance represented by the acquisition of such highly resolved data of a complex membrane, the images give new insights into the organization of this energy generating apparatus in Rsp. photometricum: (i) there is a variable stoichiometry of LH2, (ii) the RC is completely encircled by a closed LH1 assembly, (iii) the LH1 assembly around the RC forms an ellipse, (iv) the PSU proteins cluster together segregating out of protein free lipid bilayers, (v) core complexes cluster although enough LH2 are present to prevent core–core contacts, and (vi) there is no cytochrome bc1 complex visible in close proximity to the RCs. The functional significance of all these findings is discussed.

Characterization of the motion of membrane proteins using high-speed atomic force microscopy

For cells to function properly, membrane proteins must be able to diffuse within biological membranes. The functions of these membrane proteins depend on their position and also on protein-protein and protein-lipid interactions. However, so far, it has not been possible to study simultaneously the structure and dynamics of biological membranes. Here, we show that the motion of unlabelled membrane proteins can be characterized using high-speed atomic force microscopy. We find that the molecules of outer membrane protein F (OmpF) are widely distributed in the membrane as a result of diffusion-limited aggregation, and while the overall protein motion scales roughly with the local density of proteins in the membrane, individual protein molecules can also diffuse freely or become trapped by protein-protein interactions. Using these measurements, and the results of molecular dynamics simulations, we determine an interaction potential map and an interaction pathway for a membrane protein, which should provide new insights into the connection between the structures of individual proteins and the structures and dynamics of supramolecular membranes.

Filming Biomolecular Processes by High-Speed Atomic Force Microscopy

In the history of science and technology, researchers have always undertaken endeavors to enhance the degree of “directness of measurement”. With “directness of measurement”, we conceptualize that the essence of the object under investigation, the structure, dynamics, and function of biological samples in the scope of this Review, is entirely and straightforwardly assessed by the measurement, bypassing hypotheses and intricate data analysis. When the degree of measurement directness is low, conclusions derived from the gleaned data often differ depending on the formulation of hypotheses, analysis models, and the data interpretation. That is why conceptual consensus about a specific issue of the studied object is rarely reached based on indirect data. This is often the case for studies of structure–function relationship of proteins. Previously, scientists had to understand distinct qualities of proteins by measuring a sample containing a huge number of molecules. The results are ensemble-averaged quantities (often at equilibrium) that provide only limited information on the proteins of study. Proteins are dynamic in nature and work at the single-molecule level. Protein molecules fluctuate, undergo structural changes, bind to and dissociate from interaction partners, and traverse a range of energy and chemical states during molecular action. Most, if not all, of these dynamics and their statistical distributions are convoluted and hidden in ensemble averaging measurements. To overcome the limitations of ensemble measurements, single-molecule biophysics was developed more than two decades ago, with the use of fluorescence microscopy,1,2 optical spectroscopy,3,4 and optical and magnetic tweezers,5,6 whose performances were further improved by the advancements of improved optical microscopes, lasers, electronics, computers, and high-sensitivity video cameras and sensors. Using these techniques, our understanding of the functional mechanism of proteins has made significant steps forward. Moreover, super-resolution optical microscopy techniques bypassing the diffraction limit for fluorophore localization have recently been added to fluorescence microscopy.7−9 However, the degree of directness of measurement is still limited, because the protein molecules themselves are invisible in these single-molecule measurements. Protein structure is typically studied by X-ray crystallography, electron microscopy (EM), and nuclear magnetic resonance (NMR) spectroscopy. To date, these techniques have revealed detailed three-dimensional structures of over 94 000 proteins (Protein Data Bank (PDB), http://www.rcsb.org/pdb/home/home.do), with a growth rate of about 8000 novel structures per year (2010–2013). Yet, these techniques make use of ensemble averaging, and, more seriously, the obtained structures are merely limited to static snapshots of fixed conformations. Thus, the simultaneous and direct observation of structure, dynamics, and function of single protein molecules has long been infeasible, and hence the materialization of a technique allowing such an observation has long been awaited in biological sciences. An ideal microscopy technique that allows simultaneous observation of structure, dynamics, and function of single protein molecules has to meet all of the following conditions (see Table 1): (i) in-liquid specimen imaging, (ii) high spatial resolution, (iii) high temporal resolution, (iv) low invasiveness to the specimen, and (v) direct imaging of the specimen without the use of markers (in other words, resolving the structure of the specimen itself). Although efforts have been made to develop environmental electron microscopy techniques capable of observing unstained biological specimens in solutions,10 the strong electron dose that is required to achieve high contrast and spatial resolution instantaneously denatures the sample. Achieving the above-described goals by EM is a highly difficult, if not impossible, task. Conventional atomic force microscopy11 (AFM) meets most of the above-mentioned conditions, except for the third condition, that is, high temporal resolution, and the fourth condition, that is, low invasiveness, is only moderately satisfied. Table 1 Feasibility Comparison of Three Types of Microscopy

High-speed force spectroscopy unfolds titin at the velocity of molecular dynamics simulations.

Bridging the Titin Gap The muscle protein titin is a molecular spring that has been extensively studied by single-molecule unfolding experiments and by molecular simulation. However, experimental and simulated unfolding could not be compared directly because they differ by orders of magnitude in pulling velocity. Rico et al. (p 741) developed high-speed force spectroscopy to pull titin molecules at speeds that reach the lower limits of molecular dynamics simulations. Bridging the gap between simulation and experiment clarified the mechanism of conformational changes in titin. Experimental time scales previously accessible only to simulations provide insight into forced protein unfolding. The mechanical unfolding of the muscle protein titin by atomic force microscopy was a landmark in our understanding of single-biomolecule mechanics. Molecular dynamics simulations offered atomic-level descriptions of the forced unfolding. However, experiment and simulation could not be directly compared because they differed in pulling velocity by orders of magnitude. We have developed high-speed force spectroscopy to unfold titin at velocities reached by simulation (~4 millimeters per second). We found that a small β-strand pair of an immunoglobulin domain dynamically unfolds and refolds, buffering pulling forces up to ~100 piconewtons. The distance to the unfolding transition barrier is larger than previously estimated but is in better agreement with atomistic predictions. The ability to directly compare experiment and simulation is likely to be important in studies of biomechanical processes.

Structure of the Dimeric PufX-containing Core Complex of Rhodobacter blasticus by in Situ Atomic Force Microscopy

We have studied photosynthetic membranes of wild type Rhodobacter blasticus, a closely related strain to the well studied Rhodobacter sphaeroides, using atomic force microscopy. High-resolution atomic force microscopy topographs of both cytoplasmic and periplasmic surfaces of LH2 and RC-LH1-PufX (RC, reaction center) complexes were acquired in situ. The LH2 is a nonameric ring inserted into the membrane with the 9-fold axis perpendicular to the plane. The core complex is an S-shaped dimer composed of two RCs, each encircled by 13 LH1 α/β-heterodimers, and two PufXs. The LH1 assembly is an open ellipse with a topography-free gap of ∼25 Å. The two PufXs, one of each core, are located at the dimer center. Based on our data, we propose a model of the core complex, which provides explanation for the PufX-induced dimerization of the Rhodobacter core complex. The QB site is located facing a ∼25-Å wide gap within LH1, explaining the PufX-favored quinone passage in and out of the core complex.