Imaging modes of atomic force microscopy for application in molecular and cell biology

Journal name:
Nature Nanotechnology
Volume:
12,
Pages:
295–307
Year published:
DOI:
doi:10.1038/nnano.2017.45
Received
Accepted
Published online

Abstract

Atomic force microscopy (AFM) is a powerful, multifunctional imaging platform that allows biological samples, from single molecules to living cells, to be visualized and manipulated. Soon after the instrument was invented, it was recognized that in order to maximize the opportunities of AFM imaging in biology, various technological developments would be required to address certain limitations of the method. This has led to the creation of a range of new imaging modes, which continue to push the capabilities of the technique today. Here, we review the basic principles, advantages and limitations of the most common AFM bioimaging modes, including the popular contact and dynamic modes, as well as recently developed modes such as multiparametric, molecular recognition, multifrequency and high-speed imaging. For each of these modes, we discuss recent experiments that highlight their unique capabilities.

At a glance

Figures

  1. Timeline of key inventions, starting from the birth of AFM in 1986 to the latest AFM imaging modes in molecular and cell biology.
    Figure 1: Timeline of key inventions, starting from the birth of AFM in 1986 to the latest AFM imaging modes in molecular and cell biology.

    Key inventions developed over the years include: an optical detection system and fluid cell enabling contact mode AFM to operate in aqueous solution (Bio-AFM); dynamic mode AFM (DM-AFM), which oscillates the AFM tip to reduce friction while contouring the biological sample; force–distance curve-based AFM (FD-AFM), which contours the surface of a biological system while recording pixel-by-pixel a full force–distance curve; multiparametric AFM (MP-AFM), which contours the sample while mapping multiple physical or chemical properties; molecular recognition AFM (MR-AFM), which images and maps specific interactions of biological samples; multifrequency AFM (MF-AFM), which contours the sample while oscillating the cantilever tip at multiple frequencies, thus mapping various physical parameters; correlating advanced optical imaging and AFM (Opto-AFM) for the imaging of complex biological systems; high-speed AFM (HS-AFM), which speeds up the image acquisition time by a factor of ~1,000, providing access to dynamic processes in biology. Most modes cross-fertilized each other, ultimately leading to combinatorial AFM. Images adapted from: Bio-AFM, ref. 28, Macmillan Publishers Ltd; DM-AFM, ref. 45, American Chemical Society; FD-AFM, ref. 76, Wiley; MP-AFM, ref. 78, Elsevier; MR-AFM, ref. 9, Cell Press; MF-AFM, ref. 46, Macmillan Publishers Ltd; Opto-AFM, ref. 145, The Company of Biologists; HS-AFM, ref. 122, Macmillan Publishers Ltd.

  2. AFM-based imaging of native biological systems to molecular resolution.
    Figure 2: AFM-based imaging of native biological systems to molecular resolution.

    a, Basic principles of contact (left) and dynamic (right) AFM imaging modes. In contact mode, the cantilever deflection is kept constant (constant force) by adjusting the relative height between tip and sample. A topographic height change alters the cantilever deflection, which a feedback loop corrects by adjusting the tip–sample distance. The dynamic mode oscillates the cantilever close to or at resonance frequency. Height changes alter the cantilever oscillation, which is used to adjust the tip–sample distance. be, Contact mode AFM topographs. b, Cyclic nucleotide-regulated potassium channels (MlotiK1) reconstituted into lipid membranes. c,d, Rows of densely packed rhodopsin dimers distributed in the native disc membrane extracted from rod outer segments of the eye. e, Image of a living SAOS-A2 cell bundling and pulling collagen fibrils coating a substrate. To maximize contrast, the exemplified image shows the deflection of the cantilever, which changes while contouring the sample. f–h, Dynamic mode AFM topographs. f, An IgG antibody absorbed to mica and visualized with frequency modulation mode. g, Single brome mosaic viruses packed in a crystalline assembly. h, Circular plasmid DNA imaged in buffer solution by frequency modulation AFM. Red and blue arrows indicate major and minor grooves of the DNA, respectively. Panels adapted from: b, ref. 148, PNAS; c,d, ref. 30, Macmillan Publishers Ltd; e, ref. 147, Elsevier; f, ref. 46, Macmillan Publishers Ltd; g, ref. 146, Elsevier; h, ref. 45, American Chemical Society.

  3. Force-distance curve-based AFM.
    Figure 3: Force–distance curve-based AFM.

    a, Principle of recording force–distance (FD) curves by approaching (blue) and withdrawing (red) the AFM tip from the sample. The tip of the cantilever is initially away from the sample (1) to which it is brought into contact (2). During retraction (3) of the AFM tip, adhesive events may occur at different distances due to nonspecific (4) or specific (5) interactions between tip and sample. b, FD-based AFM imaging records pixel-by-pixel FD curves while contouring the sample topography. The indentation force Fi is controlled and parameters extracted include the tip–sample adhesion force Fadh, or elastic and electrostatic properties (by fitting the curve). Parameters can be displayed as coloured maps and correlated to the topography. c, Example of multiparametric FD-based AFM imaging of the elasticity and adhesion of two dividing Staphylococcus aureus cells. d, AFM force error (top) and elasticity (bottom) maps of living HaCaT keratinocytes. e, Topography (left, brown coloured) and stiffness map (top right) of nuclear pore complexes from the cytoplasmic surface. The graph (bottom right) shows the stiffness as a function of tip–sample separation recorded close to the centre of the cytoplasmic ring. Grey dots represent data points and the red curve is the average. Blue and black dashed lines are fits to the data using indentation models for spherical and conical tips, respectively. f, Top left: topograph of human protease activated receptors 1 (PAR1) in proteoliposomes recorded with a SFLLRN-ligand functionalized tip. Bottom left: overlay of topograph (grey) and adhesive interactions (red) localizes individual receptors binding the ligand. The circles numbered 1–4 indicate regions at which FD curves 1–4 were taken. Top right: FD curves exemplifying unspecific adhesion events (1 and 2) and specific ligand-receptor unbinding events (3 and 4) showing the stretching of the linker tethering the ligand to the AFM tip. Bottom right: free energy landscape of the ligand binding to PAR1 extracted from measuring the rupture force of the ligand-receptor bond at different loading rates. xu describes the distance of the bound to the transition state and ΔGbu is the binding free energy. Panels adapted from: c, ref. 79, PNAS; f, ref. 99, Macmillan Publishers Ltd. Panels reproduced from: d, ref. 78, Elsevier; e, ref. 149, Macmillan Publishers Ltd.

  4. Multifrequency AFM.
    Figure 4: Multifrequency AFM.

    a, Scheme of the deflection of the cantilever in bimodal AFM. b, Two eigenmodes of the cantilever are excited and detected. Observables associated with both eigenmodes are recorded to determine sample properties such as flexibility, deformation and viscosity. c, Separation of short-range mechanical forces and long-range magnetic interactions in ferritin. The first eigenmode contours the topography while the second eigenmode detects long-range magnetic forces. By combining both signals, the iron oxide core and the apoferritin shell are separated in the AFM image. d, Top: AFM topograph (xy frame) of GroEL proteins. Bottom: vertical profile (xz frame, taken along the red dashed line of the topography) of the hydration layers contouring four GroEL molecules. The dashed red line marks the surface of the GroEL molecules. e, Multifrequency flexural AFM of a bacteriophage Φ29 mature virion. The virion topography (left) is acquired simultaneously with multiharmonic observables images, from which the viscosity map (right) is shown. Images were recorded applying 100 pN. f, Electron microscopy image of a T-shaped cantilever designed for torsional harmonic AFM. g, Multifrequency torsional harmonics scheme for probing chemical groups of a protein using DNA labels. A DNA strand attached to the tip interacts with target DNA strands. Complementary sequences have identical colours. Panels adapted from: a, ref. 42, Macmillan Publishers Ltd; c, ref. 107, IOP; d, ref. 108, RSC; e, ref. 110, RSC; f, ref. 60, Macmillan Publishers Ltd. Panel g reproduced from ref. 112, Macmillan Publishers Ltd.

  5. HS-AFM filming proteins in action.
    Figure 5: HS-AFM filming proteins in action.

    a, Illustration of filming myosin walking along actin filaments. be, Key devices and techniques for HS-AFM. b, Small cantilever with high resonant frequency and small spring constant. c, Fast scanner suppressing impulses generated by quick displacements of piezoelectric X- and Z-scanners. d, Active vibration damping based on Q-control with mock Z-scanner. e, Feedback controller with automatic gain tuning for low-invasive high-speed imaging without causing tip-parachuting. f–i, HS-AFM images of proteins in action. f, Bacteriorhodopsin in native purple membrane recorded under dark and illumination at 1 frame per second. White triangles indicate bacteriorhodopsin trimers. Blue triangles indicate 'trefoils' that comprise three bacteriorhodopsin monomers, each belonging to an adjacent trimer. Green light was illuminated at 2 s and switched off at 3 s. On illumination, bacteriorhodopsin trimers dilate outwardly, while bacteriorhodopsin monomers contact each other in trefoils. g, Myosin V walking unidirectionally along an actin filament, showing forward rotation of the leading lever-arm on trailing head detachment from actin. Dashed white lines indicate positions of actin-bound myosin heads. h, Rotorless F1-ATPase undergoing conformational changes. Red circles indicate the highest positions of the topographs. Since a nucleotide-free β-subunit protrudes higher than ADP- and ATP-bound ones, it is observed that the unbound state rotates anticlockwise. i, Spiral filament formation by polymerization of the ESCRT-III protein Snf7 on a supported lipid membrane. Panels adapted from: f, ref. 122, Macmillan Publishers Ltd; g, ref. 123, Macmillan Publishers Ltd; h, ref. 124, AAAS; i, ref. 125, Elsevier.

  6. AFM of cellular systems.
    Figure 6: AFM of cellular systems.

    a, Schematic setup of an AFM combined with optical microscopy for the characterization of living cells. b,c, Fluorescence image (b) and correlative AFM images (c) of a macrophage (green) incubated for 3 h with cells from Candida albicans (blue). Images in panel c are enlarged views of the dashed areas shown in the fluorescence image. Internalized (bottom) and externalized (top) hyphae featuring major structural differences. d, HS-AFM topographs of the Escherichia coli bacterium. The first topograph shows the entire bacterium, while the following images of the outer membrane show moving net-like structures formed by porin trimers. The inset in the last image shows a single porin trimer. e, Mechanical confinement and morphological characterization of mitotic animal cells. The image shows overlaid differential interference contrast and histoneH2B green fluorescent protein (H2B-GFP) images of mitotic HeLa cells. The cantilever (dark shadowed) confines a single mitotic cell in the metaphase to measure the force and pressure generated by the rounding animal cell. f, Wedged cantilever applied to confine a mitotic HeLa cell and to mechanically control mitotic progression. Depicted are mitotic phases, spindle microtubules (green), chromosomes (red), nuclear envelope breakdown (NEBD), time (t), and force (Fset) and height (h) set by the cantilever. g,h, Top view of spindle characterization scheme of a confined HeLa cell (g) and fluorescence snapshots of microtubules (mTubulin–GFP) and chromosomes (H2B–mCherry) (h). Grey double arrows, metaphase plate width; white arrows, stray chromosomes. t = 0, NEBD. Panels adapted from: a, ref. 131, American Chemical Society; e, ref. 139, Macmillan Publishers Ltd. Panels reproduced from: b,c, ref. 131, American Chemical Society; d, ref. 128, Elsevier; f–h, ref. 58, PNAS.

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Affiliations

  1. Institute of Life Sciences and Walloon Excellence in Life Sciences and Biotechnology (WELBIO), Université catholique de Louvain, Croix du Sud 4-5, bte L7.07.06., B-1348 Louvain-la-Neuve, Belgium

    • Yves F. Dufrêne &
    • David Alsteens
  2. Department of Physics, Kanazawa University, Kanazawa 920-1192, Japan

    • Toshio Ando
  3. Instituto de Ciencia de Materiales de Madrid, CSIC, Sor Juana Inés de la Cruz 3, 28049 Madrid, Spain

    • Ricardo Garcia
  4. Department of Biosystems Science and Engineering, Eidgenössische Technische Hochschule (ETH) Zürich, Mattenstrasse 28, 4056 Basel, Switzerland

    • David Martinez-Martin &
    • Daniel J. Müller
  5. Department of BioNanoscience, Delft University of Technology, Van der Maasweg 9, 2629 HZ Delft, The Netherlands

    • Andreas Engel
  6. Swiss Nanoscience Institute, University of Basel, Klingelbergstrasse 80, 4057 Basel, Switzerland

    • Christoph Gerber

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