Keywords

Introduction

Viruses are outstanding examples of how the Nature evolution can achieve sophisticated systems designed to last with minimalistic resources [1]. They can be considered as nanomachines whose function and properties are not only interesting for understanding their biology, but also for applications in material science [2]. In this vein, the mechanical properties of biomolecular complexes are essential for their function, and viruses are not an exception [3]. Virus structures exhibit certain meta-stability whose modulation permits fulfilling each task of the viral cycle on time thanks to their physicochemical properties. The basic construction of non-enveloped viruses consists of the capsid, a shell made up of repeating protein subunits (capsomers) or lipids, packing within the viral genome [1]. Far from being static structures, both enveloped and non-enveloped viruses are highly dynamic complexes that shuttle and deliver their genome from host to host in a fully automatic process. These natural designed capabilities have impelled using viral capsids as protein containers of artificial cargoes (drugs, polymers, enzymes, minerals) [2] with applications in biomedical and materials sciences. Both natural and artificial protein cages have to protect their cargo against a variety of physicochemical aggressive environments, including molecular impacts in highly crowded media [4], thermal and chemical stresses [5], and osmotic shocks [6]. Thus, it is important to use methodologies that supply information about protein cages stability not only under different environments, but also its evolution upon structural changes. Structural biology techniques such as electron microscopy (EM) and X-ray are used to unveil the structure–function interplay, revealing high resolution structures of protein cages [7]. However, these methodologies require a heavy average of millions of particles present in the crystal (X-ray) or thousands of structures for the model reconstruction (cryo-EM). Thus, they provide limited information on possible structural differences between individual particles that differs from the average structure. In addition, these approaches require conditions (i.e., vacuum) far away of those where protein shells are functional (liquid), precluding the characterization of protein shells dynamics and properties in real time. Indeed, the advent of single molecule technologies has demonstrated that mechanical properties of biological molecular aggregates are essential to their function [8]. The exploration of these properties would complement the structural biology methodologies (EM and X-ray) to find the structure–function-property interplay of virus structures. Atomic Force Microscopy (AFM) may not only characterize the structure of individual protein-made particles in liquid milieu, but also to obtain physicochemical properties of each one [9]. In addition, the nano-dissection abilities of AFM allows the local manipulation of protein shells to learn about their assembly/disassembly [10]. In this chapter, we give a general overview of how to apply AFM methods to protein shells. Our tour starts with a basic review of the recipes for attaching protein cages to solid surfaces. Afterwards we describe some modes for imaging protein shells with AFM and comment on inherent artifacts, such as geometrical dilation. We describe the ability of AFM to explore virions under changing environments in the liquid cell. Subsequently we describe the nanoindentation methodology, which probes the stiffness, breaking force, brittleness, and cargo unpacking of individual protein shells. Afterwards we focus on the effects of cyclic loading on individual particles (disassembly), and the AFM/fluorescence combination for exploring protein cages packing GFP.

Insights on AFM Methodology

Cantilevers. In AFM a microcantilever with a nanometric tip located at the very extreme actuates as a force transducer which palpates the viruses adsorbed on a solid surface. Cantilevers with different spring constants are mounted on a chip. For example Olympus OMCL-RC include a variety of four spring constants: 0.39, 0.76, 0.05 and 0.10 N/m that can be used with different viruses, depending on their stability and the adsorption strength to the surface. Figure 10.1a shows and optical microscopy image of the 0.05 N/m cantilever where the pyramid tip is encircled. Olympus stopped the production of OMCL-RC cantilevers. PNP-DB and QP-BioAC tips (Nanoandmore™) with similar properties can be used. The chip holding the cantilever is attached to the quartz window of the liquid cell which is created between the surface and the cantilever holder (Fig. 10.1b). For the shake of comparison with viruses, Tetraspeck™ (Thermofisher) plastic spheres of 100 nm in diameter are spread on a glass surface that are seen as white blobs with a size of ~250 nm because of the diffraction limit.

Fig. 10.1
a. A micrographic image of the tip of an instrument has a dimension of 40 micrometers. b. A laser beam is used to attack the viral particles. c. 3 tetrahedral-shaped viruses have a dimension of 100 nanometers shown from different orientations.

Attaching protein shells on surfaces. a Optical microscopy image of an 0.05 N/m OMRC cantilever, with the encircled tip. Plastic bead of 100 nm in diameter are adsorbed on the surface for the sake of comparison. b Cartoon of the liquid cell. Protein cages and cantilever are not in scale. c HOPG, glass and mica bare substrates before attaching the viruses. d hAdV particles on HOPG, glass and mica. e Individual hAdV particles showing twofold, threefold and fivefold symmetry orientations

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Immobilization of virus particles on surfaces. Virus particles are adsorbed to the surface by using physical forces (physisorption) with the substrate, including polar, non-polar, an van der Waals interactions [11]. Physisorption traps virus particles without inducing chemical bonds that might alter their structure [12]. Each virus species has individualized features such as local charge densities or hydrophobic patches [13] that can be used for anchoring the particles, via hydrophobic and/or electrostatic interactions, on different materials, such as glass, mica and HOPG (Highly Oriented Pyrolytic Graphite). Mica and HOPG surfaces are layered materials whose preparation consist of removing the last layer with adhesive tape, exposing a fresh surface ready for experiments. Care should be taken on leaving a flat surface by avoiding dangling whiskers on the area of the meniscus because they might crash with the cantilever. HOPG presents a non-polar surface and protein cages adsorb via hydrophobic interactions [14]. In the case of human adenovirus (HAdv) virus particles can be adsorbed on HOPG, silanized glass and mica with different results. HOPG collapses most of HAdv particles, indicating a strong non-polar (hydrophobic) interaction. However, silanized glass [9] reduces the attachment interaction and allows imaging intact icosahedral particles exhibiting fivefold, twofold and threefold symmetry orientations on the surface (Fig. 10.1c). Interestingly, using NiCl2 150 mM on mica [11] induce the adsorption of HAdv particles only at the threefold symmetry orientation, thus protein particles exhibit a triangular facet (Fig. 10.1c, right). Adsorption of virus particles on surfaces also may induce a reduction of the particle height [15, 16]. From a practical point of view predictions on proteins shells adsorption are difficult to make, and one uses the try-and-error methodology to find the best conditions.

Imaging. In AFM the tip scans the sample in x, y and z directions by using piezo actuators. While x and y scanners move in a pre-established way over a square region, the cantilever bends following the surface topography. Either the cantilever deflects perpendicularly to the surface applying a normal force (Fn) (Fig. 10.2a), or it bends laterally by torsion exerting a dragging force parallel to the surface (Fl) (Fig. 10.2b). Both Fn and Fl are monitored by focusing a laser beam at the end of the cantilever, whose reflection is registered in a four-quadrant photodiode. Thus, each pixel of the image located at a particular position of the planar coordinates (x, y), will be associated with certain bending values of the cantilever Fn and Fl. If the particle is not strongly enough attached or if it is too soft, it can be swept or modified under large bending forces. To avoid this effect as much as possible, a feedback loop is engaged to Fn to move the z piezo position in such way that Fn is kept constant. In this operational approach (contact mode, Fig. 10.2c), the AFM topography map will have x, y and z coordinates. The torsion Fl of the cantilever exerts about 40 times the perpendicular bending force Fn [17]. Individual virus particles are thus susceptible to undesired modifications by lateral forces. Their size of tens of nanometers offers a large topographical aspect ratio that is difficult to track by the feedback loop. A typical approach for surpassing this limitation is using fixation agents, such as glutaraldehyde. In such conditions AFM provide images whose resolution is comparable to that of some EM images [18]. Nevertheless, since glutaraldehyde structurally reinforces the specimens [19, 20], it precludes any characterization of dynamics or properties of intact native viruses, such as assembly/disassembly or physical properties [21]. Other approaches include developing imaging modes that avoid dragging forces as much as possible. In jumping mode (JM) [22], the lateral tip displacement occurs when the tip and sample are not in mechanical contact, thereby avoiding shear forces to a large extent (Fig. 10.2d). JM performs consecutive approach-release cycles at every pixel of the sample. In each cycle, known as force vs. distance z curve (FZ, Fig. 10.2e), the z-piezo approaches tip and sample from non-contact (label 1 at Fig. 10.2e) until establishing mechanical contact (label 2 at Fig. 10.2e) and reaching a certain feedback force (label 3 at Fig. 10.2e). After a few milliseconds, the z-piezo retracts about 100 nm until releasing the tip from the surface (label 1 at Fig. 10.2e). Subsequently the scanner moves laterally to the next pixel, and the process starts again. An AFM cantilever experiences a viscous drag while moving up and down in liquid, giving rise to a hysteresis loop (Fig. 10.2e) because forward and back curves do not coincide. Although AFM dynamic modes have also able of imaging protein shells in liquid conditions, it is difficult to control the applied force [23].

Fig. 10.2
5 illustrations. An illustration of a laser projected on a photodiode is in contact mode. The scan direction is horizontal in orientation. Another method used is the jumping mode of the laser on the plate. A normal force and a dragging force act on the viral particle called the viscous drag.

AFM working modes. a and b show the normal (Fn) and lateral forces concepts, respectively. c and d indicate contact and jumping modes, respectively. e Force curve of jumping mode in liquids, indicating the viscous drag and the important stages (see text)

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Tip dilation. The typical radius of the tip apex for usual cantilevers (OMCL-RC800PSA) is ~20 nm, and it is comparable to the curvature of the virus particles diameter. In this case, tip size plays an important role on the image resolution by inducing a lateral expansion, namely dilation, of the specimen [24]. Since dilation very often impairs high resolution in proteins, it is convenient to estimate how the tip-size is going to affect to AFM images. WSxM software [25] implements a geometrical dilation algorithm that allows simulating the dilation of protein shell’s structure. By using Chimera software [26] it is possible to access to a particular protein shell structure, such as the electron microscopy model. The “Surface Color By Height” option generates a gray-scale image that captures the topography variation in a given orientation. The TIFF format of this image can be imported by WSxM software and calibrated (www.wsxm.eu). The dilation algorithm input is the tip radius, and the dilated structure is calculated. Figure 10.3 exemplifies the dilation of phi29 bacteriophage EM structure [27]. Figure 10.3a shows the 3D rendered topography data of a bacteriophage phi29 on HOPG. By using the dilation algorithm with a 10 nm in diameter tip on the PDB model of phi29 phage [27] it is possible to obtain the dilated topography (Fig. 10.3b). Dilation strongly depends on the tip size, as shown in Fig. 10.3c. The darker area of Fig. 10.3b represents the extra dilated geometry of phi29 around the electron microscopy model (light colored).

Fig. 10.3
a and b. 2 illustrations of a spherical-shaped viral particle that has a 25 nanometer in dimension. c. An illustration of a viral shell that is oblong. The circumference of the tip is demonstrated with its radius from the center to the periphery.

Dilation effects in the phi29 bacteriophage shell. a represents the AFM image of a prohead. In b the bright color illustrates a EM structural model of phi29, and the dark area indicates the dilation corresponding to a tip of 10 nm in diameter. The cartoon of c indicates the dilation as a function of the tip size: dark, red and blue curves are the topographical profiles obtained with tips of 0.5 nm, 5 and 10 radius in diameter, respectively

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Applications

Imaging at different environment conditions. The conditions of the liquid environment in the liquid cell can be gradually changed for monitoring the stability of virus capsids over time. One pump connected to the inlet of the chamber inserts the buffer at the desired final concentration, and another pump connected to the outlet withdraws liquid at the same velocity of the inlet. In this particular case (Fig. 10.4) we observe the effect of a non-ionic detergent (IGEPAL® CA-630 (Sigma-Aldrich, Saint Louis, MI, USA, CAS: 9002-93-1)) on the structure of the coronavirus transmissible gastroenteritis virus (TGEV) [28], whose structure [29] and mechanical properties [30] are identical to SARS-CoV-2. Consecutive images of the same TGEV virus particle while increasing IGEPAL concentration in the liquid cell were taken (Fig. 10.4a). This virus was continuously imaged more than 40 times while IGEPAL was increased up to ~0.06% for 80 min. The virus particle remained stable at 0.020% IGEPAL, although at 0.023% some deterioration could be recognized. In frame #26, some material was ejected from the virus and a circular crack at the top appears. This damaged structure seems stable until frame #40. For statistical analysis, the height data evolution of eight particles subjected to similar process are plotted their individual (Fig. 10.4b, thin grey) and their average (Fig. 10.4b, thick dark) height variation over time. Consecutive images of virions in buffer without detergent were taken as control (Fig. 10.4b, blue), showing that their heights were not affected by AFM scanning. Figure 10.4c shows the effect of the pH variation on the stiffness of human adenovirus particles [31].

Fig. 10.4
6 microscopic images of a virus in 0%, 0.02%, 0.023%, 0.028%, 0.033%, and 0.04% magnification to view the core of the virus. A graph on delta h versus I G E P A L has a constant and a decreasing waveform. A graph on T versus p H has an intersection of 2 curves at 6 p H approximately.

Imaging while changing the environment in the liquid cell: effect of IGEPAL concentration on TGEV virions. a Topographical images were taken while the concentration of IGEPAL was increased over time. The frame number is shown in the upper left corner and the concentration of IGEPAL in the bottom-right. b Height loss during the time course assay. Experimental curves are shown in light red and the average curve of 8 observations is shown in red. As a control, the height of 1 virus particle without detergent is shown (black). Adapted from [32]. c Evolution of the spring constant with pH in human adenovirus. Adapted from [31]

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Mechanical properties of viruses: nanoindentation. Single force curve (FZ) experiments consist on pushing on the top of a selected protein shell (Fig. 10.5a). The FZ is executed on the particle at a typical speed of 50 nm/s to allow the water leaving the virus when it is squeezed [33]. After the contact between tip and particle is stablished, FZ typically shows an approximate linear behavior, which corresponds to the elastic regime of the shell and ascribes to the mixed bending of the cantilever and sample deformation (Fig. 10.5a, b, label 2). When the z-piezo elongation surpasses the critical indentation, particle breaks (Fig. 10.5, 1.1 nN) inducing a drastic decrease of the force, that resemble the penetration of the tip apex trough the virus (Fig. 10.5a, b, label 3). Afterwards the FZ is linear again and represents the cantilever bending on the solid substrate. By performing a FZ on the substrate (Fig. 10.5b, dotted line) and assuming that it is much more rigid that the cantilever, we can obtain the cantilever deformation. In this case, the human picobirnavirus particle [34] yields and breaks after the nanoindentation experiment (Fig. 10.5c). The subtraction of sample from substrate curves allows isolating the deformation of the virus cage (Fig. 10.5d). From these data we can obtain a few mechanical parameters: Fitting of the elastic part from 0 to 5 nm results in the stiffness or spring constant of virus shell (k = 0.25 N/m). The breaking or yield force is the force value when the elastic regime finish at 5 nm of indentation (Fb = 1.1 nN). The critical indentation δc, is the deformation of the virus when it breaks (5 nm). Thin shell theory relates the protein shell stiffness with the Young’s modulus as \(k\approx E\frac{{h}^{2}}{R}\), where h is the thickness of the shell and R its radius [35]. The area enclosed between forward and backward curves from indentation 0 up to 5 nm is the energy used to break the cage. In this case, it is about 8.8 nm × nN, i. e. 8.8 × 10–18 J or 2140 kBT, which approaches the order of magnitude of the total energy used for assembling the virus [36]. In addition, the critical strain εc = δc/h, where h is the initial height of the protein cage as measured with AFM, informs about the brittleness or the mechanical stability of protein cages [15]. In this case, εc = 5/36 = 0.14, saying that the particle breaks when it is deformed 14%.

Fig. 10.5
An illustration of a viral particle in contact with a cage. In a waveform on F n versus z piezo elongation, the wave has 3 phases and a peak followed by a sharp increase. Another waveform is f n versus indentation with 2 waves. Out of 2 waves, one has a peak, and both are increasing.

Single indentation assay. a Cartoon showing the three main stages during nanoindentation experiment on a protein cage: before contact (1), during deformation (2) and after breaking (3). b Evolution of Fn along the z-piezo elongation. Forward curve exhibits the three stages commented in a. c AFM topographies before and after nanoindentation showing a crack. d Nanoindentation data extracted from b, showing the shell deformation. Inset compares topographical profiles before (blue) and after (red) de experiment obtained at the line marked by the arrows in c. Inset shows the profile of the particle

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Genome externalization and mechanical properties of cargo. Nanoindentations at low force can induce cracks, allowing the access to the inner cargo of viruses. For instance, the consecutive application of nanoindentation cycles in HAdv cracks-open the shell in a controlled fashion to probe the mechanical properties of the core [37, 38]. These mechanical properties relate with the condensation state of dsDNA. In particular, the HAdv core is formed by 35 kbp dsDNA molecule bound to positively charged viral proteins that constitute ∼50% of the core molecular weight. Approximately about 500–800 copies of protein VII (pVII) contributes with 22 kDa to the core, becoming the majority protein. Consecutive nanoindentation experiments (Fig. 10.6) performed on wild type (HAdv-wt) and mutant lacking pVII (HAdv-PVII-) human adenovirus particles provide information on the pVII function. Figure 10.6a presents the typical evolution of HAdv-wt and HAdv-VII– particles in a multiple indentation experiment. The first indentation (FIC#0) on the intact shell (Fig. 10.6a, #0) opens a crack at the icosahedral facet with a similar size to the apex of the AFM tip (∼20 nm) (Fig. 10.6a, #1), allowing a direct access of the tip to the core. Subsequent indentations present a non-linear Hertzian behavior in contrast with the linear deformation found in intact particles (Fig. 10.5). During the first indentation after capsid cracking (Fig. 10.6a, #1), HAdv-wt cores are not deformed beyond 40 nm at 3 nN, while HAdv-VII– cores undergo larger indentations, up to ∼60 nm, at forces as low as 2 nN (Fig. 10.6b). This difference indicates that it is easier for the AFM tip to penetrate/deform HAdv-VII– than HAdv-wt cores. That is, in the absence of protein VII the HAdv core is softer and less condensed. Elongated structures with a height compatible with dsDNA strands (Fig. 10.6a, yellow) appeared on the substrate surrounding HAdv-VII– particles after the first fracture of the shell (Fig. 10.6a, #1, top), but not in HAdv-wt (Fig. 10.6a, #1, bottom). Material consistent with unpacked DNA seemed to be more abundant in HAdv-VII– particles that in HAdv-wt for the equivalent indentation number (Fig. 10.6a). This qualitative observation suggests that it is easier to exit the disrupted shell for the VII-free genome than for the VII-bound one. An alternative possibility would be that the VII-free genome has a larger tendency to adsorb to the mica substrate than the VII-bound DNA. Measurements of the topographical profile across the crater produced by the successive indentations (Fig. 10.6c) support the first possibility. Because topographical profiles include both the crater and rims, we used the lowest height inside the crack as an indicator of the remaining core contents (Fig. 10.6c, insets). Plotting the evolution of this parameter along six consecutive indentations for eight HAdv-VII– and six HAdv-wt particles (Fig. 10.6c) showed that, indeed, the core components are leaving faster the HAdv-VII– than the HAdv-wt cracked particles [38]. The application of consecutive indentations to coronavirus structures (TGEV), also inform about stability and genome release [32]. These experiments unveiled two different behaviors of TGEV particles: In the first case (Fig. 10.7a), a virus particle was probed seven times with maximum forces ranging from 0.6 to 3 nN, remaining unaltered through the alternating AFM images. In fact, ~65% of the explored virus particles (N = 69) kept their height constant (Fig. 10.7c, black), indicating that their structure was elastic and not affected by the nanoindentations. However, in the second case (Fig. 10.7b) and using the same FZ number and parameters, the TGEV particle showed evident structural changes consisting of appearing a circular crater after the first FZ that enlarged after consecutive deformations until the virus appears to have been ripped open, releasing its content (Fig. 10.7, #7). This behavior was observed in ~35% of the explored particles and was accompanied by a gradual decrease in height after each indentation (Fig. 10.7c, red). It is important to remark that the initial height of the particles that remained undamaged (indentation resistive) was larger than the damaged ones (indentation sensitive) (Fig. 10.7d), a difference that was significantly different at a 95% level of confidence. By assuming that all viruses package the same amount of genome, it is likely that everyone contains a similar number of RNPs, no matter their size. Therefore, the larger height of these particles (Fig. 10.7d) suggests a loosened core state represented by the ‘eggs-in-a-nest’ assembly [39]. In this case, when the tip retracts, RNPs recover their original positions and the flexible membrane returns to its previous height. In contrast, indentation-sensitive particles (Fig. 10.7b) exhibited lower heights than the resistive ones (Fig. 10.7d). The particle crack-opened with a permanent fracture (Fig. 10.7b) of ~25 nm in depth (Fig. 10.7c, blue). The lower height indicates a higher packing fraction (‘pyramid’ packing) [39] without internal free room, preventing the RNPs from rearranging under the mechanical stress induced by the tip. In fact, structural data of SARS-CoV-2 indicate this ‘pyramid’ packing occupies ~36% of the cavity, very close to viruses with high packing fraction values, such as lambda and phi29 bacteriophages [40]. Together with the AFM tip, solidly packed RNPs act as a “hammer and anvil” on the membrane, concentrating the mechanical stress on the virus envelope and causing permanent fractures (Fig. 10.7b). Consecutive imaging of the virus particles after each nanoindentation (Fig. 10.7b) reported increasing damage, with virus height loss from 25 to 100 nm (average decrease ~40 nm; Fig. 10.7c, red).

Fig. 10.6
12 microscopic images of the viral particle are disintegrated to view the internal structure of the virus. A graph on force versus indentation has an increasing trend. An error plot on height versus 2 forms of the virus has a curve that has multiple breaking points.

Mechanical properties of human adenovirus core: the role of protein VII. a Images showing the evolution HAdv-wt and HAdv-VII– particles during multiple indentation assays. Images are colored by height, as indicated by the color bar at the right hand side. b FZ curves corresponding to the first indentation performed on the core of 8 HAdv-wt (red) and 12 HAdv-VII– (blue) particles. c Evolution of the core height (as indicated by the minimum height in the crater, grey crosses) for eight HAdv-VII– and six HAdv-wt particles. Insets: Examples of topographical profiles obtained through the crater opened by the indentation Adapted from [38]

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Fig. 10.7
8 illustrations of the micrographs of a virus disintegrating slowly have a 60-nanometer dimension. A scatter plot on delta h versus F Z has an irregular trend. A box plot on height versus damaged and undamaged particles.

Mechanical properties of TGEV coronavirus: the role of packing fraction. a and b present topographical images of TGEV virions after consecutive indentations. Particles can be classified in two groups: indentation resistive (a) and indentation sensitive (b) as shown in the topo images. Frame number indicates how many indentations were performed. c Chart evolution of virus height as a function of the indentation number (FZ#). The values of height are taken from the indented region. Black and red charts represent indentation resistive an indentation sensitive, respectively. d Height box-plot distribution of intact virus particles. Adapted from [32]

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Mechanical fatigue, disassembly and aging. The breaking force describes the largest force that the virus can survive. Single indentation experiments collapse the particle by inducing large and uncontrollable changes in its structure. It is thus difficult to derive consequences about disassembly, since in the cycle of many virus shells, for instance, disassembly takes place by losing individual capsomers in an gradual way [41]. A protein cage must also resist a constant barrage of sub-lethal collisions in crowded environments [42]. Equipartition theorem provides an estimation of the energy transferred in a molecular collision to be ~\(\frac{3}{2}{k}_{B}T\), which is far below that the energy supplied by single indentation assay experiments. Imaging of individual protein shells with AFM in jumping mode requires thousands of load cycles (FZs) at low force (~100 pN per pixel, Fig. 10.2c) [22]. A rough estimation indicates that ~10 \({k}_{B}T\) is transferred to the particle at every cycle [43], very close to the molecular collisions value. The continuous imaging of a particle enables the evaluation of any structural alteration while subjected to cycle load at low forces. Mechanical fatigue experiments have demonstrated to be a disassembly agent able of recapitulating the natural pathway of adenovirus uncoating [22]. Therefore fatigue provides additional information by providing information on shell stability under multiple deformation cycles at low force (~100 pN) [44], well below the breaking force (Fig. 10.5d). Let us exemplify the mechanical fatigue methodology in the case study of HAdv [45]. This virus travels to the cytoplasm until the nuclear pore to deliver its genome. Inside the host cell, the virus finely tunes the sequential loss of pentons (proteins located at the icosahedron corners) to render a semi disrupted capsid at the nuclear pore. Viruses with too many pentons would not be able to release their DNA through the cell’s nuclear pore, whereas those with too few pentons would liberate their genome before reaching the nucleus. Mechanical fatigue induces penton failure [44] (Fig. 10.8a), which mimics the stresses the virus sustains during its journey to the nucleus. This approach allows to study the transition kinetics of penton release as a two-state process, from where it is possible derive the spontaneous escape rate and the free energy barrier of a penton [45]. Moreover, fitting the survival probability P of penton N (Fig. 10.8b) under different fatigue forces to Weibull analysis: \(P\left(N\right)=1-\mathrm{exp}\left[-{\left(\frac{N+1}{\lambda }\right)}^{\beta }\right]\), where λ is the half-life and β determines the nature of the penton’s dynamics release. In particular, β < 1 would imply that failure rate decreases over time (Lindy effect). This effect can be found in ancient monuments, since the older something is, the more likely it will survive. The case of β = 1 would imply a Poisson process with a constant probability of penton failure over time. However, β > 1 (Fig. 10.8b) involves the phenomenon termed in materials science as aging, which means that the failure rate increases over time. This indicates that the probability for the virus of losing a penton increases if other penton has been lost before. This aging process (β > 1) accelerates the overall penton escape rate by about 50% (Fig. 10.8c) with respect to a sequence of independent escape events (β = 1). Specifically, while the first penton resists about 60 min, the second and third would last 30 and 20 min more, respectively (Fig. 10.8c). In this way, aging guarantees to have the virus particle just disrupted enough to release the genome trough the nuclear pore. Without aging, virus particle would be still quite complete and the genome could not escape (Fig. 10.8c), with negative consequences for infection.

Fig. 10.8
An illustration of a viral particle inside a nucleus that acts with aging and without aging. The R N A inside the nucleocapsid is released into the nucleus after 90 minutes. The laser points are observed at an interval of 20, 30, and 60 minutes. A graph on failure probability versus time.

Mechanical fatigue and aging of human adenovirus. a AFM of a HAdv particle showing three vacancies of pentons on the triangular facet (encircled). b Failure probability of pentons overt time for different forces. Inset show the fitting parameters of Weibull statistics. c Diagram showing the biological implications of aging on the virus infectivity (see text). Adapted from [45]

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Layer-by-layer disassembly of a multilayered virus with mechanical fatigue. Several viruses wrap their genome in several layers of protective casing, resulting in an onion-like structure. For example, the rotavirus (RV), that can induce severe diarrhea in young children, have three layers built of different proteins with various functionalities. Rotavirus subparticles may exist with only one or two of these coats, which allows to study each layer in detail [46]. Mechanical fatigue allows to explore the strength and binding of each layer (Fig. 10.9a). The RV infectious particle is a 100 nm non-enveloped triple-layered particle (TLP) composed of three concentric protein shells enclosing the dsRNA genome and the viral RNA polymerase and capping enzyme. The inner layer surrounds the eleven dsRNA genomic segments associated with the RNA-dependent RNA-polymerase VP1 (125 kDa) and the RNA-capping enzyme VP3 (88 kDa) at the pentameric positions. This thin single-layered particle (SLP), an intermediate structure that is involved in the packing and replication of the viral genome, is surrounded by a thick layer formed by 260 VP6 pear-shaped trimers (45 kDa) in the double-layered particle (DLP). This particle, which does not disassemble during the infection, constitutes the transcriptional machinery that initiates the core steps of the viral replication cycle once delivered in the host cell cytoplasm. Cyclic imaging of the TLP (Fig. 10.9b, left) at forces between 100 to 200 pN per pixel shows that, while the VP4 spikes are removed from the particle surface in a few frames (Fig. 10.9b, middle), the VP7 layer remains mostly intact (Fig. 10.9b, right) during 80 frames (light red in Fig. 10.9f). These results illustrate that the spikes are easily removed by the AFM tip and are not strongly anchored. However, the VP7 layer displays a strong resistance against fatigue. A strong binding energy between capsomers would not only result in a high resistance of individual proteins against fatigue, but also will contribute to a high breaking force when all capsomers are probed in a single indentation assay experiment [46]. The current model proposes a calcium concentration drop in endosomal compartments during RV entry as the factor that triggers VP7 disassembly and membrane penetration. In fact, calcium depletion by chelating agents (as EDTA) is used to uncoat TLP to DLP by inducing VP7 trimer dissociation. To explore the structural consequences of this process in real time, fatigue assays are carried out on TLP while EDTA simultaneously flowed in the AFM liquid chamber, to induce the gradual depletion of Ca ions of the particles (Fig. 10.9c). In these conditions, fatigue induces the neat VP7 detachment from the VP6 subjacent layer (indicated by a circle in Fig. 10.9c, #19) even before the spikes are removed. Indeed, the evolution of the topographic profiles (dark red in Fig. 10.9f) show abrupt downwards steps very close to the VP7 thickness (red arrow of Fig. 10.9f) indicating that TLP particle loses VP7 completely while keeping VP6. These results not only suggest that Ca ions mediate the interaction between VP7 and VP6 layers, but also that the absence of ions weakens the interaction between VP7 subunits. If fatigue continues, VP6 subunits are neatly removed from VP2 layer (circle in Fig. 10.9c, #29). Therefore, VP6 layer appears forming a weak shell whose interaction with the beneath VP2 layer is not very strong, since it peels off rapidly to reveal the SLP. Similar results are found on DLP. Again, fatigue induced a clean VP6 disassembly after less than 10 frames (circle in Fig. 10.9d, #8). In this case the evolution of the topographic profiles (blue in Fig. 10.9f) undergoes sharp reductions very close to the VP6 thickness, inducing the gradual uncovering of the innermost VP2 (blue arrow in Fig. 10.9f). These experiments not only illustrate a weak interaction between VP6 and VP2 layers, but also a very feeble VP6-VP6 binding energy. Finally, the thin SLP VP2 is highly unstable under fatigue (Fig. 10.9e) experiments collapsing well before reaching 10 frames (green in Fig. 10.9f).

Fig. 10.9
13 illustrations of the viral particles with laser given by A F M tip. The disintegration of the coating of viral particles is seen in the 12 microscopic images below. A graph on height variation versus frame number. T L P and D L P waveforms are increasing initially, then become constant.

Peeling a virus layer by layer with mechanical fatigue. a cartoon of the peeling experiment. Topographic evolution of TLP (b), TLP + EDTA (c), DLP (d) and SLP (e) during continuous imaging at low force (~60–120 pN) indicating the corresponding frame. f Topographic evolutions obtained at center of each virus particle (b–e) in TLP (red), DLP (blue) and SLP (green) particles. Dark red color indicates fatigue of TLP + EDTA. Red and blue arrows indicate the loss of height from TLP to DLP and for DLP to SLP, respectively. Adapted from [46]

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AFM/fluorescence combination. Here we discuss the methodology for studying the activity of individual nanoreactors based on a protein container. About 220 GFPs are packed inside P22 bacteriophage capsids [47] and its functionality is explored as a function of the mechanical stress applied with the AFM tip to the virus particle while monitoring fluorescence with Total Internal Reflection Fluorescence Microscopy (TIRFM) [48]. The integration of a single molecule fluorescence microscope with AFM requires to monitor the fluorescence signal at the surface to avoid not only the background signal of the AFM probe itself, but also the light coming from the bulk solution (Fig. 10.10a). In this approach [49] P22 bacteriophages are immobilized on a glass surface, in such way that a region with particles visualized with AFM (Fig. 10.10b, left) is correlated with their TIRFM pattern (Fig. 10.10b, right). The diffraction-limited resolution of the optical system restricted the size of the smallest spot to λ/2 (≈250 nm), which is larger than the P22 VLPs. Consequently, the particles that were too close together (arrow, Fig. 10.10b, left) appeared as a single fluorescence spot (bottom right, Fig. 10.10b, right). The inset of Fig. 10.10b (right) shows the topographic (white) and light (green) profiles of two VLPs (Fig. 10.10b, white and green dotted lines) where the particle width in the fluorescence signal is ≈500 nm, approximately five times larger than in the AFM topography. After localizing an intact virus showing fluorescence signal, it is possible to execute a nanoindentation experiment (Fig. 10.10c, black) by using a Si3N4 tip to avoid electronic quenching of GFP and isolate pure mechanical effects [49]. The fluorescence remained constant (Fig. 10.10c) during the tip approach (Fig. 10.5a, label 1). Furthermore, the light signal stayed stable even when the tip established contact with the VLP at 1 s and linearly deformed the VLP structure (Fig. 10.10c). Fluorescence quenching started at 1.2 s, coinciding with the force steps that indicated the yielding and/or breaking of the protein shell, reaching a minimum value once the particle stopped collapsing (2 s). At 2.6 s the AFM tip moved backwards, away from the surface, and the fluorescence signal remained stable until the tip was released from the VLP (3 s), coming back to a similar value to the initial magnitude (4 s). This fluoresce reduction is most likely due to changes in the integrity of the packed GFPs. In particular, GFPs can oligomerize inside the P22 cavity (internal radius of 22.2 nm) due to crowding phenomena. In the P22 prohead system there are ≈209 GFPs attached internally to the capsid with an, resulting in a nearest neighbor distance of ≈3 nm between the protein centers. Since the GFP Stokes radius is 2.8 nm, we derived an average distance of 2 Å between fluorophores. It is possible to simulate the compression of a single P22 particle between two planes (Fig. 10.10d, left). The compressed particle shows that there are around 10 unfolded GFPs inside (Fig. 10.10d, right –red-) which account for the amount of quenched light (Fig. 10.10c, green).

Fig. 10.10
An illustration of A F M tip on V L Ps over a glass surface. 2 micrographs indicate the oblong particles and bright spots of the viruses are seen. A graph on force versus time versus fluorescence has 2 waveforms. The wave for fluorescence decreases while it peaks at 2.5 seconds for force.

AFM/fluorescence combination: quenching of packed GFP inside P22 protein cages. a Presents a diagram of the AFM/TIRF system, showing the P22 VLPs (light green) and the AFM probed particle in dark green. b Simultaneous AFM and fluorescence images (right and left, respectively) of individual VLPs on the glass surface. The white arrow (left) indicates a couple of particles appearing as one spot in fluorescence (right). The inset in b-left shows a high-resolution AFM image of P22 VLP resolving the proteinaceous structure of the virus. Inset of b-right compares the lateral resolution of both microscopies by plotting the signal profile of the AFM topography (white) and the fluorescence (green) obtained at the lines indicated by the dashed white and green lines, respectively. c Chart representing the simultaneous signals of force (black) and light (green). d A simulation of the GFP aggregates (green) inside a P22 procapsid (blue) before deformation (left). In this 3D image, one half of the capsid has been removed to show the internal configuration of the GFPs located below the virus lumen. The right panel shows a deformed virus where the partially unfolded GFPs are highlighted in red. Adapted from [49]

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Conclusion and Perspective

Bacteriophage phi29 was the first virus structure to be studied with a scanning probe technique [50], one year before the invention of AFM [51]. A few years later AFM was a generalized technique for studying biological samples [52] at single molecule level. AFM was initially applied just for imaging viruses [53, 54], but it was also used to study the biophysical properties of viruses [9]. During the last 20 years AFM is gradually being considered as a tool for virus research, although is not yet at the level of the classical structure techniques such as electron or X-ray microscopies. Using AFM for measuring the binding forces of virus to cells [55] or for the assembly of 2D virus-like assemblies in real time [56] are also promising applications of AFM that, hopefully, will convince virologists that AFM is an equally valid approach for studying viruses. We hope that the applications of AFM exposed in this chapter will help in this task.