Submolecular-Scale Imaging of α-Helices and C-Terminal Domains of Tubulins by Frequency Modulation Atomic Force Microscopy in Liquid

Abstract

In this study, we directly imaged subnanometer-scale structures of tubulins by performing frequency modulation atomic force microscopy (FM-AFM) in liquid. Individual α-helices at the surface of a tubulin protofilament were imaged as periodic corrugations with a spacing of 0.53 nm, which corresponds to the common pitch of an α-helix backbone (0.54 nm). The identification of individual α-helices allowed us to determine the orientation of the deposited tubulin protofilament. As a result, C-terminal domains of tubulins were identified as protrusions with a height of 0.4 nm from the surface of the tubulin. The imaging mechanism for the observed subnanometer-scale contrasts is discussed in relation to the possible structures of the C-terminal domains. Because the C-terminal domains are chemically modified to regulate the interactions between tubulins and other biomolecules (e.g., motor proteins and microtubule-associated proteins), detailed structural information on individual C-terminal domains is valuable for understanding such regulation mechanisms. The results obtained in this study demonstrate that FM-AFM is capable of visualizing the structural variation of tubulins with subnanometer resolution. This is an important first step toward using FM-AFM to analyze the functions of tubulins.

Introduction

X-ray and electron crystallography are well-established techniques for the structural analysis of proteins with atomic-scale resolution. For example, Nogales and co-workers (1,2) determined the atomic-scale structure of an αβ-tubulin heterodimer in a Zn-sheet structure by electron crystallography. Subsequently, Gigant et al. (3) investigated the structure of tubulin complexes with stathmin-like domains by x-ray crystallography. NMR, electron microscopy (EM), and electron tomography have also been used to visualize protein structures with angstrom resolution. For example, Sosa et al. (4) and Kikkawa et al. (5) investigated the binding structures of motor proteins on the surface of tubulins by cryo-EM. In addition, Sui and Downing (6) visualized the doublet structures of tubulin microtubules by cryo-electron tomography.

Atomic force microscopy (AFM) (7,8) has also been an important tool for investigating the surface structures of proteins. AFM has some advantages in the structural analysis of proteins. For example, AFM imaging does not require chemical modification of a sample, such as staining or isotopic modification. Moreover, it can be operated in solution. Therefore, various proteins have been investigated by AFM with nanometer and subnanometer resolution. Contact-mode (CM)-AFM has been used to investigate various protein structures, including purified proteins and native membrane proteins (9–15). For example, Karrasch et al. (11) reported CM-AFM images of intermediate filaments and bacteriophages that were covalently fixed on glass substrates. Among various studies on membrane proteins, Müller and co-workers (16,17) imaged the structural changes and flexibilities of a bacteriorhodopsin with subnanometer resolution. Hoh et al. (10) investigated individual connexons of gap junctions in native membranes isolated from a rat liver. These CM-AFM studies in liquids have allowed us to determine the functional structures of proteins in their natural environment, as well as the structural changes at work (18).

One of the drawbacks of CM-AFM is that it cannot be used to image isolated biomolecular structures weakly attached to a substrate, because of the influence of the lateral friction force during imaging. To overcome this problem, investigators have used amplitude modulation (AM)-AFM (19,20). The lateral friction force is dramatically reduced by oscillating a cantilever at its resonance frequency in AM-AFM. Using AM-AFM in liquid, researchers have investigated a variety of isolated proteins. For example, Kasas et al. (21) used AM-AFM to investigate the molecular-scale structures of RNA polymerases and their activity to produce RNAs from nucleotides on mica. Their results demonstrated the ability of AM-AFM to monitor the biological activities of proteins that are weakly attached to a substrate. In another study, Möller et al. (22) used AM-AFM images of purple membranes and hexagonally packed intermediate layers to demonstrate the capability of AM-AFM for subnanometer-scale imaging of proteins in a liquid. Many other applications of CM- and AM-AFM in protein analyses have been thoroughly reviewed in previous works (23–25). In addition to the large number of biological applications of CM- and AM-AFM, investigators have developed novel imaging modes of AFM (e.g., bimodal AFM (26,27) and jumping-mode AFM (28)) to improve the performance of AFM techniques.

The above-mentioned studies showed the ability of AFM to analyze the structures and mechanical properties of proteins with subnanometer resolution in liquid. However, it is often difficult to assign a submolecular-scale contrast to a specific structure of a protein when the orientation of the deposited protein is unknown. Therefore, the orientation of randomly deposited proteins must be determined for submolecular-scale investigations of proteins by AFM. To determine the orientation of proteins by AFM, it is necessary to specify structural features such as secondary structures at the surfaces of proteins.

Recent progress in frequency modulation (FM)-AFM (29) has made it possible to perform true atomic-resolution imaging in liquid with piconewton-order loading forces (30,31). In addition, investigators have demonstrated the applicability of FM-AFM to biological studies by imaging molecular- and submolecular-scale structures of various biological systems (32–37). Hoogenboom and co-workers (32,38) investigated membrane proteins (e.g., bacteriorhodopsin) and voltage-dependent anion channels in native membranes by FM-AFM in liquid. In addition, Fukuma et al. (34) imaged individual β-strands constituting an isolated amyloid fibril deposited on a mica surface. This result demonstrated the unique capability of FM-AFM to visualize secondary structures of proteins.

Tubulins are globular proteins with a diameter of ∼4 nm (1). The αβ-tubulin heterodimer is known as a common building block of microtubules, which are cytoskeletons with a diameter of 25 nm. Microtubules serve as structural components within cells and are involved in many cellular processes. For example, microtubules act as a molecular rail for motor proteins, such as kinesin and dynein, in intracellular transportation. Previous studies suggested that such cellular functions of tubulin microtubules are regulated by post-translational modifications of the C-terminal domains of tubulins (39–41). In spite of their importance, the detailed conformation of tubulin C-terminal domains and its relation to their functions have not been clarified, even by well-established techniques such as x-ray crystallography and NMR. Although this has been ascribed to the local variations or fluctuations of tubulin C-terminal domains, direct evidence for either of these models has yet to be presented.

AFM has been used for the investigation of microtubules and other tubulin structures, such as protofilaments. Fritz et al. (20) and other groups (42,43) investigated the surface structures and elasticity of microtubules by AFM in liquid. In addition, Elie-Caille et al. (44) reported that AM-AFM imaging of tubulin protofilaments revealed the effect of paclitaxel (taxol) on molecular arrangements. They also showed that molecular-scale structures and their arrangements in tubulin protofilaments can be quantitatively analyzed by AM-AFM in liquid.

In this study, we investigated the surface structures of tubulins by FM-AFM in liquid to show the applicability of FM-AFM for identifying subnanometer-scale structures at the surfaces of tubulins. On the basis of the submolecular-scale AFM images of tubulin protofilaments, we discuss the arrangement of α-helices and C-terminal domains at the surface of tubulins.

Materials and Methods

Sample preparation

The tubulins used in this study were purified from pig brains through two cycles of polymerization-depolymerization and phosphocellulose column chromatography (45). The stock solution of the tubulins was diluted with buffer solution to a concentration of 1 mg ml⁻¹. In this study, two types of buffer solution were used to prepare tubulin protofilaments and sheet-like structures. PEM-G buffer solution (80 mM PIPES, 1 mM MgCl₂, 1 mM EGTA, 1 mM GTP, pH 6.8) was used for the preparation of tubulin protofilaments. For imaging of tubulin sheet-like structures, MES-Zn buffer solution (140 mM MES, 0.7 mM MgCl₂, 0.3 mM EGTA, 1 mM GTP, 0.7 mM ZnCl₂, pH 5.8) was used. The tubulin solution (1 mg ml⁻¹) was incubated at 37°C for 30 min to form tubulin protofilaments or sheet-like structures by polymerization. After the addition of 10 μM taxol, the solution was ultracentrifuged at 37°C and 87,000 × g for 10 min. The precipitate was depolymerized by incubation at 4°C for 1 h followed by sedimentation. The supernatant was incubated at 37°C for 30 min, and the tubulin structures were then stabilized by the addition of 10 μM taxol.

For FM-AFM imaging, tubulin solution (200 μl) was deposited onto a freshly cleaved mica surface (ϕ12 mm). The sample was incubated at room temperature (25°C) for 30 min and rinsed with PEM-G or MES-Zn buffer solution for the preparation of protofilaments or sheet-like structures, respectively.

The size distribution of tubulin structures in the buffer solution was analyzed with a laser diffraction particle size analyzer (ELSZ-2; Otsuka Electron, Tokyo, Japan). The system can detect particles with sizes from 0.6 nm to 7 μm, which is sufficient to identify different types of tubulin structures, such as heterodimers, protofilaments, and microtubules. The tubulin solution was maintained at 37°C in a disposable plastic cell during the measurement of the size distribution.

FM-AFM imaging

We performed AFM measurements using an in-house-built, ultralow-noise FM atomic force microscope (30) combined with a commercially available AFM controller (Nanonis RC-4, SPECS Zurich GmbH, Zurich, Switzerland). All AFM experiments were performed at room temperature (25°C) in PEM-G buffer solution or MES-Zn buffer solution. A commercially available silicon cantilever (PPP-NCH; Nanoworld, Headquarters, Switzerland) with a nominal spring constant of 42 N m⁻¹ and a resonance frequency of 150 kHz in liquid was used. A phase-locked loop circuit (Nanonis OC-4; SPECS) was used to detect the frequency shift and oscillate the cantilever at its resonance frequency with a constant amplitude.

Results and Discussion

Molecular-scale imaging of tubulin protofilaments

Fig. 1 a shows the size distribution of tubulin structures in the tubulin solution measured by the dynamic light scattering method. Peak i at 9.1 nm corresponds to the length of a single αβ-tubulin heterodimer. In the microtubule formation, αβ-tubulin heterodimers form a linear repeating structure with a head-to-tail arrangement, which is known as a tubulin protofilament. Thirteen to 15 protofilaments are aligned in parallel to form a microtubule with a cylindrical structure in vivo and in vitro. The existence of peak ii at 165 nm and peak iii at 1128 nm suggests that the tubulin solution contains at least two other types of tubulin structures, which are probably protofilaments and microtubules.

Figure 1.

(a) Size distribution of tubulin structures in the PEM-G buffer solution used in this experiment. (b) FM-AFM image of tubulin protofilaments deposited on mica using the same PEM-G buffer solution (Δf = −10 Hz, A = 0.38 nm, v = 200 nm/s).

Fig. 1 b shows an FM-AFM image of tubulin structures deposited onto cleaved mica using the same tubulin solution, which contains αβ-heterodimers, protofilaments, and microtubules. A number of fibrillar structures with a length of 30–200 nm are found in the AFM image. The width of the fibrillar structures (∼9.9 nm) is smaller than the diameter of a microtubule (25 nm). According to previous AM-AFM studies by Elie-Caille et al. (44) and Hamon et al. (46), tubulin protofilaments are adsorbed on cleaved mica without surface modification. In contrast, other authors (20,47) reported that microtubules are not readily attached to negatively charged surfaces, such as mica and glass, owing to electrostatic repulsion between the negatively charged microtubule surfaces and the negatively charged substrate. Thus, the tubulin structures found in the AFM image are mostly tubulin protofilaments, although the solution contains other tubulin structures.

Fig. 2, a and b, show an FM-AFM image of an isolated tubulin protofilament on mica and a model of a protofilament consisting of three αβ-tubulin heterodimers, respectively. The height profiles along lines A-B and C-D in Fig. 2 a are respectively shown in Fig. 2, c and d. As indicated by arrows in Fig. 2, a and d, the surface of the protofilament exhibits a periodic corrugation with a periodicity of ∼4 nm, in agreement with the diameter of a tubulin monomer. The average spacing of 4 nm also shows good agreement with previous results obtained by AM-AFM and EM (44,48). In addition, the height of the protofilament is also 4 nm, as shown in Fig. 2 c. In contrast, the apparent width of the protofilament in the FM-AFM image is 9.9 nm (as shown in Fig. 2 c), which is more than two times larger than the actual diameter of a protofilament (∼4 nm). This increase in the width of the fibrillar structure can be explained by the effect of the nanoscale tip geometry (49). Thus, these results demonstrate that individual tubulins constituting a protofilament can be directly imaged by FM-AFM in a physiologically relevant solution.

Figure 2.

(a) FM-AFM image of an isolated tubulin protofilament (Δf = +210 Hz, A = 1.44 nm, v = 200 nm/s). (b) Structural model of a tubulin protofilament. Height profiles were measured along lines (c) A-B and (d) C-D in panel a.

In this experiment, we were not able to obtain a submolecular-scale FM-AFM image of an isolated protofilament, although we endeavored to optimize the imaging parameters, such as the amplitude of cantilever vibration (A), the frequency shift (Δf), and the tip velocity (v), to improve the resolution of the image. In addition, we often observed the dissociation of tubulins from a protofilament during imaging. In the previous study, Elie-Caille et al. (44) achieved stable, high-resolution imaging of protofilaments deposited on mica by AM-AFM using cantilevers with k = 0.3 and 0.6 N m⁻¹. However, in our experiment, we found it difficult to perform such stable, high-resolution imaging of protofilaments. This may be because we used a relatively stiff cantilever (k = 42 N m⁻¹), and hence a transient feedback error may have caused a large loading force to be applied to the sample. Although one can increase the rigidity of a protofilament by chemical modification to bridge adjacent tubulins using linker molecules, the submolecular-scale features of its surface are also likely to be modified. Therefore, such chemical modification is not desirable for submolecular-scale AFM studies.

Submolecular-scale imaging of tubulin sheet-like structures

Fig. 3 a shows FM-AFM images of tubulin protofilaments prepared with MES-Zn buffer solution. As previously reported (1,50,51), tubulins form Zn-sheet structures in MES-Zn buffer solution. Zn ions are known to bridge tubulin protofilaments and help form a large Zn-sheet structure. However, in this experiment, we found small sheet-like structures in which protofilaments were aligned in parallel, as shown in Fig. 3 b. Fig. 3 c shows a height profile measured along the axis of a protofilament (line A-B in Fig. 3 b). The profile shows a periodic corrugation with a spacing of ∼4 nm, which agrees with that of an isolated protofilament (see Fig. 2 a). Although the tubulin structures observed in the large-scale AFM image (Fig. 3 a) look similar to the protofilaments in Fig. 1 b, we found a difference between them in high-resolution AFM images with a small scanning size (Fig. 3 b). In this study, we were unable to observe subnanometer-scale contrast at the surface of protofilaments prepared with PEM-G buffer solution, whereas we did observe such contrast on the sheet-like structures prepared with MES-Zn buffer solution. As described above, it is likely that the submolecular-scale imaging of isolated protofilaments was hindered by the high loading force resulting from the use of a stiff cantilever. Therefore, we speculate that the subnanometer-scale imaging of protofilaments in the sheet-like structures was possible because of the stabilization by Zn ions bridging adjacent protofilaments.

Figure 3.

(a) FM-AFM images of tubulin structures obtained in MES-Zn buffer solution (Δf = +25 Hz, A = 0.38 nm, v = 300 nm/s). (b) FM-AFM image of a tubulin sheet-like structure (Δf = 0.0 Hz, A = 0.30 nm, v =100 nm/s). (c) Height profile measured along line A-B in b. (d) Cutout from panel b as indicated by the dashed line in b.

A part of Fig. 3 b is magnified in Fig. 3 d to highlight the detailed subnanometer-scale contrasts observed on the tubulin sheet-like structure. The length of the scale bar in Fig. 3 d corresponds to the diameter of a tubulin (4 nm). In this experiment, we used cantilever vibration with a very small oscillation amplitude (A < 0.5 nm) to enhance the sensitivity of the frequency shift signal to the short-range interaction forces acting between the front atoms of the tip and the sample. Thus, we were able to obtain subnanometer-resolution images even with a tip diameter of a few nanometers. The image shown in Fig. 3 d demonstrates that subnanometer-scale structures within individual tubulins can be visualized by FM-AFM. The observed subnanometer-scale contrasts show a large variation. We speculate that this is caused by the difference in the rotational orientation of the protofilaments.

Although we were not able to assign all of the subnanometer-scale contrasts to a specific tubulin structure, we were able to do so in some cases, as shown in Fig. 4 a, where the rotational orientation was determined from the AFM image. In Fig. 4 a, one of the protofilaments constituting the tubulin sheet-like structure was imaged with a smaller scan size. The image shows nanoscale repeating structures corresponding to the tubulin monomers as well as subnanometer-scale features within each molecule. In particular, the fine features indicated by the arrows in Fig. 4 a exhibit a periodic corrugation with a spacing of 0.53 nm, as revealed by a cross-sectional plot taken along line A-B (Fig. 4 b). This average spacing is in agreement with the common pitch of an α-helix backbone (0.54 nm), as illustrated in Fig. 4 c, which strongly suggests that the periodic features correspond to α-helices at the tubulin surface.

Figure 4.

(a) FM-AFM image of a tubulin protofilament in a sheet-like structure (Δf = +3.0 Hz, A = 0.30 nm, v = 100 nm/s). (b) Height profile measured along line A-B in panel a (average spacing: 0.53 nm, standard deviation: 0.056, n = 12). (c) Schematic illustration of α-helix backbone. (d) Structural model of a tubulin heterodimer (PDB ID: 1JFF).

An important finding here is that the backbones of α-helices can be imaged by FM-AFM with clear contrast, even though the backbones are buried under fluctuating side chains. This result can be explained by the rigidity of an α-helix backbone. In general, an α-helix backbone is stabilized by three to four parallel hydrogen bonds per turn. Thus, a backbone with a helical structure is more rigid than the fluctuating side chains. Therefore, the interaction between the tip and the α-helix backbone may predominantly contribute to the formation of the image contrasts, whereas the interaction between the tip and the side chains may be smeared out by their random motion, leading to almost no image contrast.

The direct imaging of α-helices provides useful information for identifying the orientation of a tubulin protofilament. Although the structural model of a tubulin protofilament is known, to interpret the subnanometer-scale features in an FM-AFM image, one must determine the rotation angle of the protofilament. We compared the subnanometer-resolution FM-AFM image (Fig. 4 a) with the known structural model of the tubulin heterodimer (PDB ID: 1JFF) with various rotation angles. As a result, we found that the arrangement of the observed α-helices exhibits the best agreement with that of the structural model when the rotation angle is set as shown in Fig. 4 d. The red regions of the structural model represent α-helices at the surface of tubulins. Although it is difficult to distinguish between α- and β-tubulins in the FM-AFM image, the result allowed us to determine the directions of the (+) and (−) ends of the tubulin protofilament, as shown in Fig. 4 a.

Moreover, the good agreement between the AFM image and the structural model allowed us to interpret local features in the high-resolution FM-AFM image. The positions of the bright protrusions observed in the FM-AFM image (Fig. 4 a) agree with those of the C-terminal domains in the structural model (Fig. 4 d). This result demonstrates that the C-terminal domains of tubulins can be directly imaged by FM-AFM in liquid.

To enhance the contrast of the bright protrusions, we adjusted the color scale of the AFM image as shown in Fig. 5 a. In addition, the same FM-AFM image is shown with a 21-step color scale in Fig. 5 b for comparison of the subnanometer-scale structures within the C-terminal domains between different molecules. The AFM image shows that the bright protrusions have similar oval shapes, and the longer and shorter axes have lengths of 2–3 and 1–2 nm, respectively. The height of the protrusions is almost uniform and is ∼0.4 nm from the surface of the tubulins. However, small corrugations within the C-terminal domains exhibit variation depending on the molecules, as shown in Fig. 5 b. The results demonstrate the capability of FM-AFM to visualize structural differences between different C-terminal domains with subnanometer-scale resolution.

Figure 5.

(a) Cutout from the FM-AFM image shown in Fig. 4a. The color scale was adjusted to highlight the C-terminal domains. (b) The same image shown with a 21-step color scale. (c and d) Schematic illustrations to explain the possible imaging mechanisms of the C-terminal domains.

According to previous studies (52–54), a C-terminal domain of a tubulin consists of ∼10–25 residues and varies in length and constituent amino acids owing to the existence of different isotypes. In one of the structural models (54,55), the C-terminal domain is considered to protrude from the tubulin surface a few nanometers into the solution with significant fluctuations. Although the height of the protrusions in the AFM image (0.4 nm) is lower than the expected height of the C-terminal domain in the extended form, the tip may be scanned over the fixed end of the C-terminal domain during AFM imaging, as shown in Fig. 5 c.

In contrast, previous NMR studies using a synthetic short peptide as a model of a C-terminal domain suggested that the peptide model has a relatively compact structure under a low pH condition (pH 5), owing to the helical conformation formed by the several residues (55). Because the subnanometer-scale FM-AFM image (Fig. 4 a) was obtained under a slightly acidic condition (MES-Zn buffer, pH 5.8), it is possible that the C-terminal domains were taking a folded structure as illustrated in Fig. 5 d.

Previous studies on microtubules showed that the polyglutamylation of C-terminal domains at the surfaces of tubulins affects the binding of kinesins and microtubule-associated proteins to microtubules (39,56–58). These molecular interactions play an important role in the major functions of microtubules, including intracellular transport and cell division. To elucidate the mechanisms of these functions in detail, submolecular-scale changes at the surface of tubulins caused by chemical modification need to be investigated. The results obtained in this study demonstrate that FM-AFM can be used to visualize subnanometer-scale structures of tubulins. This capability should be of considerable value in future studies on the molecular-scale mechanisms of tubulin functions.

Conclusions

In this work, we performed molecular- and submolecular-scale imaging of tubulins by FM-AFM in liquid. In the imaging of tubulin protofilaments, individual tubulins were clearly imaged by FM-AFM. This result shows the applicability of FM-AFM to molecular-resolution imaging of isolated tubulin protofilaments. In the imaging of tubulin sheet-like structures, the individual α-helices were imaged as a periodic contrast with a spacing corresponding to the pitch of an α-helix backbone. To our knowledge, this is the first demonstration of real-space imaging of individual α-helices in liquid. By determining the position of the α-helices, we were able to identify the orientation of the tubulins. As a result, we found that the C-terminal domains are imaged by FM-AFM as bright protrusions with a height of 0.4 nm. The results obtained in this study demonstrate that FM-AFM is capable of identifying arrangements of secondary structures such as α-helices and C-terminal domains at the tubulin surface. This capability should be of considerable value in future studies on the molecular-scale mechanisms of tubulin functions.