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The AFM operates by the same principle as originally proposed for Binning, Quate and Gerber, similar to that of the scanning tunnelling microscope (STM) [61]. Compared to STM, it has the additional and important ability to image non-conducting materials. The AFM allows visualisation of how single biomolecules and biomolecular assemblies function in their native environment [62].
Figure 1.8: A schematic showing the principle of operation for an AFM or STM, where B is the path followed by the probe over the sample surface. A represents an atom adsorbed to the sample surface. Reproduced from [61].
The principle of AFM is simple to understand, as it is analogous to a blind person feeling the sample with a sharp stick: A sharp tip on a long flexible arm, known as the cantilever, is raster-scanned across a surface to determine its topography line by line [61]. The AFM monitors the bending of the cantilever through the deflection of a laser spot shining on the back of the cantilever, which is coated in gold or aluminium to enhance its reflectivity,. This is known as the optical-lever method [63]. In the simplest mode of AFM operation (contact mode), a piezo actuator moves the cantilever vertically from/to the sample to keep the cantilever bending, and therefore the applied force, constant. The topography of the surface is calculated from this piezo movement [64]. There are many modes of operating the AFM, several of which are discussed further in chapter 2.
1.4.2
Illustration of Atomic Force Microscopy Applied to Biomolec-
ular Processes
AFM is a tool to visualise and probe the nanoscale [64]. It has two major functions as a visualisation tool, first to determine the structure of biomolecules in their native state, i.e., in aqueous solution and unlabelled; and second to determine how biomolecules move and interact under physiologically relevant conditions [65]. AFM has become a well-established technique for imaging individual macromolecules at nanometre reso- lution [66], in particular for densely packed and stabilised molecules [67]. AFM has the ability to image a range of biomolecules and biomolecular assemblies from single molecules, such as nucleic acids [68]–[73] and proteins [5], [74]–[78], to whole cells [79]– [81]. AFM has contributed a great deal to biology over the last few decades, with in- creasing numbers of studies exploiting the AFM to elucidate biomolecular processes at the nanoscale, examining structure-function relationships [82]. This section provides a non-exhaustive list of applications as an illustration of AFM contributions to biology.
Figure 1.9: AFM images of DNA plasmids showing two different DNA conformations for the same plasmid. A) Right handed B-DNA, B) Left handed DNA. C) Line profiles taken along each of these images between the marked arrows showing variations in DNA structure. Adapted from [70].
DNA molecules for many years were imaged as long featureless polymer strands by AFM with little improvement in resolution [71], [83]. Over the past few years, sub- stantial improvements in the AFM have yielded increased sensitivity and resolution, allowing the apparatus to image secondary structure on single DNA molecules ph- ysisorbed to mica in solution [70]. Figure 1.9 shows variation in DNA confirmation as observed by AFM. Figure 1.9A shows the Watson-Crick right-handed B-DNA, with 1.9B showing a stretched and left-handed form that is probably induced or stabilised by interaction with the sample substrate. Figure 1.9C shows line profiles taken along each
of these images between the marked arrows showing variations in the helical repeat of the DNA.
AFM has been used to visualise the process of transcription by visualising the move- ment of RNA polymerase (RNAP) along a strand of DNA to gain mechanistic un- derstanding of transcription dynamics [84]. Nucleosome integrity and position after transcription was visualised. This is also relevant to other processes involving molecu- lar motors on DNA wrapped in nucleosomes, including DNA replication and chromatin remodeling.
Figure 1.10: Images of RNAP DNA complexes situated asymmetrically on a DNA template imaged by time lapse AFM. Collided complexes were formed by two different methods. In the one-step method collisions were caused immediately by the addition of nucleotide triphosphates (A and C). In the two-step approach a stalled intermediate (B) was formed by nucleotide omission before the missing NTP was added allowing transcription elongation to proceed causing a collision between the two RNAPs (C). Reproduced from [85].
Figure 1.10 shows how AFM has been used to obtain snapshots of of RNA polymerase (RNAP) and nucleosome collisions, including stalling and backtracking [86]. Collided complexes were formed by two different methods. In a one-step method, collisions were caused directly by the addition of nucleotide triphosphates (NTPs). Figure 1.10A shows RNAP bound to DNA prior to the addition of NTPs, and 1.10C the collided state after the addition of four NTPs. In a two-step method, a stalled intermediary step is formed by the omission of one of the key NTPs. Figure 1.10B shows the stalled intermediary NTP omission, and 1.10C the collided state where both RNAPs collide after the addition of the required final NTP. This resulted in the unexpected conclusion that collided RNAPs remain bound to the DNA strand.
Membrane proteins occur in the cell membrane and play key roles in cell-cell com- munication, transportation, and energy conversion [87]. Ion channels are membrane proteins that regulate the flow of ions across the cell membrane, and play a central role in cell regulation. Ion channels are often targeted for drug development [88]. The opening and closing of ion channels is regulated by ligands which react to the presence
of molecular species, e.g., ATP or calcium, in their local environment [89]. Information on these structures can aid in drug development, and provide valuable insight into how these channels function for signalling transduction. Ion channels, as is the case for all membrane proteins, are non-trivial to examine by other methods.
Figure 1.11: High-resolution AFM images of the ion channel MloK1. A) MloK1 im- aged in the presence of cAMP, whereby each subunit of the tetrameric channels is well resolved on each single molecule. MloK1 imaged without cAMP no submolecular structure is visible, corresponding to a closed channel. Inset (both): Averaged images of the channels. Scale bar 10 nm. Reproduced from [90].
Many ion channels have been studied extensively by AFM. Such studies include the nucleotide-modulated potassium channe MloK1 which opens in the presence of the nucleotide cAMP as shown in figure 1.11; the conformation changes of the outer mem- brane protein F (OmpF) and changes in electrostatic surface potential generated by the pore opening [91]; the opening of the ATP-gated P2X4 receptor by ATP [92]; the ar- rangement of rhodopsin in membranes extracted from bovine eye [93]; the organisation of voltage dependant ion channels in tetramers and hexamers, in native membranes [94]; and the closing of the KirBac3.1 channel in the presence of magnesium ions [89]. More recently high-speed AFM has been utilised to show the association and disso- ciation of bacteriorhodopsin trimers, shown in figure 1.12, suggesting that these form trimers before assembling into the lattice [95], [96]. AFM has also been used to show conformational changes in bacteriorhodopsin upon illumination with green light. This generates long range effects across the membrane whereby a cytoplasmic portion of each bacteriorhodopsin monomer undergoes a conformational change to alter its posi- tion and come into contact with adjacent trimers [60], [97].
Myosin V, a processive motor which acts as a cargo transporter in cells, has been extensively studied by a number of techniques [98]. Most recently Myosin V has been visualised at high resolution by AFM stepping along actin filaments as shown in figure 1.13 [99]. This study was carried out at improved resolution over previous studies [100]. It was shown that both the generation of tension in the stepping leg and the
Figure 1.12: HS-AFM images of the bacteriorhodopsin cytoplasmic surface showing association and dissociation of a bR trimer as highlighted by the red and white triangles respectively. Adapted from [95].
powerstroke can be generated without ATP, a mechanism which had not previously been postulated or observed.
Force spectroscopy, or force pulling, is a much used AFM technique for determining the structure of proteins, based on measurement of mechanical properties rather than of surface topography. E.g., unfolding experiments on the large muscle protein titin by force spectroscopy have contributed to wour understanding of single-biomolecule mechanics [102]. In force-spectroscopy experiments, a protein is attached stably to the substrate, at one end, and to the AFM tip at the other. As the AFM probe moves up the protein is stretched. The force is monitored as the extension is increased, and as each domain of the protein is unfolded, the force versus extension shows a characteristic sawtooth pattern where each sawtooth corresponds to the unfolding of a different domain [103]. This method enables the determination of intra-molecular forces.
Figure 1.13: Successive HS-AFM images showing the movement of Myosin V along an actin filament in 1 µM ATP, scale bar 30 nm. The vertical dashed lines show the centres of mass of the motor domains, and the plus sign indicates the plus end of actin. Adapted from [101].
Force spectroscopy has also been extensively used to study the mechanical properties of DNA, which are relevant for DNA transcription, gene expression and regulation. It has been shown via force spectroscopy that the covalently closed back-bone of a single
DNA strand withstands forces of at least 800 pN and that this depends on applied force, ionic strength, temperature, and base pair sequence [104].