Scanning probe microscopes (SPMs) are a family of instruments that are used to measure surface properties, and include atomic force microscopes (AFMs) and scanning tunneling microscopes (STMs). The main feature that all SPMs have in common is that the measurements are performed with a sharp probe operating in the near field; that is, scanning over the surface while maintaining a very close spacing to the surface. The STM—invented in the early nineteen-eighties by Binnig and Rohrer [4]—was the first to produce real-space images of atomic arrangements on flat surfaces. The development of the STM arose from an interest in the study of the electrical properties of thin insulating layers. This led to an apparatus in which the probe–surface separation was monitored by measuring electron tunneling between a conducting surface and a conducting probe. A few years later, Binnig and colleagues [5] announced the birth of the second member of the SPM family, the atomic force microscope (also known as the scanning force microscope, SFM). Numerous variations of these techniques have been developed since.

AFM is extensively used for imaging surfaces ranging from micro- to nanometer scales, with the objective of visualising and characterising surface textures and shapes [6]. It has evolved into an imaging method that yields structural details of biological samples, such as proteins, nucleic acids, membranes, and cells in their native environment. AFM is a unique technique for providing sub-nanometer resolution at a reasonable signal-to-noise ratio under physiological conditions. It complements electron microscopy (EM) by allowing the visualisation of biological samples in buffers that preserve their native structure over extended time periods. Unlike EM, AFM yields 3D maps with an exceptionally good vertical resolution (less than a nanometer). Additionally, the measurement of mechanical forces at the molecular level provides detailed insights into the function and structure of biomolecular systems. Inter- and intramolecular interactions can be studied directly at the molecular level. Recently, improvements in the temporal resolution were made by the development of high-speed AFM [7]. This technique is capable of observing structure dynamics and dynamic processes at the sub-second to sub-100 ms temporal resolution and 2 nm lateral and 0.1 nm vertical resolution.

Since AFM imaging can be performed in physiological conditions, high resolution imaging of the yeast cell surface can be performed on living cells. Therefore, an appropriate cell immobilisation method has to be used that avoids cell detachment by the scanning probe. Several methods have been developed and used to perform high-resolution live yeast cell imaging and analysis (i.e., force spectroscopy). Yeast cells can be trapped in the pores of a filter membrane [8] (Figure 1A), in a hydrogel [9,10], or in microfabricated microwells [11,12]. Especially the model yeast Saccharomyces cerevisiae has been imaged at the nanoscale (Table 1). The cell walls of other yeasts have also been visualised: the model pathogenic yeast Candida albicans, and the other model yeast Schizosaccharomyces pombe (Table 1). Imaging can be easily combined with nanoindentation experiments to map the elasticity of the cell surface [13].

Since the earliest examination of cellular structures, observing cells using a light microscope has fascinated biologists. Being able to observe processes as they happen with the use of light microscopy adds a vital extra dimension to our understanding of cell behaviour and function [25]. Microscopy has evolved to provide not only quantitative images but also a significant capability to perturb structure–function relationships in cells. These advances have been especially useful in the study of a wide range of biological processes, including cell adhesion and migration [26].

Recent advances in fluorescence microscopy have allowed the imaging of structures at extremely high resolutions [27]. The past decade witnessed an explosion of fluorescence microscopy-based approaches to image protein dynamics and interactions [28]. For example, fluorescence recovery after photobleaching (FRAP) or photo-activation using photo-convertible fluorescent proteins to assay protein mobility and maturation in cells [29]; and Förster resonance energy transfer (FRET) to monitor physical intra- or intermolecular associations in space and time [30,31].

Despite the advantages of standard fluorescence microscopy, ultra-structural imaging is not possible, owing to a resolution limit set by the diffraction of light (Rayleigh criterion) [32]. Therefore, the maximal spatial resolution of standard optical microscopy is around 200 nm. This limit is one to two orders of magnitude above the typical molecular length scales in cells. Several approaches have been used to break this diffraction limit (Table 2). The diffraction limit can be overcome by exploiting the distribution of fluorescence intensity from a single molecule. When imaged, a fluorophore behaves as a point source with an Airy disc point spread function. The center of mass of the function (and therefore the position of the molecule) can be obtained by performing a least-squares fit of an appropriate function (such as a Gaussian distribution) to the measured fluorescence intensity profile of the spot [33]. With a sufficient number of photons, these methods can provide a localisation of 1–2 nm (15 to 70 nm on intact cells), allowing the measurement of distances on the scale of individual proteins. Single-molecule detection offers new possibilities for obtaining sub-diffraction-limit spatial resolution [34,35,36].

Photoactivatable or “optical highlighter” fluorescent proteins (FPs) have emerged as powerful new tools for cellular imaging [38,39,40,41,42,43,44,45,46,47]. The fluorescent properties of these proteins can be altered upon illumination at specific wavelengths. They either switch between a fluorescent and non-fluorescent state (photoswitching) [48,49,50,51,52,53,54], or they change their fluorescence emission from one wavelength to another (photoconversion) [39,44,55,56]. The controlled photoconversion/switching of these proteins provides unique opportunities to mark and track selected molecules in cells in space and time [42,48,50,57,58,59]. High-density mapping of single-molecule motions can be obtained using photoactivated localisation microscopy (PALM) [60,61,62].

Another promising application of photoswitchable proteins is their use in super-resolution microscopy. This technique relies on the stochastic photoactivation and localisation of single molecules, in which a fluorescence image is constructed from high-accuracy localisation of individual fluorescent molecules that are switched on and off optically [63,64,65,66,67,68]. Microscope techniques that are based on this principle are called RESOLFT (reversible saturable optical fluorescence transitions) microscopy. RESOLFT microscopy concepts are photoactivated localisation microscopy (PALM) [63,65], fluorescence photoactivation localisation microscopy (FPALM) [69], stochastic optical reconstruction microscopy (STORM) [64,69,70,71,72], and PALM with independently running acquisition (PALMIRA) [73,74] (Table 2). Image resolution well below the Abbe diffraction limit is achieved. Labelled proteins can be localised with a precision down to about 2–10 nm. Stunning images have been obtained based on photoactivatable FPs [63,75] (Figure 1D). Recently, super-resolution microscopy has also been extended to dual-colour imaging [76]. Another recently developed super-resolution technique is STED (stimulated emission depletion) [72,77]. In a STED microscope, the focal spot of excitation light is overlapped with a doughnut-shaped spot of light of lower photon energy, quenching excited molecules in the excitation spot periphery by stimulated emission. A resolution of 15 to 70 nm has been realised to map, for example, the nanoscale distribution of proteins inside cells [78], on the plasma membrane [79], and the movement of synaptic vesicles inside the axons of cultured cells [80].

Microscopes consist of an illumination source, a condenser lens to converge the beam on the sample, an objective lens to magnify the image, and a projector lens to project the image onto an image plane, which can then be photographed or stored. In electron microscopes, the wave nature of the electron is used to obtain an image. There are two important forms of electron microscopy: scanning electron microscopy (SEM) and transmission electron microscopy (TEM). Both use electrons as the source for sample illumination. The lenses used in electron microscopes are electromagnetic lenses. For high-resolution surface investigations, two commonly used techniques are scanning electron microscopy (SEM) (Figure 1C) and AFM. The operation of the SEM consists of applying a voltage between a conductive sample and filament, resulting in electron emission from the filament to the sample. This occurs in a vacuum environment. The electrons are guided to the sample by a series of electromagnetic lenses in the electron column. The resolution and depth of field of the image are determined by the beam current and the final spot size. The electrons interact with the sample within a few nanometers to several microns of the surface, depending on the beam parameters and sample type. Along with the secondary electron emission (which is used to form a morphological image of the surface in the SEM), several other signals are emitted as a result of the electron beam impinging on the surface. Each of these signals carries information about the sample that provides clues to its composition. Two of the most commonly used signals for investigating composition are X-rays and backscattered electrons. X-ray signals are commonly used to provide elemental analysis. The percentage of beam electrons that become backscattered electrons has been found to be dependent on the atomic number of the material, which makes it a useful signal for analysing the material composition.

Since electron microscopy is conducted in a vacuum environment, it is at a disadvantage for the study of hydrated samples. To image poorly-conductive surfaces without sample charging may require conductive coatings or staining (which may alter or obscure the features of interest), or it may require low-voltage operation or an environmental chamber, which may sacrifice resolution. Recently, an electron microscopy technique was described for imaging whole cells in liquid that offers nanometer spatial resolution and a high imaging speed using a scanning transmission electron microscope (STEM) [81,82]. The cells were placed in buffer solution in a microfluidic device with electron-transparent windows inside of the vacuum of the electron microscope.

In TEM, the transmitted electrons are used to create an image of the sample. Scattering occurs when the electron beam interacts with matter. Scattering can be elastic (no energy change) or inelastic (energy change). Elastic scattering can be coherent and incoherent (with and without phase relationship). TEMs with resolving powers in the vicinity of 1 Å are now common. A relatively recent electron microscopy technique that can be used to study cells at the nanoscale is electron tomography. Electron tomography (ET) is the most widely applicable method for obtaining three-dimensional information by electron microscopy [83,84,85]. A tomogram is a three-dimensional volume computed from a series of projection images that are recorded as the object in question is tilted at different orientations. ET has the potential to fill the gap between global cellular localisation and the detailed three-dimensional molecular structure, because it can reveal the localisation within the cellular context at true molecular resolution and the shapes and three-dimensional architecture of large molecular machines. It can also reveal the interaction of individual proteins and protein complexes with other cellular components, such as DNA and membranes. A recent development is cryo-electron tomography (cryo-ET), which allows the visualisation of cellular structures under close-to-life conditions [86,87,88] (see Figure 1B as an example). Rapid freezing followed by the investigation of the frozen-hydrated samples avoids artifacts notorious to chemical fixation and dehydration procedures. Furthermore, the biological material is observed directly, without heavy metal staining, avoiding problems in interpretation caused by unpredictable accumulation of staining material. Consequently, cryo-ET of whole cells has the advantage that the supramolecular architecture can be studied in unperturbed cellular environments.

The ultrastructure of yeast cells (the model yeasts S. cerevisiae and Sc. pombe) was first studied by TEM using thin sections in 1957 [14], and the freeze-etching replica method was introduced in 1969 to obtain the fine structure of yeast cells [89]. During the next 50 years, techniques for the analysis of the ultrastructure of yeasts advanced greatly [90]. Initially, yeast cells were fixed solely with potassium permanganate (KMnO₄), and not by the widely used osmium tetroxide (OsO₄), since the thick cell wall is a barrier for the penetration of OsO₄. Finer EM images were obtained by using a double fixation with glutaraldehyde (GA) and KMnO₄ [91]. Important landmark studies have used conventional chemical fixation using GA and OsO₄—after enzymatic removal of the cell wall—to describe the cellular features of S. cerevisiae and to compare ultrastructural defects that result from mutations in key genes [92,93,94,95,96]. Next, methods using cryo-immobilisation followed by freeze substitution have been developed to provide excellent preservation of intact yeast cells [97,98,99]. These approaches involve rapid freezing of the sample with subsequent substitution treatment to replace frozen water in the sample with an organic solvent and fixatives [100]. Currently, high pressure freezing followed by freeze substitution (HPF/FS) is the method of choice for preparing cells for ET. Yeast prepared with these methods are used in 3D electron tomography studies for which sampling of the cell is performed at unprecedented resolution [88,101] (Figure 1B).

The desire to actively deliver precise amounts of biomolecules through nanosized probes initiated the development of novel microfluidic probes. Microfabrication processes have been introduced for the production of AFM cantilevers with embedded microchannels [202,203,204,205]. Microchannel cantilevers were connected to a pressure controller for active liquid handling in fluidic force microscopy (FluidFM) [206,207]. The ability to apply a pressure allows for negative pressure experiments involving suction for applications such as cell adhesion, or positive pressure experiments resulting in cell deposition on a specified spot or in controlled dispensing for applications such as the accurate delivery of bioactive compounds to a single targeted cell in physiological medium or even cell injection. FluidFM was used for the spatial manipulation of single S. cerevisiae cells [208]. Therefore, the hollow cantilever was positioned over a yeast cell and approached in AFM contact mode. An underpressure of ~50 mbar was applied to suck the cell against the channel aperture. After displacement, the cell was deposited onto the substrate with an AFM approach in contact mode, and the cell was released by applying a short overpressure pulse while retracting the probe. The underpressure single-cell immobilisation of cells on the cantilever also allows accelerating the pace of SCFS, since the conventional cell trapping cantilever chemistry can be avoided [124]. Single-cell C. albicans adhesion forces to a hydrophobic (dodecyl phosphate coated) surface were compared to adhesion to a hydrophilic (hydroxyl-dodecyl phosphate coated) surface, the C. albicans mutant ∆hgc1 (which reduces the cell surface hydrophobicity), and to S. cerevisiae adhesion to the hydrophobic and hydrophilic substrate (Table 5). Force adherence measurements of S. cerevisiae cells on bare glass and polydopamine-coated glass substrates have been performed using a microfabricated hollow cantilever made entirely from SU-8 [129] (Table 5). Highly flexible SU-8 cantilevers with integrated microchannels have been fabricated for both additive and subtractive patterning of S. cerevisiae cells [209].

The oldest and most commonly used approach for single-cell manipulation uses glass capillary micropipettes [210]. A negative pressure applied to growth media-filled capillary immersed in a cell culture dish controls the aspiration of a desired cell. A positive pressure dispenses the cell. Motion stages with multiple degrees-of-freedom were used to manually manipulate the micropipette and accurately control its tip position to perform either micromanipulation or microinjection [211]. Micromanipulators enable the controlled separation of selected living cells from suspension and even allow for isolation of prokaryotic cells [212]. They can also be used in adhesion studies, such as the interaction of a single C. albicans cell that is sucked to a micropipette with a diameter that is smaller than the cell, with a salivary pellicle-coated bead that is manipulated with a second micropipette [213].

Single cell manipulation systems that are based on capillaries are commercially available; for example: TransferMan (Eppendorf, Hamburg, Germany), PicoPipet (Bulldog Bio, Portsmouth, NH, USA), Stoelting Micromanipulators (Wood Dale, IL, USA), and miBot™ manipulator (Imina Technologies, Lausanne, Switzerland). These manipulation systems are manual, although the miBot micromanipulator is a mobile micro-robot that moves directly over the surface of the microscope base, has a nm spatial resolution, and can be remotely controlled. Micropipette cell manipulation systems that allow automatic selection and placement of a single yeast cell using vision-based feedback control have been developed [211]. A robotic micromanipulation system based on a general-purpose micromanipulator and a traditional glass micropipette was developed for pick-and-place positioning of single cells [214]. By integrating computer vision and motion control algorithms, the system visually tracks a cell in real time and controls multiple positioning devices simultaneously to accurately pick up a single cell, transfer it to a desired substrate, and deposit it at a specified location. A computer-controlled micropipette installed on an inverted fluorescence microscope was used to automatically recognise by computer vision, and both fluorescently labelled and unlabelled live cells in a Petri dish were picked up [215]. A recent developed computer vision-based automated single-cell isolation system allowed the isolation of single live cells from a very dense culture without immobilising cells on a surface [216].

Microchanneled AFM micropipettes have also been developed and used for cell adhesion and spatial cell manipulation applications (see previous section “AFM-based cell manipulation”). These AFM micropipettes are also designated as versatile nanodispensing (NADIS) systems [217]. Compared to conventional glass pipettes, this tool is particularly suitable when using substances of high cost or limited amounts, because significantly less volume is required for an experiment [218,219]. Another advantage over glass pipettes is the precise control wielded in the manipulation of sensitive targets, due to concurrent measurements of cantilever deflections without significant target damage [206]. Targets—such as functionalised surfaces or surface immobilised cells—can be precisely and gently manipulated physically, biologically, and chemically [129,207,208,220].

In the last decade, optical manipulation has evolved from a field of interest for physicists to a versatile tool widely used within life sciences [221]. Optical trapping and manipulation is a spin-off from research where lasers were used to study the effect of linear and angular momentum of light on small neutral particles. Arthur Ashkin first demonstrated that radiation pressure from a focused laser beam significantly affected the dynamics of micrometer-sized transparent and neutral particles, and two basic light-pressure forces were discovered: a scattering force in the direction of the incident light beam, and a gradient force in the direction of the intensity gradient of the beam [222]. The scattering component of the force works as a photonic “fire hose” pushing the particle in the direction of light propagation. The gradient force can be explained by a dipole in an inhomogeneous electric field that experiences a force in the direction of the intensity field gradient of the laser beam [223]. Using these forces, small particles (such as cells) can be accelerated, decelerated, and trapped in three dimensions.

Optical tweezers use light to levitate a particle (cell) of distinct refractive index [224]. The trapped cell is suspended at the waist of the focused (typically infrared) laser beam. The displacement of the cell from the focal center results in a proportional restoring force, and can be measured by interferometry or back-focal plane detection. Optical tweezers use a high gradient of optical pressure to guide cells by focusing a laser beam through a high numerical aperture (N.A.) lens on the cells. High optical intensity of about 10¹⁰ mW/cm² may cause damage on the cells, and are not suitable for long-term cell manipulation [225]. Micro-meter-sized homogeneous particles (or cells) can be trapped with forces ranging from a few pN to several tens of pN, depending on the optical properties of the particles and of the medium [226]. It is possible to track the position of the trapped particle with sub-nanometer accuracy at high (several MHz) repetition rates [227]. Due to the rapid advances in laser technology, optical manipulation setups have been developed that have become relatively uncomplicated. Optical manipulation is easily integrable with various microscopy setups, including confocal, super-resolution, or multiphoton microscopes. It allows for high spatial and temporal resolution, and interaction forces can be minimised.

Optical tweezers have been used to manipulate yeast cells, such as in cell trapping, cell positioning, and cell sorting (Table 7, Figure 5). Optical trapping was used to isolate single yeast cells from a mixture of two strains that were distinguishable in fluorescence microscopy [228]. An optical tweezer was used for the rapid separation and immobilisation of a single yeast cell by concomitant laser manipulation and locally thermosensitive hydrogelation [229]. Optical tweezers can be used to trap single yeast cells for further analysis, such as Raman microspectroscopy [230], time-lapse fluorescence microscopy to determine single-cell internal pH [231] (Figure 5C), or to study single cell dynamics by monitoring GFP-tagged proteins [232] (Figure 5D). Yeast cells are conducive to direct optical tweezing and can be used for single-cell force studies [233]. The effect of various factors (such as the ionic strength and the nature of the counter-ion in the solutions) on the adhesion and detachment force of yeast cells on glass was assessed [234]. Compared to AFM, magnetic tweezers, and more conventional ways of studying cell adhesion (such as shear-flow cells), optical tweezers present several advantages: direct measurements in physiological conditions, clear criterion to evaluate the proportion of adhering cells, and ease of examining the heterogeneity of cell behaviours in the population. However, the optical tweezer method is limited to low adherence forces (~1 to 100 pN) owing to the low refractive index of cells, and is sensitive to the cell optical heterogeneity.

Optical gradient forces generated by fast steerable optical tweezers are highly effective for sorting small populations of cells in a lab-on-a-chip environment (Figure 5). Reliable sorting of yeast cells in a microfluidic chamber by both morphological criteria and by fluorescence emission was demonstrated [235]. More than 200 yeast cells could be contact-free immobilised into a high-density array of optical traps in a microfluidic chip [236] (Figure 5B). The cell array could be moved to specific locations on the chip, enabling the controlled exposure of cells to reagents and the analysis of the responses of individual cells in a highly parallel format using fluorescence microscopy. Additionally, single cells were sorted within the microfluidic device using an additional steerable optical trap. Optical tweezers were used to spatially and temporally control pathogenic C. albicans and Aspergillus fumigatus and place them in proximity to host cells, which were subsequently phagocytosed [237,238].

Optical tweezers can also be used to investigate the complex system of mechanical interactions taking place inside a living cell [226,249,250]. The viscoelastic properties of living Sc. pombe were investigated by studying the diffusion of lipid granules naturally occurring in the cytoplasm [251,252]. Optical manipulation techniques, such as optical tweezing, mechanical stress probing, or nano-ablation allow handling of probes and sub-cellular elements (such as organelles and individual molecules) with nanometric and millisecond resolution [254]. A near-infrared optical tweezer was used for yeast cell manipulation and micro-ablation [255]. This micro-nanosurgery system is based on a pulsed ultraviolet laser that induces plasma formation for intracellular surgery in live culture cells with submicron precision. Optical tweezers allow force probing of organelles and single molecules in vivo [256,257]. PicoNewton forces—such as those involved in cell motility or intracellular activity—can be measured with femtoNewton sensitivity, while controlling the biochemical environment. A method to perform a correct force calibration inside a living yeast cell (Sc. pombe) was developed [257]. This method takes the viscoelastic properties of the cytoplasm into account, and relies on a combination of active and passive recordings of the motion of the cytoplasmic object of interest. Absolute values for the in vivo viscoelastic moduli of the cytoplasm as well as the force constant describing the optical trap were determined.

Instead of using surface chemistry to prevent or allow cells to attach to certain regions, electromagnetic forces can be used to control cell positioning [149] and adhesion [258,259]. Electrical fields are very suitable for cell and bioparticle manipulation, with the advantages of strong controllability, easy operation, high efficiency, and minimal damage to targets [260]. Electrokinetic motion of cells refers to the migration of electrically charged or uncharged particles in a liquid medium or suspension in the presence of an electric field [261]. Electrical forces for manipulating cells at the microscale include electrophoresis and dielectrophoresis (DEP) [148,149]. Electrophoretic forces arise from the interaction of a cell’s charge and a uniform or non-uniform electric field, whereas dielectrophoresis refers to the motion of polarised (uncharged) particles in only a non-uniform electric field [262]. Based on the applied electric field, DEP can be broadly divided into AC (AC DEP, classical DEP), DC (DC DEP), insulator-based DEP (iDEP, DC-iDEP), combined AC/DC (AC-iDEP), and travelling wave DEP (twDEP). In AC DEP, an array of metal electrodes is embedded inside a microdevice (such as a microfluidic chip) to generate a spatially non-uniform electric field, and can be used to separate particles by changing the medium property and frequency of the applied electric field [263,264]. It can also eliminate any electrophoretic (EP) and electroosmotic (EO) effect [265]. In DC DEP, the spatially non-uniform electric field is created by specially-designed insulators, such as electrically non-conducting obstructions or hurdles in a microdevice, and electrodes that are positioned at the ends of the microfluidic channels [263]. The DC electric field results in an EO force and eliminates the need for an external pump, which is required in the case of AC DEP. The combination of AC/DC DEP can transport cells by using electrokinetic effects (such as electroosmosis), and AC DEP can be used to separate cells. In the travelling wave DEP, transport and separation of cells can be performed with the AC electric field [266]. In this case, the spatial nonuniformity of the phase of the electric field is used to transport the particle, and the nonuniformity in magnitude of the field is used to separate the particles.

DEP and electrophoretic forces have been used to create microsystems that separate cell mixtures into their component cell types or act as electrical “handles” to transport cells or place them at specific locations [267] (Figure 6C). DEP has also been applied for cell sorting [268], focussing, filtration [269], and assembly [262]. DEP has been used to characterise cells, for example, to monitor cell viability changes (including morphology and internal structure) and isolate viable cells with minimal or no damage [270,271]. Electrophoretic and/or electroosmotic pumping can also be used to control and drive cell transport in microfluidic chip channels [272]. DEP tweezers have been developed that allow the positioning of a single cell in three dimensions or transfer a single cell to any designated area [260]. A DEP tweezer consisting of a sharp-tip glass needle with a pair of electrodes and using a pDEP force could hold a single yeast cell at the end of the micromanipulator [267]. The design of the tweezers was not adequately optimised for one-by-one manipulation; therefore, a round-tip shape for the DEP-based tweezers that shifts the electric field to the centre of the tweezers’ tip due to the smooth geometry at the tip is most suitable for single-cell manipulation [273].

DEP traps for single-cell patterning in physiological solutions have been developed [274] (Table 8). DEP manipulation and trapping of yeast cells has been included in microdevices such as microfluidic chips [272,275,276,277]. Live and dead yeast cell separation was achieved with the “headlands and bays” electrodes [278]. Live yeast cells are attracted to the regions of the maximum field, while the dead ones are repelled to the regions of the minimum field, resulting in the separation of live and dead yeast cells. The separation of live and dead yeast cells by DEP could be enhanced by using the cross-linking agent glutaraldehyde (since glutaraldehyde selectively cross-links nonviable cells to a much greater extent than viable cells due to the higher cell wall permeability of nonviable cells) [279]. Live and dead S. cerevisiae cells were sorted by using AC DEP [280], multifrequency DEP [281], or AC/DC DEP using a quadrupole electrode array [282] (Figure 6A). Yeast cells could be pre-concentrated by trapping using DEP, and separated depending on their vitality by using hydrodynamic, DC electrophoretic, and DC electroosmotic forces [271]. Live and dead yeast cells were characterised based on dielectric properties [283,284]. By combining DEP and image processing, dielectrophoretic spectra of cells can be acquired [285]. From this, the membrane properties of the cells can be obtained. DEP was used to control the rotation and vibration of patterned yeast cell clusters [286] (Figure 6B). This strategy is based on the cellular spin resonance mechanism, but it utilises coating agents to create consistent rotation and vibration of individual cells.

Magnetophoresis was applied to pattern yeast cells using a micromagnetic array [289]. Therefore, the diamagnetic S. cerevisiae were placed in an aqueous solution enriched in paramagnetic ions, and micromagnets that produce high magnetic field gradients were used. S. cerevisiae were coated with a single-layer of Fe₃O₄ nanoparticle-doped alginate hydrogel, which allowed their manipulation by a magnetic field [293]. Magnetic and electric manipulation of single or multiple yeast cells in a microfluidic channel was demonstrated using a microelectromagnet matrix and a micropost matrix [276]. The yeast cells labelled with magnetic beads were trapped by the microelectromagnet matrix, whereas the unlabelled cells were trapped by micropost matrix-generating electrical fields. The setup is suitable for the efficient sorting of yeast cells in a microfluidic chip. Yeast cells were trapped in a three-dimensional magnetic trap in an aqueous solution of paramagnetic ions [294].

Magnetic tweezers are similar in concept to optical tweezers; a magnetic particle in an external magnetic field experiences a force proportional to the gradient of the square of the magnetic field [295]. High forces can be achieved with relatively small magnetic field strengths, provided a very steep field gradient can be generated. The fields generated by sharp electromagnetic tips [296] or small permanent magnets [297] have been used to apply forces in excess of 200 pN on micron-sized magnetic particles. Magnetic tweezers are capable of exerting forces in excess of one nN (electromagnetic tweezers), and can be used to manipulate—and importantly, rotate—magnetic particles ranging in size from 0.5 to 5 μm. Magnetic tweezers are unique in that they afford passive, infinite bandwidth, force clamping over large displacements.