Yeast Nanobiotechnology

Abstract

Yeast nanobiotechnology is a recent field where nanotechniques are used to manipulate and analyse yeast cells and cell constituents at the nanoscale. The aim of this review is to give an overview and discuss nanobiotechnological analysis and manipulation techniques that have been particularly applied to yeast cells. These techniques have mostly been applied to the model yeasts Saccharomyces cerevisiae and Schizosaccaromyces pombe, and the pathogenic model yeast Candida albicans. Nanoscale imaging techniques, such as Atomic Force Microscopy (AFM), super-resolution fluorescence microscopy, and electron microscopy (scanning electron microscopy (SEM), transmission electron microscopy (TEM), including electron tomography) are reviewed and discussed. Other nano-analysis methods include single-molecule and single-cell force spectroscopy and the AFM-cantilever-based nanomotion analysis of living cells. Next, an overview is given on nano/microtechniques to pattern and manipulate yeast cells. Finally, direct contact cell manipulation methods, such as AFM-based single cell manipulation and micropipette manipulation of yeast cells, as well as non-contact cell manipulation techniques, such as optical, electrical, and magnetic cells manipulation methods are reviewed.

1. Introduction

Nanotechnology is the ability to work at the atomic, molecular, and supramolecular levels (on the scale of ~1–100 nm) to understand, create, and use material structures, devices, and systems with fundamentally new properties and functions resulting from their small structure [1]. Nanobiotechnology is defined as a field that applies nanoscale principles and techniques to understand and transform biosystems (living or nonliving) and that uses biological principles and materials to create new devices and systems integrated from the nanoscale [2]. The biological and physical sciences share a common interest in small structures (the definition of “small” depends on the application, but can range from 1 nm to 1 mm) [3]. A bacterial cell is approximately 1 μm, a yeast cell 5 μm, and a mammalian cell is 10 μm when rounded and 50 μm when fully spread in attached culture. A vigorous trade across the borders of these areas of science is developing around new materials and tools (largely from the physical sciences) and new phenomena (largely from the biological sciences). The physical sciences offer tools for the synthesis and fabrication of devices for measuring the characteristics of cells and sub-cellular components and of materials useful in cell and molecular biology. Biology offers a window into the most sophisticated collection of functional nanostructures that exist. Nanobiotechnology offers new solutions for the transformation of biosystems, and provides a broad technological platform for applications in several areas—including bioprocessing in industry, molecular medicine, investigating the health effects of nanostructures in the environment, improving food products (food conservation), and improving human performance [2].

This review discusses nanobiotechnological analysis and manipulation techniques that have been especially applied to yeast cells. Nanoscale imaging methods that allow imaging at nanometer resolution are reviewed: atomic force microscopy (AFM), super-resolution fluorescence microscopy, and electron microscopy (including scanning electron microscopy (SEM), transmission electron microscopy (TEM), and electron tomography). Force spectroscopy for the analysis of single biomolecule interactions or unfolding on live yeast cells, as well as single-cell force spectroscopy, and the recently developed AFM-based nanomotion analysis of cells is reviewed and discussed. Since single-cell analysis has increasingly been recognised as the key technology for the elucidation of cellular functions which are not accessible from bulk measurements of the population level, nano/micro single-yeast cell manipulation techniques are reviewed. Yeast cell patterning techniques, such as microcontact printing, mechanical cell patterning, and the use of robotic cell printing are discussed. Finally, direct-contact (such as AFM-based and micropipette-based) and non-contact (such as optical, electrical, and magnetic cell) yeast cell manipulation techniques are reviewed.

2. Yeast Nanobiotechnological Analyses

2.1. Nanoscale Imaging

We use microscopy in order to see objects in more detail. The best distance that one can resolve with optical instruments (disregarding all aberrations) is about 0.5 times the wavelength of light, or the order of 250 nm with visible radiation. High-resolution microscopy techniques that are used for nanoimaging and nanoscale characterisation have been developed in the last 20 years. They can be divided into three categories: optical microscopes, scanning probe microscopes (SPMs), and electron microscopes. Recently-developed microscopy-based technologies can also be used to control and manipulate objects at the nanoscale—i.e., single-cell as well as single-molecule manipulation and analysis.

2.1.1. Atomic Force Microscopy

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. Examples of AFM imaging of yeast cell surfaces.

Yeast Type	AFM Analysis	Objective	Refs
C. albicans	Imaging, cell surface elasticity	Effect of antifungal caspofungin	[16]
	Imaging, cell elasticity	Imaging mode evaluation	[17]
	Imaging, force spectroscopy using concanavalin A-functionalised tips	Mapping of adhesive properties	[18]
Candida parapsilosis	Imaging, adhesion force	Surface morphological characterisation	[19]
S. cerevisiae	Imaging	Immobilisation method	[8]
	Imaging	Immobilisation method	[9]
	Imaging, force spectroscopy using concanavalin A-functionalised tips	Mapping cell wall polysaccharides	[20]
	Imaging, cell elasticity	Mapping of cell elasticity	[21]
	Imaging	Cell surface change on thermal and osmotic stress	[22]
	Imaging, motion analysis	Nanomechanical motion analysis	[23]
	Imaging	Effect of electromagnetic field and antifungal nystatin on the cell wall	[24]
	Imaging, cell elasticity	Immobilisation method	[10]
	Imaging	Immobilisation method	[11]
	Imaging, cell surface elasticity	Effect of antifungal caspofungin	[16]
Sc. pombe	Imaging	Cell surface change on thermal and osmotic stress	[22]

). 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].

2.1.2. Light Microscopy

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. Super-resolution optical microscopy techniques (adapted from [37]).

Technique	Description	Spatial Resolution	Timescale
Fluorescence imaging with one-nanometer accuracy (FIONA)	Localises and tracks single-molecule emitters by finding the centre of their diffraction-limited point-spread function (PSF).	~1.5 nm	~0.3 ms
Single-molecule high-resolution colocalisation (SHREC)	Two-colour version of FIONA. Two fluorescent probes with different spectra are imaged separately and then localised and mapped onto the plane of the microscope.	<10 nm	~1 s per frame
Single-molecule high-resolution imaging with photobleaching (SHRImP)	Uses the strategy wherein, upon photobleaching of two or more closely-spaced identical fluorophores, their position is sequentially determined by FIONA, starting from the last bleached fluorophore.	~5 nm	~0.5 s per frame
Nanometer-localised multiple single-molecules (NALMS)	Uses a similar principle to single-molecule high-resolution imaging with photobleaching to measure distances between identical fluorescent probes that overlap within a diffraction-limited spot.	~8 nm	~1 s per frame
Photoactivatable localization microscopy (PALM)	Serially photoactivates and photodeactivates many sparse subsets of photoactivatable fluorophores to produce a sequence of images that are combined into a super-resolution composite.	~2 nm	~1 min
PALM with independently running acquisition (PALMIRA)	Records non-triggered spontaneous off–on–off cycles of photoswitchable fluorophores without synchronising the detector to reach faster acquisition.	~50 nm	~2.5 min
Single particle tracking PALM	Combines PALM with live-cell single fluorescent particle tracking.
Stimulated emission depletion (STED)	Reduces the excitation volume below that dictated by the diffraction limit by coaligning one beam of light capable of fluorophore excitation with another that induces de-excitation by stimulated emission.	~16 nm	~10 min
Stochastic optical reconstruction microscopy (STORM)	Small sub-populations of photoswitchable fluorophores are turned on and off using light of different colours, permitting the localisation of single molecules. Repeated activation cycles produce a composite image of the entire sample.	<20 nm	~mins

). 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].

2.1.3. Electron Microscopy

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).

2.2. Force Microscopy

AFM techniques have turned out to be a suitable and versatile tool for single-molecule interactions (

Table 3. Examples of yeast molecule interaction studies using single-molecule force spectroscopy (SMFS).

Cell Type	Interacting Molecule 1	Interacting Molecule 2	Rupture Force (pN)	Refs
C. albicans	Als5p	Fibronectin	2800 ± 600	[103]
S. cerevisiae	Prion protein Sup35 hexapeptide	Prion protein Sup35 hexapeptide	—	[104]
	Prion protein Sup35 hexapeptide antiparallel hairpin structure	Prion protein Sup35 hexapeptide antiparallel hairpin structure	32–134	[105]
	Nucleoporin	Nucleoporin	—	[106]
	Nucleoporin	Importin	—
	Flo1p	Flo1p	300 (100–600)	[107]

) and for probing the physical properties of microbial cell surfaces [102]. Especially, it has been used to study yeast surfaces: to determine nanomechanical properties of the cell wall, map cell wall proteins (Figure 2A), molecular recognition forces (receptor–ligand interaction), and characterise biomolecules by single-molecule unfolding (

Table 4. Examples of yeast receptor–ligand interaction studies using single-molecule force spectroscopy (SMFS) on cell surfaces.

Cell Type	Cell Receptor	Ligand	Rupture Force (pN)	Refs
C. albicans	Cell surface β-mannan	Anti-β-1,2-mannoside antibodies	41 ± 14	[108]
	Cell surface β-glucans	Anti-β-1,3-glucan antibodies	38 ± 10	[108]
	Cell wall chitin	WGA 1 lectin	65 ± 19	[108]
	Cell wall	Streptococcus mutans exoenzyme glycosyltransferase B	1000–2000	[109]
C. glabrata	Epa6p	Hydrophobic surface	-	[110]
	Cell surface β-mannan	Anti-β-1,2-mannoside antibodies	54 ± 9	[108]
	Cell surface β-glucans	Anti-β-1,3-glucan antibodies	41 ± 8	[108]
	Cell wall chitin	WGA 1 lectin	41 ± 8	[108]
S. cerevisiae	Cell surface α-mannan	Con A 2 lectin	75-200	[111]
	Cell surface α-mannan	Con A 2 lectin	92 ± 35	[108]
	Cell surface β-glucans	Anti-β-1,3-glucan antibodies	42 ± 7	[108]
	Cell wall chitin	WGA 1 lectin	54 ± 19	[108]
	Wsc1p-His-tagged	NTA-Ni2+	-	[112,113,114]
	HA 3-tagged Ccw12p	Anti-HA antibody	69.3 ± 31.4	[115]
	Ste2p	α-factor	250	[116]
S. pastorianus	Flo protein	Glucose	121 ± 53	[111]
	Flo protein	Con A 2	117 ± 41	[111]

¹ WGA: wheat germ agglutinin; ² Con A: concanavalin A; ³ HA: human influenza hemagglutinin.

). For these types of analyses, the force sensing capabilities of the AFM are used. AFM-based force spectroscopy exerts pulling forces on a single attached molecule by retraction of the tip in the z direction (perpendicular to the x–y scanning plane). Cantilever bending is detected by the deflection of a laser beam onto a position-sensitive detector, such as a quadrant photodiode. A piezoelectric actuator stage is used to control the positioning of the sample relative to the tip. AFM-based force spectroscopy is also used to study single cell interactions (cell–cell and cell–substrate adhesion).

Single-cell force spectroscopy (SCFS) assays on living cells have been applied to measure the strength of cell adhesion down to the contribution of single molecules [119,120,121] (Figure 2B,C). AFM-based SCFS is currently the most versatile method for the study of adhesive interactions of cells with other cells, proteins, and surfaces, since SCFS offers a large range of detectable forces (from 10 pN to 100 nN), and offers precise spatial (1 nm to 100 µm) and temporal (0.1 to >10 min) control over the adhesion experiment and experimental parameters [120]. A living cell can be attached to a tipless AFM cantilever and the interacting partner (molecule or cell) on a substrate-coated surface. Alternatively, the living cell can be fixed on a surface, and the tip functionalised with the interacting molecule. AFM force spectroscopy with a single cantilever-bound cell can be used to investigate cell–cell and cell–matrix interactions. The approach and withdrawal of this cell to and from its surface can be precisely controlled by parameters such as applied force, contact time, and pulling speed, benefiting from the AFM’s high-force sensitivity and spatial resolution. The data collected in these experiments include information on repulsive forces before contact, cell deformability, maximum unbinding forces, individual unbinding events, and the total work required to remove a cell from the surface (

Table 5. Examples of yeast using single-cell (SCFS) force spectroscopy studies.

Cell Type	Interaction Partner	Variables	Refs
C. albicans	Staphylococcus aureus	Deletion of ALS3 (adhesion gene)	[123]
	Hydrophobic DDP 1 coated surface	Surface hydrophilicity, hydrophobicity, deletion of HGC1, compared to S. cerevisiae	[124]
	C. albicans hyphae	Deletion of ALS3 and ALS1 (adhesion genes)	[125]
	DC-SIGN 2	Differences in the N-mannan structure of the cell wall	[118]
C. glabrata	Adhesin Epa6p	Surface hydrophilicity, hydrophobicity, expressed and deleted EPA6	[110]
S. cerevisiae	Abiotic surface	Surface hydrophilicity, hydrophobicity BSA coating, life cycle stage, glutaraldehyde-treated cells	[126]
	Silica surface	Different silica with defined roughness	[127]
	Methacrylate polymers surface	Polymer imprinted and non-imprinted surface	[128]
	Bare and polydopamine-coated glass	Polydopamine coating	[129]

¹ DDP: dodecyl phosphate; ² DC-SIGN: dendritic cell-specific intercellular cell adhesion molecule-3 (ICAM-3)-grabbing non-integrin.

, Figure 2B,C). Force spectroscopy can identify cell subpopulations and characterise the regulation of cell adhesion events with single-molecule resolution [122].

2.3. Nanomotion Analysis

New sensor technologies based on microcantilevers have recently been developed [130,131]. Nanomechanical oscillators are increasingly being used for the detection of very small masses [132] or for nanostress sensing in molecular biology [133,134]. Cantilever resonators have been shown to possess a mass resolution in the pico- to femtogram ranges in both air [135] and liquid [136,137]. Many of the available systems are limited by the need to perform the measurements in air or in a humid environment, and most rely on the detection of the replication of the cells on the surface of the sensor. Thanks to the many advantages they offer, microcantilevers have recently been explored as nanosensors for cell studies; they are highly sensitive, selective, label-free, real-time, and provide in situ detection capabilities [138]. Single cell detection and monitoring on the cantilever sensor has been reported for S. cerevisiae cells [139,140], E. coli and Bacillus subtilis [140,141], HeLa cells [142], mouse lymphoblasts [140], and human lung carcinoma and mouse lymphocytic leukemia cells [143]. Cell growth detection has been demonstrated by monitoring resonance frequency changes of cantilevers as the mass increases from immobilized S. cerevisiae and fungal Aspergillus niger spores on the surface of the cantilevers in humid air [144]. S. cerevisiae cells were deposited onto the cantilever surface, and its bending as a function of time corresponded to the yeast growth behaviour [138].

Recently, the metabolic state of living organisms that are immobilized on the cantilever surface could be detected by cantilever nanomotion analysis in physiological conditions [145,146,147]. In nanomotion analysis mode, the sample is directly deposited onto the cantilever, and the analysis is performed with the functionalised cantilever in liquid. This differs from nanomechanical resonators, where the liquid sample is flowed through a capillary in the cantilever (Figure 3A). If the sample is alive, its nanometric-scale motions are transmitted to the cantilever, causing it to oscillate. These oscillations are detected by monitoring the cantilever displacements with the traditional laser–photodiode system; a typical set-up is depicted in Figure 3Ba. The cantilever and the sample of interest are immersed in an analysis chamber equipped with an inlet and an outlet that permits measurement in liquids, and, importantly, the exchange of liquids during measurements. It has been observed that any type of organism induces oscillations of the cantilever that only last while the organism is alive [147]. Once an efficient killing agent is applied, the cantilever oscillations stop. The exact origin of these vibrations is still under investigation. In the case of motile organisms, such as mammalian cells or flagella-equipped bacteria such as E. coli, the answer is straightforward. However, in the case of immotile microorganisms such as yeast or Staphylococcus aureus, the explanation is more challenging. Probably, a direct momentum transfer between the sample’s surface proteins that undergo conformational changes and the cantilever plays an important role [146].

Figure 3Bb shows a typical nanomotion experiment with C. albicans. The AFM cantilever was pre-treated with glutaraldehyde and incubated in a solution containing the cells. Some Candida cells attached onto its surface. The cantilever was eventually inserted into the growth medium-filled analysis chamber, and its oscillations were recorded. After the injection of a buffer solution containing 10 µg/mL of caspofungin (an antifungal drug to which Candida is sensitive) in the analysis chamber, the cantilever oscillations dramatically decreased. This drop became noticeable after only 10 min post-caspofungin exposure. Such an application can be very efficient (in a timeframe of minutes) for the detection of chemicals to which living organisms are sensitive, or for simple assessment of the presence of living organisms in extreme environments.

3. Yeast Cell Patterning and Manipulation

3.1. Yeast Cell Patterning

Manipulating the physical location of cells is useful both to organize cells in vitro and to separate cells during screening and analysis [148,149]. The quest to manipulate cells on length scales commensurate with their size has led to a host of technologies exploiting chemical, mechanical, optical, electrical, and other phenomena. The major cell-patterning methods include patterning on adhesive micropatterns, mechanical cell patterning, and robotic cell patterning [150]. Cell-adherence methods have been especially developed for the adhesion of mammalian cells, but have also been developed for yeast cell patterning (

Table 6. Applications of yeast cell patterning.

Yeast Type	Cell Patterning Method	Issue Addressed	Refs
S. cerevisiae	Patterning on adhesive micropatterns	Microcontact printing of concanavalin A	[157]
	Mechanical cell patterning in microfluidic microchambers	Monitoring dynamics of single-cell gene expression	[158]
	Robotic cell printing	Systematic profiling of cellular phenotypes	[159]
	Mechanical cell patterning using trap barriers	Single cell gene expression analysis	[160]
	Robotic cell printing	Localisation of the yeast proteome during polarised growth	[161]
	Mechanical cell patterning in microfluidic microchambers	Quantitative analysis of the yeast pheromone signalling response	[162]
	Patterning on adhesive micropatterns	Microcontact printing of biotinylated bovine serum albumin	[163]
	Mechanical cell patterning using trap barriers	Whole lifespan microscopic observation	[164]
	Mechanical cell patterning in microfluidic single-cell microwells	Real-time cellular responses of the mating MAPK pathway	[165]
	Mechanical and chemical patterning in microchambers	Molecular phenotyping of aging in single cells	[166]
	Mechanical cell patterning using trap barriers	Single cell analysis of yeast replicative aging	[167]
	Mechanical cell patterning in elongated cavities	Monitoring the dynamics of cell division	[168]
	Mechanical cell patterning using trap barriers	Studying ageing and dynamic single-cell responses	[169]
	Patterning in microcavity array by negative pressure, and embedded in agarose gel layer	Long-term single cell growth observation	[170]
	Mechanical cell patterning using trap barriers	Automated measurements of single-cell aging	[171]
	Mechanical cell patterning using trap barriers	High-throughput analysis of yeast replicative aging	[172]
Sc. pombe	Mechanical patterning in culture microchambers	Mechanical mechanisms redirecting cell polarity and cell shape in fission yeast	[173]
	Mechanical patterning in single-cell microwells	Determination of the mechanical forces involved in cell growth	[174]
	Mechanical patterning in microchambers	Time-lapse fluorescence observation of the effect of a microtubule-inhibiting drug	[175]
	Mechanical trapping in single-cell cavities	Fission yeast synchronisation	[176]
	Mechanical barrier single-cell trapping	Lon-term observation using super-resolution fluorescence microscopy	[177]
	Mechanical patterning in chemostat microchambers	Long-term single-cell analysis	[178]
	Mechanical patterning in culture microchambers	Studies of cellular aging	[179]

MAPK: Mitogen-activated protein kinases.

). A variety of different patterning techniques have been developed to present adhesive ligands at a range of scales to investigate biological events, pushing the envelope on the minimum feature down to the nanometer scale [151,152,153,154,155,156].

Microcontact printing has become the most popular technique [180]. A polydimethylsiloxane (PDMS) stamp with desired microfeatures is fabricated using soft lithography methods, and is used to print adhesive biomolecules onto the culture substrate [157,181]. For yeast cell adhesion, the lectin concanavalin A (which binds to cell wall mannose and glucose aminoglycans) can be used as an adhesive molecule. S. cerevisiae was also immobilised on cholesterol-modified microcontact-printed spots [163]. Despite its popularity, microcontact printing has several drawbacks for cell biology labs, such as the requirement of a clean room to microfabricate the stamp, and variations in the quality of the protein transfer [182].

In mechanical cell patterning, mechanical barriers capture the cells at specified spots. Cells can be trapped in microchambers (Figure 4A), microwells (Figure 4C), or by cell trap barriers (Figure 4D) (Table 6). Various microfabrication techniques have been used to fabricate microwell substrates for cell cultivation [150]. The microwell can have a diameter from several hundred micrometers up to the dimensions of a single cell [174]. Single-cell microwell arrays allow large numbers of cells to be stimulated and analysed (usually by fluorescence microscopy) in a massively parallel fashion [165,183]. Single-cell analysis has increasingly been recognised as the key technology for the elucidation of cellular functions, which are not accessible from bulk measurements on the population level [184,185]. Yeast cells have been trapped in microfluidic microchambers by using inlet and outlet valves [186,162] (Figure 4A). Culture chambers that are open on both sides [173,179,187] or on one side [176] have been constructed. These chambers fit single-cell dimensions and confine the cells. These culture chambers are suitable for non-adherent cells, such as yeast and bacteria [188].

Mechanical cell trap barriers have also been used to capture cells from suspensions in fluidic devices [158,189,190]. Fluid flow pushes the cells into the traps, and, therefore, these cell traps are also designated as hydrodynamic cell traps [191]. Barriers have been designed with a small fluidic leak that allowed single-cell trapping [160,177,192] (Figure 4D).

To create cellular microarrays, cells can be spotted or “printed” using a fluid-dispensing device (“cell printer”) [150]. It is essential to obtain a highly reproducible number of living cells per spot and an optimised printing process that is qualified for the reproducible production of microarrays with cells that keep their vitality and function for analysis. Spot formation techniques are categorised as “contact printing” and “non-contact” printing [193,194]. Robotic yeast cell contact printing was initially used to print cells on an agar growth medium by using fluid-dispensing devices or pads [195], or cells were grown in multiwell culture plates and printed on a glass slide for high-throughput imaging [159] (Figure 3B), or only short-time analyses on living cells were performed. More often, non-contact-based devices are used to produce cellular arrays, such as modified inject printers or piezo-driven tips [196,197,198,199]. In non-contact printing techniques, the liquid metering is not determined by the complex interplay of the pin, the liquid, and substrate, but is separated from the substrate, because no contact between the printing tool and the substrate occurs. The fluid is ejected as a flying droplet or jet towards the surface from a certain distance, which makes metering more precise. One concept of non-contact printing is based on syringe–solenoid-driven printers, where a reservoir and a high-speed microsolenoid valve are connected to a high-resolution syringe (e.g., the M2-Automation, synQUAD, or Genomic Solutions system). Further non-contact microarrayers are piezoelectrically driven, where a technology similar to the one used in an ink-jet printer is used (e.g., M2-Automation, MicroDrop, PerkinElmer, Scienion, GeSim) [200,201]. A piezo-actuator is fixed at the top of the dispenser tip. The squeezing of the tip forced by the piezo-actuation induces droplet ejection out of the capillary. The fast response time of the piezoelectric crystal permits fast dispensing rates (kHz range), and the small deflection of the crystal generates droplets from tens of picoliters to a few nanoliters.

3.2. Direct Contact Cell Manipulation

3.2.1. AFM-Based Cell Manipulation

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].

3.2.2. Micropipette Manipulation of Single Yeast Cells

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].

3.3. Non-Contact Cell Manipulation

3.3.1. Optical Manipulation of Single Yeast Cells

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. Examples of yeast cell manipulation using optical manipulation.

Yeast Type	Issue Addressed	Refs
C. albicans	Control and manipulation of pathogenic yeast for live cell imaging and interaction with host cells	[237,238]
Hanseniaspora uvarum and S. cerevisiae	Confinement of an individual H. uvarum cell by S. cerevisiae cells increases the average generation time	[240]
S. bayanus	Study of growth pattern of cells under line optical tweezers generated by time-shared multiple optical traps	[241]
S. cerevisiae	On-chip single-cell separation and immobilisation using optical manipulation and thermosensitive hydrogel	[229]
	Real-time detection of hyperosmotic stress response in optically trapped single yeast cells using Raman microspectroscopy	[230]
	Optical manipulation of cells to microscopically observe environmentally-induced size modulations and spatial localisation of GFP-tagged proteins to elucidate various signalling pathways	[232]
	Optical trapping and surgery of living cells using two operational modes of a single laser	[242]
	Selection and positioning of single cells combined with microscopy analysis in a microfluidic channel; cycling of GFP-tagged Mig1p and Msn1p between the cytosol and nucleus	[239]
	Optical trapping and fluorescence microscopy investigation of the internal pH response and membrane integrity with increasing temperature	[232]
	Automated transportation of single cells	[243]
	Development of a microfluidic array cytometer based on refractive optical tweezers for parallel trapping, imaging, and sorting of individual cells	[236]
	Microfluidic sorting of arbitrary cells with dynamic optical tweezers	[235]
	Development of graded-index optical fibre tweezers with long manipulation length	[244]
	Position yeast cells in a microfluidic chamber to study glycolytic oscillations	[245,246]
	Tomographic phase microscopy with live cell rotation using holographic optical tweezers	[247]
	Development of a photonic crystal optical tweezer to trap an array of yeast cells	[248]
Sc. pombe	Displacement of the lipid granules	[249]
	Displacement of the nucleus	[250,251]
	Laser ablation of microtubules in vivo	[226]
	In vivo anomalous diffusion and weak ergodicity breaking of lipid granules	[252]
	Quantitative determination of optical trapping strength and viscoelastic moduli inside living cells	[253]

, 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.

3.3.2. Electrical and Magnetic Manipulation of Yeast Cells

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. Examples of yeast cell electrical and magnetic manipulation.

Yeast Type	Manipulation Method	Issue Addressed	Refs
S. cerevisiae	Electrophoresis and electroosmosis	Cell transport in microfluidic channels	[272]
	Dielectrophoresis	Live and dead cell separation	[278]
	Magnetic patterning	Demonstration of magnetic micromanipulation of magnetically labelled cells	[276]
	Electroosmosis	Cell transport via electromigration in polymer-based microfluidic devices	[288]
	Dielectrophoresis (AC DEP)	Sorting live and dead cells	[280]
	Dielectrophoresis (AC DEP)	DEP tweezer for single cell manipulation	[267]
	Dielectrophoresis	Multiple frequency DEP separation and trapping of live and dead cells	[281]
	Diamagnetic trapping	Cell magnetic trapping in an array using a CoPt micromagnet array	[289]
	Magnetophoresis	Contactless diamagnetic trapping of cells onto a micromaget array	[289]
	Electrophoresis	Electrophoretic cell manipulation in a microfluidic device	[290]
	Dielectrophoresis (AC DEP)	Separation of yeast cells from blood cells in a microfluidic chip	[268]
	Dielectrophoresis	Live and dead cell separation	[279]
	Dielectrophoresis	Microfluidic chip for guiding cells by AC electrothermal effect and capturing by nDEP trap	[277]
	Dielectrophoresis (DC DEP)	Separation of a mixture of S. cerevisiae and Escherichia coli cells	[291]
	Dielectrophoresis (AC/DC DEP)	Sorting live and dead cells	[282]
	Dielectrophoresis, electroosmosis, electrophoresis	High-throughput trapping of cells, separation of live and dead cells	[271]
	Dielectrophoresis	Cell manipulation and immobilisation using photo-crosslinkable resin inside microfluidic devices	[292]
	Dielectrophoresis	Controlled rotation and vibration of cell clusters	[286]
	Magnetic manipulation	Magnetic manipulation of Fe3O4-doped hydrogel-coated cells	[293]

). 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.

4. Conclusions

In recent years, single-molecule and single-cell analysis and manipulation techniques have been developed and applied to the study of yeast cells. Single-cell analysis has increasingly been recognised as the key technology for the elucidation of cellular functions, which are not accessible from bulk measurements at the population level. Various techniques are now available for the analysis of a single cell; with the aid of these techniques, many biological questions can be answered. A microfluidic device is now a suitable technique for single-cell analysis, because a microfluidic system can be manipulated with high throughput, and the amount of sample from a single cell is limited. As it became obvious from this review, the newly developed nanotechniques have been largely applied to the model yeasts S. cerevisiae and Sc. pombe for fundamental eukaryotic cell biology research, and the pathogenic model yeast C. albicans for elucidating the molecular basis of pathogen–host interactions.

High-resolution imaging techniques can provide up to single-biomolecule resolution. The most widely used imaging methods are scanning probe microscopy (i.e., AFM), super-resolution fluorescence microscopy, and electron microscopy. Their characteristics, advantages, and limitations are compared in

Table 9. Comparison of high-resolution techniques for imaging yeast cells (adapted from [303]).

Characteristic	AFM	Electron Microscopy (SEM, TEM)	Super-Resolution Fluorescence Microscopy (PALM, STORM, SIM)
Resolution	~10 nm 1	~1–10 nm	~5–50 nm
Live cell	Yes	No	Yes
Sample preparation requirement	Little	Little to substantial	Little to moderate
Sample preparation time	10 min–1 d	2 h–5 d	30 min
Image acquisition time	~5 min	5–10 min	Up to 24 h
Equipment cost	€150,000–350,000	€500,000	€250,000–500,000
Operational costs	Low	High	Moderate
Advantages	Localisation (and force spectroscopy) of single proteins; observation of dynamic processes; various environments (temperature, liquid, air, etc.)	Imaging of the cell ultrastructure at very high resolution	Time resolution.
Disadvantages	Only the cell surface is analysed; only one single cell at a time; slow temporal resolution; various sources of artifacts, such as cell or tip alteration	Fixation artifacts; no dynamics; no information on physical properties of proteins	Labelling is required

¹ Depends on the flatness of the surface; the provided value refers to the resolution for observing cells.

. As can be noticed, nanoscale imaging methods are complementary, and they are therefore combined in recently developed imaging platforms, such as bio-AFM and super-resolution fluorescence microscopy [298,299], or the integration of EM and super-resolution microscopy in correlative light and electron microscopy (CLEM) [300,301,302]. The nanoscale exploration of surfaces of microbes such as yeast cells using AFM has expanded rapidly in the past years. Using AFM topographic imaging, the surface structure of live cells under physiological conditions is achieved with unprecedented resolution. Real-time imaging allows dynamic events to be followed. Chemical force microscopy (CFM)—in which AFM tips are functionalized with specific functional groups—can be used to measure interaction forces on the surface of live yeast cells. Molecular recognition imaging using spatially resolved force spectroscopy, dynamic recognition imaging, or immunogold detection can be used to localize specific receptors, such as yeast adhesins. Quantitative analysis of cell–cell or cell–substrate interactions can be performed with a number of techniques, where AFM single-cell force microscopy, optical tweezers, magnetic tweezers, and micropipette manipulation are the most popular. Understanding the fundamental forces involved in the adhesion of yeast cells is important not only in microbiology, to elucidate cellular functions (such as ligand-binding or biofilm formation), but also in medicine (host-pathogen interactions) and biotechnology (cell aggregation). These force spectroscopy techniques are compared in

Table 10. Comparison of force spectroscopy techniques [295,304].

Characteristic	Optical Tweezers	Magnetic Tweezers	AFM	Micropipette
Type	Point	Global/point	Point	Point
	Non-contact	Non-contact	Contact	Contact
Spatial resolution (nm)	0.1–2	5–10	0.5–1	-
Temporal resolution (s)	10−4	10−1–10−2	10−3	-
Stiffness (pN nm−1)	0.005–1	10−3–10−6	10–105	0.01–1000
Force range (pN)	0.1–100	10−3–102	10–104	1–1000
Probe size (µm)	0.25–5	0.5–5	100–250
Energy dissipation	Yes	No	No	No
Surface considerations	No	No	Yes	Yes
Features	Low noise and drift dumbbell geometry; access inside a cell	Force clamp, bead rotation, specific interactions; access inside a cell	High-resolution imaging	Controlled deposition/transfer of selected cells
Limitations	Photodamage, sample heating, non specific	No manipulation (force hysteresis)	Large high-stiffness probe, large minimal force, non specific	Low throughput

. These techniques are complementary, since each technique is most suitable for a specific force range.

Several micro-nanomanipulation tools for cells have been developed. The methods can be based on direct-contact mechanical cell manipulation (such as AFM-based or micropipette-based manipulation), or based on non-contact cell manipulation (such as optical, electrical, and magnetic cell manipulation). Examples of these tools are the microchannel-embedded AFM microcantilevers that can be used to suck up one selected yeast cell, which can be further manipulated (positioned for patterning, pushed to another cell or substrate to perform SCFS), and robotic cell printing with picolitre volume dispensing. The manipulation of the physical location of cells is useful both to organise the cells in vitro for single-cell analysis and for specific cell–cell interaction analyses. Another recently developed tool is the use of the AFM cantilever as a very sensitive nanosensor that can detect the metabolic activity of living yeast cells, and even monitor protein conformational changes [305]. An optical tweezer can be used to manipulate several cells in 3D in a contactless way, and can also be applied as a micro-nanosurgery tool by using the nano-ablation option of the laser. Inside cell manipulation of structures has been demonstrated for optical and magnetic tweezers, and opens new possibilities for non-invasive cell organelle manipulation activities. Magnetic tweezers allow cell rotation, which can be important for cell surface location-dependent interactions (e.g., cell–cell interaction analysis during mating).

Electric and magnetic force can be used to trap and position cells at some physical location, to monitor cell viability and separate live from dead cells, transport cells in devices such as lab-on-a-chip to develop automated assays, and to characterise cell properties (e.g., by determining dielectrophoretic spectra of cells).