**Contents**



## **About the Editor**

**Ronnie G. Willaert**, Associate Professor, has extensive expertise in yeast research (*Saccharomyces cerevisiae, S. pastorianus, Candida albicans*, and *C. glabrata*) and single-molecule biophysics (high-resolution microscopy, i.e., confocal laser microscopy, AFM, force spectroscopy, and scanning probe lithography), yeast space biology research and hardware development, protein science (yeast adhesins), cell (yeast) immobilization biotechnology, fermentation technology, and brewing science and technology.

### *Editorial* **Yeast Biotechnology 3.0**

#### **Ronnie G. Willaert 1,2**


Received: 23 July 2020; Accepted: 27 July 2020; Published: 29 July 2020

**Keywords:** *Saccharomyces cerevisiae*; non-*Saccharomyces* yeasts; fermentation-derived products; fermented beverages; wine; beer; coffee bean fermentation; flavor; itaconic acid production; bioethanol production; bioreactors; yeast micro- and nanobiotechnology

#### **1. Yeast Biotechnology 3.0**

This Special Issue is a continuation of the first and second "Yeast Biotechnology" Special Issue series of the journal *Fermentation* (MDPI). This issue compiles the current state-of-the-art of research and technology in the area of "yeast biotechnology" and highlights prominent current research directions in the fields of yeast micro- and nanobiotechnology, brewer's yeasts and beer fermentation, wine yeasts and wine fermentation, coffee bean fermentation and new developments in biochemicals production by yeasts. We very much hope that you enjoy reading it and are looking forward to the next Special Issue "Yeast Biotechnology 4.0" to appear in 2020–2021 (https://www.mdpi.com/journal/fermentation/special\_issues/yeast\_4).

#### **2. Yeast Micro- and Nanobiotechnology**

Living cell microarrays in microfluidic chips allow the non-invasive multiplexed molecular analysis of single cells. Yvanoff et al. [1] developed a simple and affordable perfusion microfluidic chip containing a living yeast cell array composed of a population of green fluorescent protein (GFP)-tagged *Saccharomyces cerevisiae* clones. Mechanical patterning in microwells and robotic piezoelectric cell dispensing in the microwells were combined to construct the cell arrays. The developed microfluidic technology has the potential to be easily upscaled to a high-density cell array, allowing one to perform dynamic systems biology (proteomics and localisomics) experiments on growing cells.

Yeast resistance to antifungal drugs is a major public health issue. Fungal adhesion onto the host mucosal surface is still a partially unknown phenomenon that is modulated by several actors, among which fibronectin plays an important role. Targeting the yeast adhesion onto the mucosal surface could lead to a highly efficient treatment for *Candida* infections. A nanoscale approach to study the behavior of the pathogenic yeast *C. albicans* was develop by Kohler et al. [2]. Using atomic force microscopy (AFM)-based detection of the nanoscale motions of the yeast cells, it was demonstrated that strongly adhering strains reduce their nanomotion activity upon fibronectin exposure, whereas low adhering *C. albicans* remain unaffected. These results open novel avenues to explore cellular reactions upon exposure to stimulating agents and to monitor, in a rapid and simple manner, the adhesive properties of *C. albicans*.

#### **3. Brewer's Yeasts and Beer Fermentation**

Due to changing lifestyle trends and legislation, there is a growing demand for non-alcoholic beers (NABs). In recent years, production methods have been improved and the use of non-*Saccharomyces* yeasts has been investigated. Non-*Saccharomyces* yeasts are interesting, since fruity ester aromas can be introduced. Bellut et al. [3] evaluated several *Cyberlindnera* strains for NAB production. It was demonstrated that the selected *Cyberlindnera subsuciens* was suitable to produce a fruity NAB. The outcome strengthens the position of non-*Saccharomyces* yeasts as a serious and applicable alternative to established methods in NAB brewing.

For some years, there has also been a new trend in using non-conventional yeasts to change the aroma profile of traditional beers. Canonico et al. [4] proposed the use of *Torulaspora delbrueckii* to obtain a beer with a distinctive aromatic taste. *S. cerevisiae*/*T. delbrueckii* mixed fermentations resulted in beers with increased concentrations of some aromatic compounds such as ethyl hexanoate, α-terpineol, and β-phenyl ethanol and an emphasized note of fruity/citric and fruity/esters notes.

Brewer's yeast flocculation is a well-appreciated characteristic of industrial brewer's strains, since it allows the removal of the cells from the beer in a cost-efficient way. However, many industrial strains are non-flocculent and genetic interference to increase the flocculation characteristics is not appreciated by the consumers. Optimization of the brewer yeast towards a more flocculating phenotype can lead to a more efficient beer production and a higher final beer quality. An attractive approach to enhance the attributes of microorganisms is the adaptive laboratory evolution (ALE) approach. Kayacan et al. [5] applied ALE to non-flocculent industrial *S. cerevisiae* brewer's strains using small continuous bioreactors and obtained an aggregative "snowflake" phenotype. It was demonstrated that ALE increased the sedimentation behavior and that no major flavor changes in the produced beer was detected.

#### **4. Wine Yeasts and Wine Fermentation**

Before cryopreservation was an established method to store wine yeasts, strain collections were stored at room temperature on agar slants in glass reagent tubes covered with vaspar and sealed with cotton plugs. Matti et al. [6] characterized 60 strains from the old wine yeast collection from the Geisenheim Yeast Breading collection and confirmed the suitability of storing yeasts by this old method. White wine fermentations and post-fermentation aroma analyses were performed. It was shown that this old strain collection bears treasures for direct use either in wine fermentations or for incorporation in yeast breeding programs aimed at improving modern wine yeasts.

Yeasts naturally occur in vineyards on the grapes and consequently in wines. Kaˇcániová et al. [7] identified yeasts on 30 grape varieties and 60 wine samples. MALDI-TOF mass spectrometry was used for the identification of yeasts and a total of 1668 isolates were identified. The most isolated species from the grapes was *Hanseniaspora uvarum*, and from wine, it was *S. cerevisiae*.

The selection of the yeast(s) is one of the most important "tools" for modulating flavor and color in wines. Therefore, Vilela [8] reviewed the role of *Saccharomyces* and non-*Saccharomyces* yeasts, as well as lactic acid bacteria, on the perceived flavor and color of wines and the choice that winemakers can make by choosing to perform co-inoculation or sequential inoculation. This choice will help them to achieve the best performance in enhancing these wine sensory qualities, avoiding spoilage and the production of defective flavor or color compounds.

During the fermentation of wine, the malolactic fermentation (MLF), which is performed by lactic acid bacteria, takes a prominent role, since it influences the wine flavor and its microbiological stability. Izquierdo-Cañas et al. [9] studied the effects of simultaneous inoculation of a selected *S. cerevisiae* yeast strain with two different commercial strains of wine bacteria *Oenococcus oeni* at the beginning of the alcoholic fermentation on the kinetics of the MLF, wine chemical composition, and organoleptic characteristics in comparison with spontaneous MLF in Tempranillo grape must. It was shown that co-inoculation reduced the overall fermentation time by up to 2 weeks, resulting in a reduced volatile acidity. The fermentation-derived wine volatiles profile was distinct between the co-inoculated wines and spontaneous MLF and was influenced by the selected wine bacteria. Co-inoculation resulted in wines with very little lactic acid and buttery flavors.

Icewine is a sweet dessert wine that is fermented from the juice of naturally frozen grapes. The high concentration of sugars in Icewine juice results in considerable osmotic stress in the

fermenting *S. cerevisiae*. Yeast can combat this stress by increasing the internal concentration of glycerol by activating the high osmolarity of the glycerol response to synthesize glycerol and by actively transporting glycerol into the cell from the environment. Muyssen et al. [10] investigated the role of the glycerol/H<sup>+</sup> symporter Stl1p in Icewine fermentations. Therefore, a strain of the common Icewine yeast *S. cerevisiae* K1-V1116 that lacks *STL1* was constructed using a developed CRISPR-Cas9-based genome editing method. The results demonstrate that glycerol uptake by Stl1p has a significant role during osmotically challenging Icewine fermentations, despite potential glucose downregulation.

There is a high interest in monitoring the changes in biochemical compounds that are changed during the alcoholic wine fermentation, since the management of the alcoholic fermentation is crucial in shaping the wine quality. Berbegal et al. [11] demonstrated the use of proton-transfer reaction-mass spectrometry coupled to a time-of-flight mass analyzer (PTR-ToF-MS) to monitor on-line volatile organic compounds (VOCs). The effect of multiple combinations of two *Saccharomyces* strains and two non-*Saccharomyces* strains (*Metschnikowia pulcherrima* and *Torulaspora delbrueckii*) on the content of VOCs in wine was assessed.

#### **5. Co**ff**ee Bean Fermentation**

Yeast fermentation of coffee beans improves the functionality of the beans and the quality of the coffee. Haile and Kang [12] evaluated the effect of green coffee bean fermentation with *Wickerhamomyces anomalu*. They demonstrated an improved functionality of the coffee beans, which was reflected into an increased total phenol and total flavoid content and a reduced total tannin content, and an improvement of the 2,2-diphenyl-1-picrylhydrazyl radical scavenging assay and the ferric reducing antioxidant power. Significant differences were also found in the superoxide dismutase activity.

The quality of coffee can also be improved by fermenting the mucilage layer of the coffee mixture with lactic acid bacteria and yeasts. da Silva Vale et al. [13] studied the effect of co-inoculation of *Pichia fermentans* and *Pediococcus acidilactici* on metabolite production during fermentation and the volatile composition of the coffee beans. They demonstrated an improved fermentation efficiency and a positive influence on the chemical composition of the coffee beans

#### **6. New Developments in Biochemicals Production**

Over the last few years, intense research has been focused on the generation of alternative renewable biofuels by fermenting agriculture waste using yeasts. One of the trends is the production of bioethanol by the fermentation of cellulosic and hemicellulosic biomass. The fermentation and assimilation of xylose (the second most abundant hemicellulosic carbohydrate) is still a bottleneck in the efficient production of bioethanol, since the conventional yeast *S. cerevisiae* cannot consume xylose. However, non-conventional yeasts, such as *Spathaspora passalidarum* and *Pichia stipitis,* can utilize xylose. Selim et al. [14] reviewed recent advances in xylose metabolizing yeasts, with special emphasis on *S. passalidarum* for improving bioethanol production.

Itaconic acid is an interesting biochemical for the polymer industry, since it can be produced from renewable substrates (such as lignocellulosic based hydrolysates) by fermentation and can replace petrochemical-based chemicals. It can be produced by the filamentous fungus *Aspergillus terreus*. Recently, alternative itaconic acid-producing yeasts such as the basidiomycetous yeasts of the family *Ustilaginaceae*, have been studied. Krull et al. [15] evaluated *Ustilago rabenhorstiana* as an alternative natural itaconic acid producer. By the optimization of media components and process parameters, a final itaconic acid concentration of 50 g L-1 using fed-batch fermentation was obtained. Moreover, itaconic acid was produced from different sugar monomers based on renewable feedstocks and the robustness against weak acids as sugar degradation products was confirmed.

**Acknowledgments:** The Belgian Federal Science Policy Office (Belspo) and the European Space Agency (ESA) PRODEX program supported this work. The Research Council of the Vrije Universiteit Brussel (Belgium) and the University of Ghent (Belgium) are acknowledged to support the Alliance Research Group VUB-UGhent

NanoMicrobiology (NAMI), and the International Joint Research Group (IJRG) VUB-EPFL BioNanotechnology and NanoMedicine (NANO).

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


© 2020 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Article* **Robotic Cell Printing for Constructing Living Yeast Cell Microarrays in Microfluidic Chips**

**Charlotte Yvano**ff **1,2,\*, Stefania Torino 1,3 and Ronnie G. Willaert 1,2,4**


Received: 31 January 2020; Accepted: 11 February 2020; Published: 14 February 2020

**Abstract:** Living cell microarrays in microfluidic chips allow the non-invasive multiplexed molecular analysis of single cells. Here, we developed a simple and affordable perfusion microfluidic chip containing a living yeast cell array composed of a population of cell variants (green fluorescent protein (GFP)-tagged *Saccharomyces cerevisiae* clones). We combined mechanical patterning in 102 microwells and robotic piezoelectric cell dispensing in the microwells to construct the cell arrays. Robotic yeast cell dispensing of a yeast collection from a multiwell plate to the microfluidic chip microwells was optimized. The developed microfluidic chip and procedure were validated by observing the growth of GFP-tagged yeast clones that are linked to the cell cycle by time-lapse fluorescence microscopy over a few generations. The developed microfluidic technology has the potential to be easily upscaled to a high-density cell array allowing us to perform dynamic proteomics and localizomics experiments.

**Keywords:** cell printing; piezoelectric dispensing; *Saccharomyces cerevisiae*; GFP-tagged yeast clone collection; living cell microarrays; microfluidic chip; dynamic single-cell analysis

#### **1. Introduction**

Cell assays have been miniaturized by growing cells in multiwell plates with increasing well number from 96 to 384 and 1536 wells and decreasing well volume from 280 μL down to 3 μL. These experiments are typically integrated in a robotic analysis platform. Major drawbacks of robotic platforms are the expense of the instrumentation, the cost of experimental consumables, closed systems (batch growth), and still relative medium throughput compared to recently developed microfluidic chips. As cell collections grow, further miniaturization of cell assays is needed to increase parallelism of the analyses. Cell microarrays provide an attractive solution, as they could increase the throughput significantly [1–3]. A cellular microarray consists of a solid support wherein small volumes of different biomolecules and cells can be displayed in defined locations, allowing the multiplexed interrogation of living cells and the analysis of cellular responses [4,5]. Living cell microarrays have been combined with microfluidic bioreactors, which provide multiple advantages for multiplex dynamic analyses and high-throughput screening [4,6]. Cellular arrays are emerging as important tools for functional genomics, drug discovery, toxicology, and stem cell research [4,7,8]. A major advantage of cell microarrays over microtiter plates is the opportunity to measure parameters on hundreds of individual single cells and average them, instead of measuring the parameters of a whole cell population. An interesting application where living yeast cell arrays are used is "dynamic proteomics" in which living cell microarrays using a fluorescent protein (e.g. GFP)-tagged yeast clone collection and automated time-lapse microscopy is used for the rapid acquisition of *in vivo* quantitative data about the

dynamic proteome and protein subcellular localization ("localizomics"). This enables the identification of members of protein complexes and coregulated proteins, and the unravelling of signaling pathways such as exposure to stress compounds, antimicrobials, or mating conditions [9–13].

Mechanical cell patterning, where mechanical barriers capture the cells at specified spots, has been frequently used to create cellular microarrays [14] in microfluidic chips. Yeast cells have been mechanically patterned in (single-cell) microwells [15,16], microchambers [17,18], and mechanical trap barriers [19–21]. As for the creation of classical DNA microarrays, a fluid-dispensing device can be used to spot or "print" living cells in an array format [4]. Dispensing techniques are categorized as contact and non-contact dispensing [22,23]. Robotic contact printing (e.g. printing DNA microarrays) was initially used to print cells on a semi-solid agar growth medium [24]. Today, mostly non-contact-based devices are used to produce living cellular arrays [25–28]. Here, the fluid is ejected as a flying droplet or jet toward the surface from a short distance. 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 [29,30]. Typically, droplet volumes of 10 to 20 nanoliter are the lower dispensing limit. Another concept is piezoelectrical dispensing, where a technology similar to the one used in an ink-jet printer is used [31,32]. A piezo actuator is fixed around a glass capillary close to the end of the tip. The squeezing of the capillary 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. Robotic cell printing can be used to easily create a living cell microarray composed of a clone collection and at a positional xy accuracy of a few micrometer.

Here, we developed a perfusion microfluidic chip containing a living GFP-clone collection that can be used to perform dynamic proteomics and localizomics experiments and demonstrate its performance. We combined mechanical patterning and robotic cell printing to produce living cell arrays. Soft lithography was used to create a 102-microwell array in the epoxy resin SU-8 (Structured by UV-8) on top of a glass coverslip substrate. A commercial microchannel top plate was used to close the microfluidic chip. Piezo dispensing was optimized for the development of living yeast cell arrays in microfluidic chips. We demonstrated that a clone collection can successfully be printed into the microwell array and yeast cells can be grown in the microwells in continuous mode. Finally, time-lapse fluorescence imaging was performed using 6 selected GFP-tagged clones demonstrating that the approach is suitable to perform dynamic analyses of protein expression and protein localization in living cells.

#### **2. Materials and Methods**

#### *2.1. Yeast Strain and Media*

Clones from the *S. cerevisiae* Yeast GFP Clone Collection (ThermoFisher Scientific, Waltham, MA, USA) were used [33]. We selected 34 clones linked to the cell cycle (Table S1) and revived them from the cryo-stock on YPD-agar plates (yeast extract 10 g/L, peptone 20 g/L, dextrose 20 g/L, agar 15 g/L). Single colonies were then transferred to liquid cultures in Synthetic Complete (SC) medium supplemented with 2% (*m*/*v*) glucose. The yeast cells were grown overnight in a shaking incubator at 170 rpm and 30 ◦C. Prior to cell printing, the cultures were diluted in Phosphate Buffered Saline (PBS) to an optical density at 600 nm (OD600) of 0.5.

#### *2.2. Fabrication of the Microwell Substrate and the Microfluidic Chip*

The microfluidic chip consists of two parts: a bottom glass slide with SU-8 microwells and a top plate containing a microfluidic channel with in- and outlet ports (Figure 1). The microwell substrate was produced in-house whereas the top plate was obtained commercially (sticky-Slide I 0.4 Luer, Ibidi, Gräfelfing, Germany). Microwells were fabricated by SU-8 photolithography on glass coverslips. Glass coverslips (75 × 25 mm, thickness of 170 μm; Ibidi, Gräfelfing, Germany) were cleaned in acetone

(Carl Roth, Karlsruhe, Germany) and 2-propanol (Carl Roth, Karlsruhe, Germany) for 15 minutes each, then rinsed with ultrapure water and finally blow-dried. The glass slides were then exposed to air plasma at 100 W, 50 kHz, for 5 minutes (Plasma System Cute, Femto Science, Dongtangiheung-Ro, Korea). The negative photoresist SU-8 2050 (Kayaku Advanced Materials, Westborough, MA, USA) was spin-coated onto the glass slides at 3500 rpm for 30 seconds in order to reach an approximative thickness of 50 μm. Next, the SU-8 was soft-baked for six minutes at 95 ◦C on a hot plate. The SU-8 slides were then aligned with the photomask (film photomask, Selba, Versoix, Switzerland) in the mask aligner UV-KUB3 (Kloé, Saint-Mathieu-de-Tréviers, France) and illuminated with 365 nm ultraviolet (UV) light (intensity of 35 mW/cm2) for 10 seconds. Following UV exposure, the SU-8 slides were post-baked for six minutes at 95 ◦C on a hot plate. Finally, the slides were treated with SU-8 developer for seven minutes and next washed with 2-propanol and blow-dried. The dimensions of the wells were then evaluated by optical microscopy (Nikon Eclipse Ti2, Nikon, Tokyo, Japan) with a 10x objective (for the diameter and pitch distance) and with a 3D profilometer (Profilm 3D, Filmetrics, San Diego, CA, USA) for the well's depth. The microfluidic chip was assembled by pressing the sticky top plate (sticky-Slide I 0.4 Luer, Ibidi, Gräfelfing, Germany) to the microwell substrate.

**Figure 1.** Construction of the microfluidic chip containing the cell microarray. (**a**) The bottom substrate: the SU-8 microwell array on the glass coverslip. (**b**) The top plate containing the channel, inlet and outlet (Ibidi, Gräfelfing, Germany). (**c**) The integrated microfluidic chip obtained by sticking the top plate to the bottom plate using double-sided sticky tape. (**d**) The microwells constructed by SU-8 UV-photolithography.

#### *2.3. Cell Printing*

Cell printing was performed using a non-contact iTWO-400 dispenser (M2 Automation, Berlin, Germany) (Figure S2a). The printer is established in an environmental enclosure for live cell printing that contains a HEPA filter and recirculating air is sterilized with a UV lamp. The environmental temperature can be controlled as well as the relative humidity. Moreover, the instrument deck contains a cooling system for source plates and target plates with dew point control, which prevents evaporation during spotting. Samples to be printed are manually dispensed in 384-multiwell plates (ShallowWell 384-multiwell plate, Thermo Fischer Scientific, Waltham, MA, USA), which are then mounted onto the "source plate" locations of the robot (Figure S2a). Likewise, the substrate to be printed is mounted onto the "target plate" locations of the robot (Figure S2a). The actual printing is performed with a piezo dispenser made of a borosilicate glass capillary surrounded by a piezo ceramic actuator (PDMD, M2 automation, Berlin, Germany), which is able to shoot pico- to nanoliter droplets at high frequency with high volume and position accuracies. Finally, the instrument deck is also equipped with a wash station, which enables to clean the tip of the piezo dispenser after sample aspiration and sample printing, in order to avoid cross-contamination. A typical printing run consists in aspirating the sample to be printed from the "source plate", washing the outside of the tip to get rid of contaminants that

could impair the shooting, shooting the sample at its desired position on the "target plate" and finally dispensing the remaining sample at the washing station and washing the outside of the tip so that it is cleaned for the next sample. Each step of this procedure can be specified in the software controlling the robot. Additionally, the robot is equipped with a camera annexed to the piezo dispenser, which enables us to verify the successful printing of the samples.

Ultrapure water was printed with a pulse duration of 15 μs and an amplitude voltage of 75 V. The environmental temperature was around 28 ◦C, the relative humidity was controlled at 50% and the temperature of the target plate was at 15 ◦C. These parameters were maintained throughout each of the cell printing experiments. For optimization purposes, ultrapure water was printed on standard glass coverslips (24 × 24 × 0.17 mm) and in microwells of commercial nanotiter plates (Microfluidic ChipShop, Jena, Germany) made of the cyclo-olefin copolymer material Topas.

Prior to cell printing, the SU-8 microwells were coated with a solution of concanavalin A (Con A) (Con A from *Canavalia ensiformis*, Sigma Aldrich, Overijse, Belgium) at 2 mg/mL in H2O, with 5 mM CaCl2 and 5 mM MnSO4. Sixty drops of Con A were printed per well with the parameters mentioned above and were left to incubate for 15 minutes before being allowed to dry.

Yeast suspensions were prepared at an OD600 of 0.5 in PBS and 40 droplets were printed into each well with the parameters mentioned above. Multiple yeast suspensions were successively printed in different wells of the microfluidic chip. Therefore, the piezo dispenser was thoroughly washed between each sample to avoid cross-contamination. The washing procedure consisted in first discarding the old sample by ejecting 30 μL of liquid at a flow rate of 30 μL/s using the syringe pump and next, flushing the outside of the piezo dispenser with ultrapure water for five seconds at the wash station. The yeast cells were left to sediment for 10 minutes. The microfluidic chip was closed and connected to a syringe filled with SC medium via silicone tubing. SC medium was gently perfused into the channel of the microfluidic chip with a syringe pump (KD Scientific, Holliston, MA, USA) at a flow rate of 25 μL/min. Finally, the complete set-up (microfluidic chip and syringe pump) was installed on a microscope for direct imaging or kept at 4 ◦C overnight to image the next day.

#### *2.4. Microscopy*

The microfluidic chip and syringe pump set-up were installed on a Nikon Eclipse Ti2 epifluorescence microscope (Nikon, Tokyo, Japan) for time-lapse imaging. The microfluidic chip was inserted into a temperature-controlled chamber (Ibidi, Gräfelfing, Germany), which was mounted onto an automated scanning stage (ProScan III, Prior Scientific Instruments, Cambridge, United Kingdom). The syringe pump was placed next to the microscope and the syringe was covered with a syringe heater (New Era Pump Systems, Farmingdale, NY, USA). Both the temperature controller and the syringe heater were set at 30 ◦C for the duration of the time-lapse experiments.

For the growth experiment with 34 clones, bright field images of yeasts were acquired every 20 minutes, for 18 hours with a 20x objective. At the final time point (18 h), the yeasts were also imaged with a 60x objective in bright field and fluorescence. The GFP fluorescence was observed by exciting the sample with a LED light source (pE-300white, CoolLED, Andover, United Kingdom) and detecting it through a FITC filter. For the growth experiment with six clones, the yeasts were imaged in bright field and fluorescence and images were recorded every 30 minutes for three hours using a 60x objective.

#### *2.5. Image Processing*

Images acquired by the camera of the iTWO-400 dispenser were post-processed using Fiji [34]. More precisely, they were stitched together to form the pictures shown in Figure 2, Figure 3, and Figure 4 using the "Grid/Collection Stitching" plugin [35]. The bright field and GFP-fluorescent pictures were also processed with Fiji for background correction and manual stack alignment.

**Figure 2.** Optimization of the printing on a glass substrate. Water droplets were piezo dispensed (pulse duration of 15 μs and voltage of 75 V) as a 3 × 3 array on a glass coverslip: (**a**) 100 droplets at a frequency of 50 Hz with a pitch of 1000 μm; (**b**) 100 droplets at a frequency of 50 Hz with a pitch of 750 μm; (**c**) 100 droplets at a frequency of 50 Hz with a pitch of 500 μm; (**d**) 50 droplets at a frequency of 25 Hz with a pitch of 400 μm; (**e**) 25 droplets at a frequency of 25 Hz with a pitch of 300 μm; (**f**) 25 droplets at a frequency of 25 Hz with a pitch of 250 μm; (**g**) 10 droplets at a frequency of 5 Hz with a pitch of 250 μm; (**h**) five droplets at a frequency of 4 Hz with a pitch of 200 μm.

**Figure 3.** Optimization of piezo printing into microwells. (**a**) Commercial microtiter plate (Microfluidic ChipShop, Jena, Germany) containing three microwell arrays with square microwells of varying width; array A: width of 400 μm and pitch of 1125 μm, array B: width of 200 μm and pitch of 563 μm, array C: width of 100 μm and pitch of 281 μm. Water droplets were piezo dispensed (pulse duration of 15 μs, amplitude voltage of 75 V and frequency of 50 Hz) as (**b**)a2 × 2 array in array A wells at 200 droplets/well, (**c**)a4 × 4 array in array B wells at 25 droplets/well, (**d**)a8 × 8 array in array C wells at five droplets/well.

**Figure 4.** Con A and yeast cells dispensing into the microwells of the microfluidic ship (pulse duration of 15 μs, voltage of 75 V and frequency of 50 Hz): (**a**) 60 droplets of Con A were printed, (**b**) 40 droplets were printed from the yeast solution (OD600 of 0.5).

#### **3. Results**

#### *3.1. Construction of the Microfluidic Chip*

The microfluidic chip (Figure 1c) was made of two parts: the microwell array on a glass slide (Figure 1a) and the top plate with a microfluidic channel (Figure 1b). The microwell array was produced in the epoxy-based photoresist SU-8 through a standard photolithography protocol. The mask design of the microwell array consisted of three rows of 34 wells with a well diameter of 300 μm and a pitch distance of 850 μm (Figure S1). Visual inspection of the microwells after photolithography showed that the SU-8 wells matched the mask's dimensions (Figure 1d). Furthermore, the measured well's depth was approximatively 50 μm (Figure S1b), which fits the expected thickness of the SU-8 layer, according to the protocol mentioned earlier. Altogether, these results prove the photolithography process to be successful and a quick method to produce the microwells in the microfluidic chip.

The overall length and thickness of the microwell array were 28.4 mm and 2.0 mm, respectively. These dimensions fit within the microfluidic channel of the top plate that was used to seal the microfluidic chip and connect the inlet and outlet tubes. The top plate was commercially available and consisted of a channel of 50 × 5 × 0.4 mm (l × w × h) surrounded by a double-sided tape, which enabled us to quickly and easily stick it onto the bottom substrate. Once the microfluidic chip was sealed (after filling with the cells), it was connected to silicone tubing and a syringe filled with SC growth medium. The medium was then perfused at a very low flow rate (25 μL/min) with a syringe pump in order to ensure nutrient renewal for cell growth.

#### *3.2. Living-Cell Microarray Development*

First, piezo dispensing parameters were optimized by printing a water droplet array on a glass substrate. Piezo dispensing is accomplished by applying rectangular voltage pulses to the piezo ceramic actuator (Figure S2c). For the duration of each pulse, the actuator tube contracts a few micrometers, thereby initiating a pressure wave that causes the ejection of a droplet. The amplitude and the duration of the pulse influence the volume and the velocity of the droplet. The higher the amplitude (applied voltage) and the longer the pulse, the bigger the droplet and the higher its velocity. The frequency of the pulses is directly correlated to the duration of the pulse and also influences the droplet's volume. Finally, the viscosity of the sample to dispense also has an influence on the printing parameters. Therefore, piezo dispensing needs to be optimized for each solution that has to be printed. In this work, yeast suspensions were prepared in PBS prior to printing.

Considering that PBS solution and ultrapure water have similar viscosities; we first optimized the piezo dispensing parameters with ultrapure water for simplicity reason. Typically, we started by determining the amplitude and duration of the voltage pulses that were giving a reliable ejection of water droplets. We obtained a stable shooting of ultrapure water with an amplitude of 75 V and a pulse duration of 15 μs. These parameters resulted in a droplet volume ranging between 70 to 80 pL (Figure S2b). Next, we created 3 × 3 arrays of water droplets with decreasing pitch distance on a glass substrate (Figure 2). We started by shooting 100 drops/spot at 50 Hz with a pitch distance of 1000

μm, 750 μm and 500 μm. We could not print arrays with pitch distances lower than 500 μm without merging of neighboring spots. Hence, we reduced the droplet number and printed 50 drops/spot at 25 Hz with a decreasing pitch distance from 400 to 300 and 250 μm. Surprisingly, printing half the droplet number did not allow halving the pitch distance as expected. Additional testing established that five drops/spot at 4 Hz with a pitch distance of 200 μm were the limit parameters at which a 3 × 3 array of water droplets could be stably printed on a glass slide. This optimization process aimed at determining the minimal spot sizes and pitch distances that can be stably printed in order to increase the throughput of the platform.

Secondly, we evaluated the filling of microwells with water droplets using piezo dispensing. Therefore, we initially used a commercially available microtiter plate (size of a microscope slide) that contains squared microwells with dimensions of 400 × 400 μm (array A), 200 × 200 μm (array B), 100 × 100 μm (array C), and height of 20 μm (Figure 3). The pulse duration (15 μs) and amplitude voltage (75 V) were as for printing on the glass substrate. With a frequency of 50 Hz, the number of droplets decreased from 200 drops per well for array A to 25 drops per well for array B and five drops per well for array C.

Once the piezo-dispensing parameters were established with ultrapure water, the printing protocol was optimized for living cells. First, the microwells were filled with 60 droplets of Con A at a pulse duration of 15 μs, amplitude voltage of 75 V and frequency of 50 Hz (Figure 4a). Con A is a lectin, which binds to the mannose glycans at the yeast cell wall [36]. Hence, Con A was used as a coating, which anchored the yeast cells to the bottom of the microwells so that they were not flushed away when SC medium was perfused during continuous cultivation. Next, the density of printed yeast cells per well was optimized. The final objective of this study consisted in developing a method that allowed monitoring the growth of single yeast cells as well as their expression of GFP-tagged proteins by time-lapse fluorescence microscopy. Therefore, the printing process had to deliver only a few cells per well. To do so, two parameters were adapted: the cell density of the yeast suspensions to print and the number of droplets/well. The latter was evaluated first and 40 drops/well provided optimal filling of the microwell without overflowing (Figure 4b). Since this parameter was kept constant for all the living cell experiments, the cell density of the yeast suspensions was then optimized. Yeasts suspensions at OD600 of 0.8, 0.5, and 0.2 were tested (Figure S2d). The suspension at an OD600 of 0.8 proved difficult to print since this high cell concentration increased the viscosity of the sample, thereby preventing the droplet ejection. On the other hand, cell suspensions at an OD600 of 0.2 were possible to shoot but did not deliver yeast cells in every well. The suspension at an OD600 of 0.5 was then selected, as it could be reproducibly printed and guaranteed the presence of a significant number of single cells per well. All the yeast printing experiments were performed with the following optimized parameters: pulse duration of 15 μs, amplitude voltage of 75 V, and a frequency of 50 Hz.

Finally, since the objective was to print multiple clones from the GFP-tagged yeast collection in different wells of the microfluidic chip, a last printing procedure had to be optimized, namely the washing step between samples in order to avoid cross-contamination. The washing step includes the disposal of the printed sample followed by the cleaning of the outside of the piezo dispenser capillary. Discarding the sample can be done either by applying pressure to the liquid path of the robot or by ejecting it with a syringe pump. Indeed, the iTWO-400 robot is equipped with both a pressure unit and a syringe pump, which enable to flow liquid through the piezo dispenser on a passive or active basis, respectively. With the pressure unit, the liquid continuously flows at a fixed fluid flow rate for as long as the pressure is applied. With the syringe pump, a specified volume of liquid flows at a specified flow rate. Cleaning the outside of the piezo dispenser is performed at the washing station of the robot (Figure S2a), and only the washing time can be adapted. To evaluate the efficiency of the washing step on avoiding cross-contamination, wells from a 384-multiwell plate (MTP) were filled with both a yeast suspension in PBS (at OD600 of 0.5) and a colored ink according to a designed pattern (Figure 5a). The samples were then printed on solid agar medium in a petri dish and left to grow at 30 ◦C for 24 hours before evaluation. In the first experiment, the printed samples were discarded by

means of pressure (at 450 mbar) for 10 seconds, and the outside of the piezo dispenser was cleaned for five seconds. Although most of the design was correctly printed, one cross-contaminated spot was visible (Figure 5b). The experiment was then repeated; however, the samples were discarded in the wash station by means of the syringe pump. The cleaning of the piezo dispenser was kept at five seconds. Using this procedure, no cross-contamination was observed (Figure 5c).

**Figure 5.** Evaluation of the washing protocol to avoid cross-contamination. (**a**) Printing pattern to evaluate the washing protocol of the piezo dispenser. Transparent wells contain a yeast solution in PBS, and the colored wells contain a blue dye (no cells). (**b**) Washing protocol based on pressure resulted in 1 cross-contamination colony. (**c**) Washing protocol based on the syringe pump resulted in spotting without cross-contamination.

#### *3.3. Growth of Yeast Cells in the Microfluidic Chip*

A selection of 34 GFP-tagged *S. cerevisiae* clones that are related to the cell cycle were printed as triplicates into the well array (Figure 4, S2e), and the chip was closed by sticking the microfluidic channel to the microwell substrate (Figure 1). Time-lapse microscopy was used to follow the growth during 18 h. An overview of all wells is presented in Figure S3 and some selected wells in Figure 6. At the end of the growth experiment, the GFP-tagged proteins were visualized.


**Figure 6.** Yeast growth in some selected wells (see Figure S3 for the full overview).

To evaluate the suitability of the developed yeast chips to perform dynamic analyses of protein expression and protein localization in single yeast cells, a small set of 6 clones were selected, i.e. Cdc39, Cla4, Bem1, Shs1, Cdc14 and Cdc28. A time-lapse experiment was performed where these clones were observed at higher resolution (600× magnification) during 3 h (Figure 7).

**Figure 7.** Time-lapse fluorescence microscopy of selected GFP-tagged clones related to the cell cycle.

#### **4. Discussion**

To create a living GFP-tagged yeast array, we combined mechanical patterning by constructing an array of microwells with cell printing (robotic cell patterning), which allows the controlled placement of the GFP-tagged clones in the selected wells. The cells were trapped in the wells by sticking them to the glass bottom of the microwells using the lectin Con A. Microfluidic chips where a GFP-tagged yeast clone collection was patterned as an array into microchambers have been previously developed [10,37]. Also, a microfluidic perfusion system where a robotic printed yeast array on agar and sandwiched with a track-etched membrane has been described [38]. These designs and fabrication methods are much more complex and difficult to construct. Due to the open design, robotic patterning of living cells in microwells is much more flexible in creating different filling designs. Additionally, it could be used to create high-density arrays without increasing much the complexity of the microfluidic chip design and construction.

A hybrid SU-8 on glass microfluidic chip containing a living cell array was developed. The direct fabrication of SU-8 microwells on the glass coverslip substrate is a simple, low cost and a high precision method that is suitable to construct disposable biochips [39]. SU-8 is biocompatible and has also the advantage that high density well arrays at high aspect ratios could be created [40]. A microwell array of 3 × 34 microwells containing microwells with a diameter of 300 μm and depth of 50 μm were created. The SU-8 microwell layer stuck well to the glass substrate. The bonding of the SU-8 microwell array to the glass could also withstand temperature shifts to low temperature (refrigerator). This allowed to store the chip filled with yeast cells for a few days before the growth experiment was performed.

Piezo printing was selected as the method to dispense a Con A protein solution and the GFP-tagged yeast collection into the microwells. We performed these dispensing steps with a piezoelectric dispenser (also called drop-on-demand ink-jet printing) [41,42]. Piezo printing of proteins and cells has been used previously for various applications including enzyme printing for glucose biosensors [43], protein arrays [44], netrin-1 adhesive micropattern construction [45], bacterial *Escherichia coli* arrays [46,47], bacterial and yeast cells on cantilever array sensors [48], *S. cerevisiae* cells on an agar layer [38], and mammalian cells [49].

We demonstrated that printing a small array on a flat glass substrate is possible and this method could be used to create cell arrays on glass substrate in a cheap and easy way. However, the droplet size was not proportional to the droplet volume which is a limitation to increase the throughput. We expected that half the number of droplets would result in half the spot volume, half the spot size and that twice the number of spots could be printed in the array. However, this was not the case. Also, significant variations in the droplet locations occurred, which resulted in arrays that were not perfectly arranged, which can result in merging of the spots in case of a small pitch distance (Figure 2e,f).

Printing of cells into microwells compared to on a flat surface has many advantages. Spatial confinement in the microwell results in higher resolution printing: smaller well sizes than droplet spot sizes and smaller pitch distances can be obtained. Droplet position in the wells is more accurate than droplets on a flat surface (Figure 2). Additionally, the liquid–air interface is reduced for microwells compared to droplets on a surface, resulting in a reduced evaporation rate. All these benefits result in higher throughput and stable printing.

The printer setup allowed to aspirate a yeast solution from an MTP (multiwell plate) well and deposit picoliter droplets containing living cells at the target microwell, and to perform this consecutively for all filled MTP wells. A tip dispenser washing protocol based on syringe pump cell solution ejection was successful to avoid cross contamination between different wells since syringe pump ejection occurred at a much higher flow rate than pressure-based ejection and could remove all yeast cells from the piezo tip. A GFP-tagged clone collection in a 3 × 34 cell array was cultivated during 18 h (Figure 6 and S3). This experiment demonstrated that a clone collection of 34 clones in triplicate could be successfully printed at a cell concentration that allows single cell observation during several generations.

As a proof-of-concept experiment to observe changes in GFP-tagged protein expression and cellular location, we selected GFP-tagged proteins that play a role during the yeast growth cycle. The Cdc39 protein is a subunit of the CCR4-NOT1 core complex that has multiple roles in the regulation of mRNA levels [50,51]. A high fluorescence intensity covering the cytoplasm in the mother and daughter cell can be observed (Figure 7) since it is present at a large number of protein molecules per cell (4300) [52]. Cla4p is a Cdc42p-activated signal transducing kinase that is involved in septin ring assembly, vacuole inheritance, cytokinesis, and sterol uptake regulation [53]. It is distributed in the cytoplasm and the bud. Higher intensity spots can be observed at the site where the bud appears (Figure 7). Bem1p is involved in establishing cell polarity and morphogenesis and functions as a scaffold protein for complexes that include Cdc24p, Ste5p, Ste20p, and Rsr1p [54]. A high intensity spot can be observed at the bud site and the growing bud (Figure 7). Shs1p is a component of the septin ring that is required for cytokinesis [55]. Initially, it is present at the cell periphery and moves to the bud site to create the septin ring (Figure 7). Cdc14p is a phosphatase required for the mitotic exit [56]. It is present in the nucleus and the nucleolus (Figure 7). Cdc28p is a cyclin-dependent kinase (CDK) catalytic subunit and master regulator of mitotic and meiotic cell cycles [57,58]. It alternately associates with G1, S, and G2/M phase cyclins. It is observed initially in the cytoplasm and next in the nucleus (Figure 7). These results demonstrate that the microfluidic chip can be used to perform dynamic experiments with subcellular resolution of fluorescently-tagged proteins.

In the future, the number of microwells in the array could be upscaled allowing to perform dynamic proteomics and localizomics experiments. This cell microarray-based systems-biology platform could be used to detect directly chemical disturbances in a small-molecule compound screen of the proteome in contrast to genetic approaches that are based on chemical-genetic interactions and necessitates a multistep indirect approach [59]. The genome-wide tagging of proteins of *S. cerevisiae*, including with GFP, has already provided a vast resource of such information [9,11,33,60]. Analysis of this high-resolution, high coverage localisation data set in the context of transcriptional, genetic, and protein-protein interaction (PPI) data revealed the combinatorial logic of transcriptional co-regulation and spatial-temporal regulation of proteins, and provided for example a comprehensive view of trafficking and signalling regulatory interactions within and between organelles in eukaryotic cells. As demonstrated here, dynamic movements from one location to another can also be followed as the cell proceeds through the cell cycle. Dynamic movements of proteins have also been described in yeast cells that respond to environmental stresses such as dithiothreitol (DTT) stress [9], hydrogen peroxide stress [9], osmotic stress by potassium chloride [37], and nitrogen starvation [9] or chemical perturbations by rapamycin [11], hydroxyurea [11], or methyl methane sulfonate [10].

#### **5. Conclusions**

We developed a perfusion microfluidic chip containing living yeast cell arrays that allows long term cultivation of the yeast cells and high-resolution time-lapse fluorescence microscopy. The creation of the cell array was based on mechanical patterning in SU-8 microwells and piezoelectric filling of the microwells. The microfluidic chip was closed by sticking a top plate that contained the microfluidic channel and inlet and outlet to the microwell substrate. The developed technology and method were validated by a growth experiment of a clone collection of 34 clones (in triplicate) that are linked to the cell cycle. Additionally, a set of six selected GFP-tagged clones were observed by time-lapse fluorescence microscopy at high resolution to observe single cell protein expression and subcellular location of the GFP-tagged proteins.

Future technological challenges lie in the further upscaling of the technology and procedures to construct genome/proteome-wide cell microarrays and analyzing cells dynamically on a whole proteome level. Dynamic proteomics profiling information based on chemical compound (such as e.g. drug compound) perturbation should allow to determine the target(s) and mechanism of action (MoA) [61]. For example, this technology could lead to the discovery of novel antifungal molecules, which are highly desired due to the limited number of available antifungal drugs and the fast emergence of multiresistant pathogens [62,63].

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2311-5637/6/1/26/s1, Figure S1: Construction of the SU-8 microwell array on the glass substrate, Figure S2: The robotic piezo dispenser, Figure S3: Growth of yeast cells in the microfluidic chip, Table S1: Selected *S. cerevisiae* GFP clones that were used in the growth experiment.

**Author Contributions:** Conceptualization, C.Y. and R.G.W.; methodology, C.Y., S.T., and R.G.W.; formal analysis, C.Y.; investigation, C.Y.; resources, R.G.W.; writing—original draft preparation, C.Y. and R.G.W.; writing—review and editing, C.Y., S.T., and R.G.W.; supervision, R.G.W.; project administration, R.G.W.; funding acquisition, C.Y. and R.G.W. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by The Belgian Federal Science Policy Office (Belspo) and the European Space Agency (ESA) PRODEX program grant number FluoCells and Yeast Bioreactor projects. C.Y. was funded by FWO for the funding of the SB PhD grant. The Research Council of the Vrije Universiteit Brussel (Belgium) and the University of Ghent (Belgium) are acknowledged to support the Alliance Research Group VUB-UGent NanoMicrobiology (NAMI), and the International Joint Research Group (IJRG) VUB-EPFL BioNanotechnology & NanoMedicine (NANO).

**Acknowledgments:** We acknowledge Rouslan Efremov for the use of lithography equipment and Gangadhar Eluru for allowing us to use his design of the microwell mask. The Hercules Foundation (FWO Flanders) is acknowledged for the funding of the Raith Voyager e-beam equipment (AUGE/13/19).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Article* **Yeast Nanometric Scale Oscillations Highlights Fibronectin Induced Changes in** *C. albicans*

#### **Anne-Céline Kohler 1,\*, Leonardo Venturelli 1, Abhilash Kannan 2, Dominique Sanglard 2, Giovanni Dietler 1,3, Ronnie Willaert 3,4,5 and Sandor Kasas 1,2,6**


Received: 18 December 2019; Accepted: 19 February 2020; Published: 21 February 2020

**Abstract:** Yeast resistance to antifungal drugs is a major public health issue. Fungal adhesion onto the host mucosal surface is still a partially unknown phenomenon that is modulated by several actors among which fibronectin plays an important role. Targeting the yeast adhesion onto the mucosal surface could lead to potentially highly efficient treatments. In this work, we explored the effect of fibronectin on the nanomotion pattern of different *Candida albicans* strains by atomic force microscopy (AFM)-based nanomotion detection and correlated the cellular oscillations to the yeast adhesion onto epithelial cells. Preliminary results demonstrate that strongly adhering strains reduce their nanomotion activity upon fibronectin exposure whereas low adhering Candida remain unaffected. These results open novel avenues to explore cellular reactions upon exposure to stimulating agents and possibly to monitor in a rapid and simple manner adhesive properties of *C. albicans.*

**Keywords:** *Candida albicans*; adhesion; fibronectin; nanomotion; atomic force microscope (AFM)

#### **1. Introduction**

Yeast biotechnology is a recent field where nanotechniques are used to manipulate and analyze yeast cells and cell constituents at the nanoscale [1]. Among the nanotechniques, AFM-related approaches played a major role in unveiling morphological, mechanical and biochemical properties of yeast [2–4]. Recently, our team demonstrated that living cells attached onto a soft cantilever induce nanometric scale oscillations (referred to as nanomotion) that stop as soon as the organism dies [5]. Commercially available atomic force microscopes (AFM) or dedicated devices easily detect these oscillations. Nanomotion detection has been applied to numerous biological samples such as proteins, single organelles, and a plethora of living cells such as prokaryotes (bacteria) and eukaryotes (fungal, vegetal and mammalian cells) [6]. The most straightforward application of the technique is the ultra-rapid antibiotic sensitivity test (AST). AST can be performed within an hour as compared to long-lasting traditional AST methods, which depend on the replication rate of the bacteria [7–9]. The test consists in attaching the organism of interest onto an AFM cantilever and monitoring its oscillations as a function of time upon addition of antibiotics in the analysis chamber. It is worth noting

that the nanometric scale oscillations do not only reflect the living or death state of the organisms but also its activity [5,10].

Fungal infections are a major public health issue nowadays; it is estimated that every year fungi infect about 1.2 billion people [11]. *C. albicans* is a common fungal pathogen that belongs to the human microbiome of healthy individuals [12]. This commensal relationship is a complex interplay of candidial and human factors. However, impairment of the host immunity or the normal host microbiota can lead to *C. albicans* infection (candidiasis) [13]. *C. albicans* is the predominant cause of virtually all types of candidiasis [14]. The first step of the infection is the adhesion of *C. albicans* onto the host. This step is an essential determinant of pathogenesis, as it allows *C. albicans* to attach to host cells and to form biofilms or to disseminate in the host blood vessels. The biofilm increases yeast cell resistance to antifungal therapeutics and protects it from the host immune system [15]. *C. albicans* has developed multiple ways to colonize and infect host cells and tissues. One such mechanism is the specific ligand–receptor interaction through a whole range of adhesins displayed on the yeast cell wall [16–18]. These cell wall proteins are capable of recognizing protein ligands [16], glycolipids [19–22] and carbohydrates [23–29] on the host cells. Fibronectin is an important protein ligand of the host extracellular matrix (ECM) that plays an essential role in *C. albicans* adhesion [30]. Furthermore, targeting fibronectin has shown to alter *C. albicans* biofilm formation [31]. Therefore, a better understanding of the yeast–fibronectin interaction could lead to novel therapeutic options to fight candidiasis.

In this work, we applied nanomotion analysis to monitor the oscillatory activity of *C. albicans* upon exposure to fibronectin. We used an AFM-based nanomotion detector to follow the evolution of cellular oscillations in the absence and the presence of fibronectin on strongly and poorly adherent *C. albicans* cells. Interestingly, these two isolates reacted very differently to the interaction with fibronectin. These preliminary results demonstrate the potential of nanomotion analysis to monitor ligand–receptor interactions in a label free manner.

#### **2. Materials and Methods**

#### *2.1. Yeast Strains*

The *C. albicans* isolate 101 and CEC 3675 were kindly provided by Salomé Leibundgut and Christophe D'Enfert laboratories [32], respectively. The yeasts were cultured in yeast-extracted peptone-dextrose (YPD) medium (1% m/v yeast extract (Difco Laboratories, Fisher Scientific, Hampton, NH, USA), 2% m/v peptone (Difco Laboratories, Fisher Scientific, Hampton, NH, USA) and 2% m/v glucose (Sigma, St. Louis, MI, USA)) overnight at 30 ◦C with shaking (160 rpm).

#### *2.2. Experimental Procedures*

Rectangular tipless cantilevers (qp-CONT, NanoandMore GmbH, Wetzlar, Germany), with a nominal spring constant of 0.1 N/m and an average resonant peak in liquids of 8 kHz, were coated with 2 mg/mL of concanavalin A (Con A) (Sigma, St. Louis, MI, USA) for 30 min at room temperature. After removing the excess of Con A, the yeast cells were placed in contact with the cantilever for 1 h at room temperature to allow them to attach to its surface. Poorly attached *C. albicans* cells were removed by washing gently with YPD medium. Finally, the *C. albicans* covered cantilever was inserted into the analysis chamber containing 2 mL of filtered (0.2 μm syringe filter, Merck Millipore, Burlington, MA, USA) YPD medium. The measurements were performed at room temperature in YPD medium and in YPD medium containing 25 μg/mL of fibronectin (Sigma, USA). Fibronectin was directly added inside the chip reservoir. For the experiments performed with antifungals, caspofungin (Sigma, USA) was diluted in the YPD present in the analysis chamber to reach a final concentration of 100 μg/mL.

#### *2.3. Nanomotion Detector*

The cantilever oscillations were collected in real time using an in-house developed nanomotion detection device. The system relies on a laser-based signal transduction as typically used in commercial AFMs. A typical experiment lasted for 2 h. The control experiments were carried out for at least 4 h.

#### *2.4. Software and Nanomotion Analysis*

The cantilever oscillations were recorded and saved at 20 kHz using a USB-4431 DAQ card (National Instruments, Austin, TX, USA). The data acquisition program was developed in LabView. A dedicated Python program was used to process the recorded data and to display the deflection of the cantilever as a function of time. The software first removes the low frequency cantilever displacement signal by calculating a first order fit of the raw signal (deflection of the cantilever) by taking 20 seconds-long window frames. The obtained fit is then subtracted from the raw signal to remove thermally induced cantilever deflection. The thermal drift essentially occurs at the beginning of the experiment and during the fluid exchange procedures. The thermal drift free signal is further processed to obtain its variance in 10 seconds-window frames.

#### *2.5. Viability Assay*

Cells were placed inside a commercially available microfluidic chip (Ibidi, Planegg, Germany), and stained with calcofluor white (Sigma, USA), according to the manufacturer's instructions. To detect dead cells, propidium iodide (PI, Sigma, USA) was added to the YPD medium and the fluorescence of the yeast cells was recorded using an Axiovert microscope (Zeiss, Oberkochen, Germany).

#### *2.6. Adhesion Assay*

Adherence of *C. albicans* to TR146 cells was measured using the protocols previously described [33,34] with slight modifications (Figure S1). TR146 cells grown as monolayers in 6-well plates were incubated with 100 *C. albicans* cells for 20 min at 37 ◦C. The supernatant was carefully removed and spread on YPD agar plates to determine the number of non-adherent fungal cells. The adherent fungal cells that were left behind in the 6-well plates were rinsed with PBS and were overlaid with melted Wort agar at 40 ◦C. The plates were incubated at 30 ◦C for 36 h to count the colonies. Adherence was determined as the ratio of the number of colonies grown on Wort agar to the number of colonies grown on Wort agar and the number of colonies grown from the culture supernatant.

#### *2.7. Statistical Analysis*

Statistical analysis of nanomotion experiments were performed with the Python package Scipy. We performed the non-parametric Mann–Whitney U test for the three independent replicates. We used standard student t-test to process the adhesion assay on three independent replicates using the Graphpad Prism software.

#### **3. Results**

To assess a putative differential reaction of strongly and weakly interacting *C. albicans*to fibronectin, we quantified the adhesion of two different isolates, 101 and CEC3675, on oral keratinocytes (TR 146). As shown in Figure 1 isolate 101 was measured to have a significantly higher adhesion compared to isolate CEC3675.

To investigate the *C. albicans*–fibronectin interaction we used an in house nanomotion detector depicted in Figure 2A. The set up consists in an analysis chamber filled with liquid (in our case YPD) containing the cantilever to which yeast cells are attached (Figure 2B). The cantilever oscillations were recorded (Figure 2C) and processed to display the signal variance as a function of time (Figure 2D).

**Figure 1.** *C. albicans* isolates 101 and CEC3675 adhere differently to oral keratinocytes. Percentage of adherence of both isolates. Statistical analysis (*n* = 3) was done using standard t-test. The asterisk represents *p* < 0.05.

**Figure 2.** Nano-mechanical sensor system. (**A**) Representative image of a cantilever with attached *C albicans* cells. Scale bar 40 μm. (**B**) Schematic of the experimental system and data collection. (1) Liquids to be injected into the analysis chamber. In our case YPD, YPD containing fibronectin, and YPD containing caspofungin. (2) Analysis chamber with the AFM cantilever and *C. albicans* attached onto its surface (green circles). (3) Super luminescent diode. (4) Four-segment photodiode. (5) Optical microscopy with camera. (6) Liquid waste. (7) In-house dedicated electronics and National Instruments data acquisition card. (8) Desktop computer. (**C**). The collected raw data are processed; and (**D**). analyzed using the variance of the signal.

Using this system, we monitored the nanomotion pattern of *C. albicans* isolates 101 and CEC3675 in the absence and presence of fibronectin (Figure 3). Before addition of fibronectin, both isolates behaved similarly (Figure 3B). However, in the presence of fibronectin, nanomotion activity (variance) of isolate 101 drastically decreased (from 0.9 ± 0.5 to 0.3 ± 0.1) (Figure 3). In contrast, isolate CEC3675

did not present a significant decrease. To confirm that the drop of signal was not due to a change in the temperature, nor convective currents that can appear upon addition of a liquid in the analysis chamber, we performed control experiments, simultaneously, with another nanomotion detector. These experiments consisted in injecting the same quantity of medium, instead of fibronectin, into the analysis chamber. The obtained results showed no significant difference in the nanomotion pattern, for both isolates, upon addition of YPD media (Figure S2). Additionally, we assessed the number of cells present on the cantilever before and after the experiment to determine if the reduced signal was caused by cells being detached from the cantilever. The analysis of the images taken by the optical microscope located above the nanomotion detector (as depicted in the schematic in Figure 2A) confirmed that no cells detached from the cantilever throughout the experiments (Figure S3).

**Figure 3.** *C. albicans* isolate 101 and CEC 3675 react differently to fibronectin. (**A**). Representative graph of the normalized variance of isolate 101 in YPD (blue) and in YPD with fibronectin (orange). The decrease of the normalized variance is clearly visible between the two conditions. (**B**). The mean of the normalized variance (experiment in triplicate) represented as a bar plot for isolate 101 compared to isolate CEC 3675. Error bars are the confidence of intervals. Statistical analyses were done using Mann–Whitney U test, the asterisk represents *p* < 0.05.

To further exclude another cause of the decrease of the nanomotion signal for isolate 101, such as premature cell death, we monitored *C. albicans* viability by nanomotion and fluorescence microscopy in the absence and presence of fibronectin. Eventually the cells were killed by the antifungal caspofungin. As shown in Figure 3A, the variance of the nanomotion signal drastically dropped after the drug injection. The fluorescent viability test did not show any effect of fibronectin on the cellular viability as it can be noticed in Figure 4B. Similarly, fibronectin also did not have any effect on the viability of isolate CEC3675 (Figure S4).

**Figure 4.** Viability assay of *C. albicans*. (**A**). Nanomotion signal of *C. albicans* isolate 101 in the absence (blue curve) and presence of fibronectin (orange curve), and after killing the cells by the antifungal caspofungin (green curve). (**B**). Representative fluorescence images of *C. albicans* isolate 101 in the absence (left panel) and presence of fibronectin (middle panel), and after killing (right panel). Scale bar 5 μm.

#### **4. Discussion**

*C. albicans* infection is a multistep process, consisting in the binding of *C. albicans* on epithelial cells. In a first adhesion step, the *C. albicans* adhesins of the agglutinin-like sequence (Als) family bind to ECM proteins of the host such as fibronectin [35,36], laminin and collagen. The attachment of the yeast cell to the host is followed by the penetration and transmigration of hypha into host cells, which then leads to vascular dissemination as soon the hypha reaches blood vessels. In this study we only explored the interaction of fibronectin with the yeast form. Adhesins playing a role in the planktonic *C. albicans* adhesion are the Als family members Als1 [27] and Als5 [37], Eap1 [38–40], Csh1 (cell surface hydrophobicity) [41,42], Ihd1 [43,44] and members of the SAP family [45–47]. It has been shown that Als1, Als5, and Csh1 interacts with fibronectin; Sap9 and Sap10 can interact with the ECM proteins collagen and vimentin. It has not yet been demonstrated that fibronectin is a ligand for Sap9/10, Eap1 and Ihd1.

Here, we used nanomotion detection to monitor the oscillation pattern of planktonic *C. albicans* cells upon exposure to fibronectin. Two different clinical isolates that showed a different adhesive phenotype, were used. The isolate 101 adhered significantly stronger to the host epithelial cells compared to isolate CEC3675. Nanomotion experiments showed that fibronectin affects isolate 101 significantly more than CEC3675. This drop of the nanomotion signal indicates a modification of the cellular activity upon fibronectin—*C. albicans* interaction. These results suggest that the initiation of adhesion related signaling in the yeast cell upon fibronectin attachment is mediated by the interaction with adhesins. Potential adhesion candidates that have been shown to interact with fibronectin are

Als1, Als3 and Csh1. The complete elucidation of the molecular mechanisms involved in the process are still unclear and deserve further research. We plan to investigate which specific adhesin(s) is (are) involved in the observed activity reduction. Additionally, the effect of other ligands such as laminin, collagen IV, fibrinogen and gelatin [28,48–50] should also be investigated.

This work demonstrated the ability of nanomotion detection to monitor in real time and in a label-free manner cellular activity changes induced by interacting ligands. Activity changes induced by increasing glucose concentration were observed for *Escherichia coli* in a previous study [5]. This technique opens novel avenues to detect cellular activation or inhibition induced by ligand–receptor interactions.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2311-5637/6/1/28/s1, Figure S1: Schematic representation of the adhesion assay protocol, Figure S2: Effect of the injection of YPD medium in the analysis chamber, Figure S3: Density of yeast cells on the cantilever, Figure S4: Viability assay of isolate CEC3675.

**Author Contributions:** Conceptualization, A.-C.K.; methodology, A.-C.K. and L.V.; software, A.-C.K.; validation, A.-C.K., L.V. and A.K.; formal analysis A.-C.K., investigation, A.-C.K., L.V., A.K.; resources, S.K., writing—original draft preparation, S.K. and A.-C.K.; writing—review and editing all the authors; visualization, A.-C.K., A.K.; supervision, A.-C.K., S.K.; project administration, S.K., G.D.; funding acquisition, S.K., G.D., D.S., R.W. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung 200021-144321, CRSII5\_173863 and 407240\_167137, the Gebert Rüf Stiftung GRS-024/14, NASA NNH16ZDA001N-CLDTCH, EPFL and ESA PRODEX project Yeast Bioreactor.

**Acknowledgments:** The authors thanks C. d'Enfert for providing the *Candida albicans* strains and S. Leibundgut for highly constructive discussions. The technical assistance of Danielle Brandalise is acknowledged. The Belgian Federal Science Policy Office (Belspo) and the European Space Agency (ESA) PRODEX program supported this work. The Research Council of the Vrije Universiteit Brussel (Belgium) and the University of Ghent (Belgium) are acknowledged to support the Alliance Research Group VUB-UGent NanoMicrobiology (NAMI), and the International Joint Research Group (IJRG) VUB-EPFL BioNanotechnology & NanoMedicine (NANO).

**Conflicts of Interest:** The authors declare no conflict of interest.

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