**1. Introduction**

Pathogenic microorganisms such as bacteria and fungi are able to successfully attack and colonize the host organism only while living in colonies. Colonies ensure the maintenance of appropriate local environmental conditions, increase local concentrations of many released compounds, such as proteolytic enzymes, and enhance the resistance to the host immune cells [1]. The development of infection is associated with the creation of biofilms - microbial assemblies that cover significant areas of host tissue. A critical factor for the optimal existence of microorganisms within biofilms is proper communication between pathogenic cells. It ensures synchronization of morphological changes, growth, gene expression and secretion of many compounds. The cell-to-cell communication system, called quorum sensing (QS) involves numerous quorum sensing molecules (QSMs), whose presence and concentration in the biofilm regulate biofilm functioning [2]. The first fungal microorganism in which the QSM mechanisms were identified was *Candida albicans*—an opportunistic yeast-like pathogen that resides on the skin and mucous membranes, which, due to a wide range of virulence factors, is responsible for the development of serious and hardly curable infections—candidiases [3–6]. In the properly functioning *C. albicans* QS, the autoregulatory QSMs are involved, including the best known farnesol (FOH) [3], farnesoic acid (FA) [7], and tyrosol (TR) [8,9]. In addition, *C. albicans* secretes phenylethanol and tryptophol, however, whether these two aromatic alcohols can act as QSMs in this

fungus still remains to be established [5]. FOH is a sesquiterpene alcohol made up of three isoprene units that is secreted by *C. albicans* into the extracellular space, reaching a concentration of over 50 μM, but the local QSM concentrations may be significantly higher [10,11]. *C. albicans* ATCC 10,231 strain was reported to secrete FA instead of FOH [12]. The presence of QSMs regulates the expression of many genes, including those responsible for the production of yeast virulence factors [13,14]. FOH was shown to work in an autoregulatory fashion, inhibiting the transition of *C. albicans* from the yeast-like to filamentous forms [3], thus blocking the formation of biofilm [15,16]. Its action is contrary to the second QSM-TR, a tyrosine-related alcohol that stimulates hypha production during the early stages of biofilm development [8]. FOH also affects the expression of genes responsible for the protection of *Candida* spp. against oxidative stress [17,18]. Moreover, FOH is used by fungi in the coexistence with *Pseudomonas aeruginosa* where this QSM down-regulates quinolone production by bacteria, thus enabling the coexistence of these two species [19]. Upon contact with the host, FOH presents immunomodulatory properties [20], affecting the efficiency of macrophages by decreasing their phagocytic activity [21], with the stimulation of the inflammatory cytokine expression [22]. FOH is also involved in blocking of monocyte differentiation into immature dendritic cells (DCs) and modulation of the DC's ability to induce T cell proliferation and activation of neutrophils [23].

Neutrophils (PMNs) can identify *C. albicans* and respond very quickly to the appearance of fungal cells by phagocytosis or the release of structures called neutrophil extracellular traps (NETs) [24,25]. However, the mechanism used by neutrophils to select between these two processes is still unknown. NETs are built of DNA backbone decorated with granular proteins like elastase, cathepsin G, proteinase 3 and myeloperoxidase (MPO), responsible for effective killing of pathogens. The process of NET release is also a mechanism of cell death, which results in the rupture of the cell membrane and the release of cellular content into the extracellular space [26]. In contrast, phagocytosis uses membrane tubulovesicular extensions (cytonemes) to capture pathogens without the neutrophil's death [27]. These components of NETs allow to defend against the hyphal form of *C. albicans* cells, which due to their size, cannot be effectively phagocyted [28]. *C. albicans* cells are trapped within the NET structures and then killed by granular enzymes and reactive oxygen species [26,29]. Many studies have indicated that some of *C. albicans'* virulence factors can activate NET release, the process named netosis [28–30]. Among them are glucans and mannans—the components of the fungal cell wall, as well as secreted aspartyl proteases (Saps), all of which can stimulate the netosis. However, the studies showing that the number of NETs significantly increases upon contact with the filamentous form of the pathogen, indicated that the morphology of fungal cells and, consequently, their size can determine the type of neutrophil response [28,30].

Although the influence of FOH on neutrophils has been demonstrated [23] there is no information about the possible netosis activation by QSMs, especially QSMs released by *C. albicans*. Therefore, the aim of this study was to determine the potential of FOH, FA, and TR to trigger NET release and to find the netosis signaling pathway activated by QSMs, as well as to verify their chemotactic properties.

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

#### *2.1. Isolation of Neutrophils*

Human neutrophils were isolated from EDTA-treated whole blood delivered by the Regional Blood Donation Center, Kraków, Poland, obtained from healthy donors. The neutrophil-containing fraction was isolated by Pancoll gradient separation [30]. The fraction containing neutrophils and erythrocytes was mixed with a solution of polyvinyl alcohol (1%) and incubated for 20 min at room temperature. The upper layer was collected and centrifuged. Erythrocytes were removed by lysis in a hypotonic solution. Cells were washed and resuspended in phosphate buffered saline (PBS). The neutrophil purity was typically >95%, as assessed by forward-scatter and side-scatter flow cytometric analyses.

#### *2.2. Viability Assay*

#### 2.2.1. Caspase 3/7 Activity

The cell apoptosis was analyzed by measuring the activity of proapoptotic caspases 3/7. Neutrophils (2 <sup>×</sup> 105 cells/well), suspended in solutions of FOH (trans,trans−3,7, 11-Trimethyl−2,6,10-dodecatrien−1-ol; Sigma-Aldrich, St. Louis, MO, USA), FA (Echelon Biosciences Inc, Salt Lake City, UT, USA) or TR (Sigma-Aldrich, St. Louis, MO, USA) at variable concentrations, were placed in the wells of a 96-well white microplate and incubated for 1 h at 37 ◦C, at 5% CO2. Then, the cells were washed with PBS, and 100 μL of Caspase-Glo® 3/7 Reagent (Caspase-Glo® 3/7 Assay, Promega, Madison, WI, USA) was added to each well, the plate was gently mixed by shaking at 300 rpm for 30 s and the chemiluminescence was measured continuously for 2 h at 37 ◦C.

#### 2.2.2. Annexin V/Propidium Iodide Analysis (Flow Cytometry)

The disturbance of the cell membrane was monitored using Annexin V (AnV) and propidium iodide (PI). Neutrophils (1 <sup>×</sup> 106 cells/sample) were placed in an eppendorf tube, washed twice with PBS, and resuspended in solutions of FOH or FA at variable concentrations. Unstimulated cells served as a negative control, and phorbol 12-myristate 13-acetate (PMA)-treated neutrophils represented a positive control. Cells were incubated for 1 h at 37 ◦C, at 5% CO2, washed three times with PBS, and stained with propidium iodide and fluorescein isothiocyanate (FITC)-labeled Annexin V for 15 min, according to supplier's instruction (Dead Cell Apoptosis Kit with Annexin V-FITC and PI, Invitrogen, Carlsbad, CA, USA). Then, cells were analyzed with flow cytometry (LSR Fortressa, BD, San Jose, CA, USA).

#### *2.3. Analysis of ROS Production*

The production of reactive oxygen species (ROS) by neutrophils was analyzed using chemiluminescence measurements. Neutrophils (2 <sup>×</sup> 105 cells/well) were suspended in 160 <sup>μ</sup>l of Krebs-Ringer phosphate buffer containing freshly prepared luminol solution (10–6 M) and allowed to settle for 15 min at 37 ◦C, 5% CO2 in the wells of 96-well white microplate. Then, 10 μl of FOH, FA, or TR were added at variable concentrations. Untreated neutrophils were used as a negative control, and cells stimulated with 25 nM PMA as a positive control. The chemiluminescence of luminol was recorded for one hour with one-second integration time, using a BioTek Synergy H1 microplate reader.

#### *2.4. Analysis of NET Quantity and Image*

Neutrophils (2.2 <sup>×</sup> 105 cells/well) were seeded into the well of 96-wells black microplate in 150 <sup>μ</sup><sup>L</sup> of RPMI-1040 and allowed to settle for 15 min at 37 ◦C, 5% CO2. Then, neutrophils were stimulated with 150 μL of FOH, FA or TR at variable concentrations in RPMI-1040. Unstimulated cells and cells treated with 25 nM of PMA served as negative and positive controls, respectively. Plates were incubated at 37 ◦C, 5% CO2 for 3 h and then analyzed, as follows:

For the visualization of NETs: the samples were fixed with 3.8% paraformaldehyde for 15 min and washed with PBS. Sytox Green (Thermo Fisher, Waltham, MA, USA.) dye was added to each well at the final concentration of 1 μM to visualize extracellular DNA. Imaging was performed using a Nikon Eclipse Ti microscope (Nikon Instruments, Melville, NY, USA).

For extracellular DNA quantification: the cells were washed three times with PBS and then 50 μL micrococcal nuclease (MNase, 1 U/mL) was added to cleave and release small fragments of extracellular DNA, and the microplates were incubated at 37 ◦C for further 20 min. The enzymatic reaction was stopped by the addition of EDTA solution (100 μg/mL), and after cell centrifugation (350× *g*, 5 min) 50 μL of supernatant was transferred into 96-well black microplate containing Sytox Green at a final concentration of 1 μM. Fluorescence was measured using the Biotek Synergy H1 microplate reader at the excitation wavelength of 465 nm and the emission wavelength of 525 nm.

#### *2.5. Identification and Quantification of Myeloperoxidase*

Neutrophils (2.2 <sup>×</sup> 105 cells/well) were seeded into the well of 96-wells black microplate in 150 <sup>μ</sup><sup>L</sup> of RPMI-1040 and allowed to settle for 15 min at 37 ◦C, 5% CO2. Then, neutrophils were stimulated with 150 μL of FOH at a concentration of 100 μM and 200 μM.

For the visualization of MPO: the samples were fixed with 3.8% paraformaldehyde for 15 min and washed with PBS. 50 μL of 1:100 diluted primary mouse anti-MPO antibodies (Abcam, Cambridge, UK) was added to each well and incubated for a night at 4 ◦C. Then, cells were washed three times with PBS and incubated with 1:500 diluted secondary Alexa Fluor 555 anti-mouse antibodies (Abcam, Cambridge, UK) for 1 h at 37 ◦C. Cells were washed two times with PBS, and imaging was performed using a Nikon Eclipse Ti microscope.

For the quantification of MPO: the quantitative determination of MPO was performed using Human MPO ELISA Kit (Wuhan Fine Biotech Co., Ltd., Wuhan, China). The cells were washed three times with PBS and then 50 μL micrococcal nuclease (MNase, 1 U/mL) was added to release DNA-bounded proteins, and the microplates were incubated at 37 ◦C for further 20 min. After cell centrifugation (350× *g*, 5 min) 100 μL of supernatant was transferred into anti-MPO pre-coated wells of plate, and the manufacturer's instruction was followed.

#### *2.6. Recognition of Receptors Involved in Netosis*

Neutrophils were preincubated with specific antibodies or inhibitors prior to stimulation with *C. albicans* factors. Neutrophils (1 <sup>×</sup> <sup>10</sup>6) were preincubated for 30 min at 37 ◦C in RPMI-1640 medium with 1 μg/mL of blocking antibodies directed against TLR2, TLR4 (Invivogen, Toulouse, France), CD11a, CD11b, CD16, CD18 (BioLegend, San Diego, CA, USA) or isotype control antibody—IgG (Abcam, Cambridge, UK).

#### *2.7. Analysis of Protein Kinase C (PKC)*

The activity of PKC was monitored using PepTag® Non-Radioactive Protein Kinase Assays (Promega, Madison, WI, USA). Neutrophils (5 <sup>×</sup> 106 in 500 <sup>μ</sup>L of PBS per well) were stimulated with 100 μM and 200 μM FOH in 12-well microplate. Negative and positive controls were prepared as described above. After 1 h of incubation at 37 ◦C, at 5% CO2, cells were washed with PBS, resuspended in 500 μL of cold PKC extraction buffer, and homogenized in the cold. Lysates were centrifugated for 5 min at 4 ◦C, 14,000× *g* and supernatants were purified on diethylaminoethyl (DEAE) cellulose resin. To the reaction solution containing 5 μL of PepTag® PKC Reaction Buffer, 2 μg of PepTag® C1 Peptide, and 5 μL of sonicated PKC Activator, 10 μL of purified samples or 4 μL of Protein Kinase C at concentration of 2.5 μg/mL (as a positive control) were added, followed by incubation at 30 ◦C for 30 min. Then, the reaction was stopped by placing the tube in a 95 ◦C heating block for 10 min. 1 μL of 80% glycerol was added to each sample, and then samples were electrophoretically separated on a 0.8% agarose gel at 100 V for 15 min. The phosphorylated peptide migrated to the cathode (+), while the non-phosphorylated peptide migrated to the anode (-). The gel was photographed on a transilluminator.

### *2.8. Analysis of ERK1*/*2*

The amount of total and phosphorylated ERK1/2 was quantified using SimpleStep ELISA Kit (Abcam, Cambridge, UK). Neutrophils (1 <sup>×</sup> 10<sup>6</sup> in 500 <sup>μ</sup>L of PBS per well) were stimulated with 100 μM and 200 μM FOH in 12-well microplate. Negative and positive controls were prepared as described above. After 1 h of incubation at 37 ◦C in 5% CO2, cells were lysed using a Cell Extraction Buffer (Abcam, Cambridge, UK). The protein concentration in the lysate was determined using the Bradford assay [31]. Then, 50 μL of lysate was mixed with 50 μL of antibody cocktail (anti-ERK1/2–total or anti-pT202/Y204–phosphorylated ERK1/2 provided by the manufacturer) in wells of SimpleStep pre-coated 96-well microplate, according to manufacturer's instruction. After 1 h of incubation with gentle shaking at room temperature, the wells were washed 3 times with PBS and then TMB (3,3 ,5,5 -tetramethylbenzidine) solution was added. After 15 min, the reaction was stopped with a Stop solution, and absorbance at 450 nm was recorded using a BioTek Synergy H1 microplate reader.

#### *2.9. Analysis of Netosis Signaling Pathway*

For the analysis of netosis signaling pathway, selected enzyme inhibitors were used. Neutrophils were pre-treated for 30 min prior to stimulation with different inhibitors: 30 μM piceatannol (Syk inhibitor; Sigma-Aldrich, St. Louis, MO, USA), 10 μM PP2 (Src inhibitor; Calbiochem, Darmstadt, Germany), 10 μM UO126 (ERK inhibitor; Cell Signaling Technology, Beverly, MA, USA) or 5 μM DPI (NADPH oxidase inhibitor; Sigma-Aldrich, St. Louis, MO, USA). Then, the cells were stimulated and analyzed as described above.

#### *2.10. Analysis of Neutrophil Chemotaxis*

Neutrophil migration was evaluated by the 24-well microchemotaxis chamber technique (Transwell®-Clear inserts, Corning, NY, USA). Neutrophils were labeled for 10 min with 1 μM CellTracker Red solution (Invitrogen, Carlsbad, CA, USA), washed three times with PBS, and placed in the upper compartment of the chamber (1 <sup>×</sup> 106 cells/well). Samples of FOH (various concentrations of up to 400 μM) or fMLP (1 μM; used as a positive control) were placed in the lower compartment. PBS was used as a negative control. The compartments were separated by a membrane with 3 μm pores. The chambers were incubated at 37 ◦C in 5% CO2 atmosphere for 1 h. Neutrophil migration was monitored by fluorescence microscopy (Motic AE31E, MoticEurope, Barcelona, Spain), and the number of cells migrated into the lower compartment of the chamber was quantified.

#### *2.11. Statistical Analysis*

Each of the experiments was repeated at least three times, obtaining consistent results. Two replicates were performed in each experiment. The graphs show the results of a single representative experiment.

Statistical analysis was performed with the GraphPad Prism 7 software (GraphPad Software, CA, USA). The statistical significance was assessed by ANOVA and Dunnett's multiple comparisons post-test.

#### **3. Results**

#### *3.1. Farnesol but Not Farnesoic Acid or Tyrosol Triggers NET Formation*

Neutrophils are the cells with high microbiocidal potential, and their high killing efficiency is due to their ability to recognize many "foreign" molecules released by pathogens. Owing to these properties and because their active cells move under the chemoattractant gradient, neutrophils release their antimicrobial molecules directly at the site of infection. In our current study, we focused on the role of *C. albicans*-released QSMs, in particular FOH.

We verified the neutrophil responses to contact with QSMs released by *C. albicans,* focusing on the PMN ability to release NETs. The stimulation of PMNs with FOH showed the dose-dependent responses with NET release within the whole range of concentrations tested (Figure 1a). The highest level of NETs released by FOH-treated neutrophils was observed for 250 μM FOH, and it reached about 60% of the positive control.

Different results were obtained for FA, a farnesol derivate. PMNs did not release NETs in response to any of the examined concentrations of FA. Although at the concentration of 250 μM the fluorescent signal increased significantly, it was probably a result of cell death induced by this FA concentration. Similar results were observed for TR, the third examined QSM. No released NETs were identified within the entire range of examined TR concentrations.

These quantitative results were confirmed by fluorescence microscopy analysis (Figure 1b), on which Sytox Green-stained neutrophils, stimulated with FOH showed cloud-like structures located around the human cells and composed of green-labeled DNA.

**Figure 1.** Release of neutrophil extracellular traps (NETs) by quorum sensing molecule (QSM)-treated neutrophils. Neutrophils were treated with QSMs at variable concentrations for 3 h. Unstimulated cells served as a negative control. (**a**) The released NETs were digested with micrococcal nuclease (MNase) and collected supernatants were stained with Sytox Green and the fluorescence intensity was measured. Data represent the mean fluorescence ± standard error of the mean (S.E.M.). from two replicates. ANOVA and Dunnett's multiple comparisons post-tests were used. Asterisks denote statistical significance (*p* > 0.1234 ns, \* *p* ≤ 0.0332, \*\*\*\* *p* < 0.0001). (**b**) The extracellular DNA was stained with Sytox Green and visualized with fluorescence microscopy.

To verify the NETs production during FOH treatment, also the extracellular localization of myeloperoxidase was determined, using specific, fluorescent antibodies. During netosis, MPO is moved from granule to the nucleus and then released together with DNA in the form of NETs.

Quantitative analysis of MPO in the samples containing neutrophils treated with FOH, was performed using ELISA. For this purpose, the netting cells were prior washed with PBS, and the extracellular DNA was digested with MNase to liberate bound MPO. In Figure 2a, we showed MPO concentration in the cell supernatants, determined by ELISA. The amount of detected MPO correlated with the DNA quantity, confirming that the observed DNA structures belong to the NETs. Additionally, MPO/DNA complexes were visualized microscopically (Figure 2b). As was shown in the figure, the protein location correlated with DNA clouds released by PMNs, confirming activation of netosis by FOH.

**Figure 2.** Identification of myeloperoxidase in farnesol (FOH)-treated neutrophils. Neutrophils were treated with FOH at a concentration of 100 μM and 200 μM for 3 h. Unstimulated cells served as a negative control. (**a**) The released DNA was digested with MNase, and a concentration of myeloperoxidase (MPO) was analyzed in collected supernatants using ELISA method. Data represent the mean concentration ± S.E.M. from two replicates. ANOVA and Dunnett's multiple comparisons post-tests were used. Asterisks denote statistical significance (*p* > 0.1234 ns, \* *p* ≤ 0.0332, \*\* *p* ≤ 0.0021). (**b**) The extracellular DNA was stained with Sytox Green, while MPO was identified with primary mouse anti-MPO antibodies and secondary Alexa Fluor 555-labeled anti-mouse antibodies. Samples were visualized with fluorescence microscopy.

#### *3.2. Farnesol Treatment of Neutrophils Does Not Lead to Cell Apoptosis*

In order to confirm that the observed effect of the DNA release the cell was associated with the mechanism of netosis, but not cell death like apoptosis, the activation of proapoptotic caspases was analyzed. PMNs were exposed to FOH and FA at the concentration range of 0–1200 μM for one hour, and caspase 3 and 7 activities were measured using the chemiluminometric method (Figure 3).

**Figure 3.** Caspase 3/7 activity in QSM-treated neutrophils. Neutrophils were treated with FOH and farnesoic acid (FA) at variable concentrations, and incubated with Caspase-Glo® 3/7, followed by continuous measurement of chemiluminescence for 2 h. Data represent mean values of luminescence from two replicates ± S.E.M. ANOVA and Dunnett's post-tests were used. Asterisks denote statistical significance (*p* > 0.1234 ns, \* *p* ≤ 0.0332, \*\* *p* ≤ 0.0021, \*\*\* *p* ≤ 0.0002, \*\*\*\* *p* < 0.0001).

The results presented in Figure 3 indicated that FOH at the concentration range of 0–200 μM did not activate proapoptotic caspases in neutrophils. However, higher concentrations of FOH led to apoptotic cell death. FA-treated neutrophils showed activation of apoptosis in a dose-dependent manner within the whole tested range of concentrations. This explains the absence of NETs released in the presence of FA, because activation of caspases 3 and 7 inhibits the release of NETs [32].

Besides, the neutrophils were incubated with FOH and FA at concentrations of 100 μM and 200 μM for one hour and then labeled with PI and AnV. The cell viability analysis based on PI and AnV-FITC was performed using flow cytometry. The results (Figure 4a), comparison to the 3/7 caspase analysis, also allow assessing the potential of PMNs for the release of NETs. The changes in the distribution of cell population labeled with PI and AnV (Figure 4b), characteristic to netosis, were previously described and used by Masuda et al. [33].

**Figure 4.** Flow cytometric analysis of neutrophil apoptosis. Neutrophils were stimulated with FOH or FA, labeled with annexin V-FITC and propidium iodide, and analyzed by flow cytometry. The results are presented as a percentage ratio of the signal detected for whole cell population and showing no cell death (PI−AnV−), early apoptosis (PI−An+) and late apoptosis (PI+AnV+). Cells stained only with propidium iodide (PI+AnV−) represent the necrotic or NET-forming cells. For each sample, data were collected for 100,000 neutrophils. (**a**) Data are presented as a part of total cell population, (**b**) the diagrams represent AnV and PI cell distribution.

The results showed four groups of cells: AnV−PI<sup>−</sup> identified as live; AnV+PI<sup>−</sup> identified as early apoptotic; AnV+PI<sup>+</sup> identified as apoptotic; AnV−PI<sup>+</sup> identified as necrotic. However, the obtained cell distribution patterns, especially for AnV+PI<sup>+</sup> cells, were identical for cells in the netosis, represented by PMNs stimulated with PMA. Approximately 35%–40% of FOH-treated neutrophils showed positive labeling for both AnV and PI, similarly to the PMA-treated positive control. This indicated that these cells lost the integrity of the cell membrane, however, the comparison of these results with the activation of caspases led to the conclusion that observed changes did not result from apoptotic cell death. The low percentage of PI-positive neutrophils suggests that DNA released outside the cells was not due to mechanical destruction or necrotic death. The presence of approximately 15% of AnV+PI<sup>−</sup> cells suggests that neutrophils were in the ongoing netosis process. In contrast, FA-treated PMNs did

not show apoptotic and necrotic cell traits within the tested range of QSM concentrations, within one hour of contact with FA.

#### *3.3. Farnesol Induces Rapid ROS Production*

The activation of NADPH oxidase, resulting in the production of ROS is a key step in the ROS-dependent netosis pathway. The chemical stimulation of neutrophils with PMA leads to the release of a high amount of ROS within minutes of activation. We checked the ability of the examined QSMs to activate NADPH oxidase and release ROS. PMNs were treated with selected doses of FOH or FA, and the activity of NADPH oxidase was measured using a chemiluminescence-based assay.

Changes in chemiluminescence intensity over the time showed in Figure 5 were proportional to the amount of ROS produced by PMNs. The stimulation of neutrophils with FOH at the concentration of 100 and 200 μM caused the rapid activation of NADPH oxidase and release of ROS at a time similar to PMA-stimulated cells. The level of ROS released by PMNs treated with 100 μM FOH reached about 25% of the positive control response, while activation with 200 μM FOH resulted in two-fold higher ROS production. Release of ROS by neutrophils correlated with the production of NETs, suggesting that FOH activates the ROS-dependent mechanism of netosis.

**Figure 5.** The time course of reactive oxygen species (ROS) generation by neutrophils in response to FOH and FA. ROS production by neutrophils (2 <sup>×</sup> 105 cells/well) was monitored at 37 ◦C for 1 h by the luminol chemiluminescence method after suspension of cells in FOH or FA. In the reference samples, the neutrophils were treated with 25 nM phorbol 12-myristate 13-acetate (PMA) or left in Krebs–Ringer phosphate buffer (a negative control).

The response of neutrophils to FA presented relatively low chemiluminescence signal, confirming that this QSM does not participate in NET release.
