A confocal microscopy based method to monitor extracellular pH in fungal biofilms

A confocal microscopy based method to monitor extracellular pH in fungal biofilms Abstract pH in fungal biofilms is important for a variety of fungal infections and industrial applications involving fungal biofilms, but to date, it has never been measured directly inside the biofilm matrix. In the present study, a new methodology was developed allowing for confocal microscopy based monitoring of extracellular pH inside fungal biofilms. Monospecies biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans were stained with the pH dependent ratiometric probe C-SNARF-4, imaged with a confocal microscope, and a digital image analysis procedure was developed to determine pH in the extracellular matrix. As a proof of concept, pH developments at the biofilm–substratum interface were monitored for 1 h after exposure to glucose. Observed pH drops differed considerably between the different species and also between replicate biofilms of the same species. Candida albicans biofilms showed the highest acidogenicity, with pH drops occurring much faster than in planktonic culture. pH ratiometry with C-SNARF-4 is a valuable tool to get insight into fungal biofilm metabolism and may shed new light on both disease-related and industrially relevant processes in fungal biofilms. Aspergillus, Candida, confocal laser scanning microscopy, Cryptococcus, C-SNARF-4, Saccharomyces INTRODUCTION In the past decade, the concept of fungal biofilm formation has gained increasing importance in mycology. Fungal biofilms are involved in a multitude of human diseases, especially in immune compromised patients, but also in connection with indwelling-device-related infections (Williams and Ramage 2015). On the other hand, fungal biofilms possess a huge potential for industrial use. The metabolic activity in surface-attached fungi is employed in microbial fuel cells, for purposes of bioremediation and in the food industry (Wang and Chen 2009; Gutiérrez-Correa et al.2012). In the medical field, Candida species, such as Candida albicans and Candida dubliniensis, are among the most important opportunistic fungal pathogens. In the oral cavity, they are frequently associated with endodontic infections (Siqueira and Sen 2004; Persoon, Crielaard and Ozok 2016), periodontal disease (Sardi et al.2010) and with denture stomatitis (Salerno et al.2011). Moreover, Candida biofilms have been implicated in vulvovaginal infections (Harriott et al.2010; Sherry et al.2017), in chronic wound infections (Dowd et al.2011) and in infections originating from intravascular and urinary catheters (Cauda 2009). In recent years, increasing evidence has accumulated linking biofilm formation to infections with Cryptococcus neoformans and Aspergillus spp., most notably Aspergillus fumigatus. Cryptococcus neoformans has been shown to form biofilms on indwelling devices, such as cardiac valves (Banerjee, Gupta and Venugopal 1997) and prosthetic joints (Johannsson and Callaghan 2009; Shah, Shoham and Nayak 2015), and is the main cause of cryptococcal meningoencephalitis (Sloan and Parris 2014). Aspergillus fumigatus biofilms are involved in severe infections of the upper and lower airways (Loussert et al.2010), and have been associated with cases of endocarditis (Escande et al.2011; McCormack and Pollard 2011). In sharp contrast to these detrimental effects, biofilms of Aspergillus spp. have also been exploited for industrial applications, such as the degradation of polymers (Upreti and Srivastava 2003; Sangeetha Devi et al.2015), the removal of heavy metals (Dias et al.2002) and the production of food ingredients and enzymes (Karaffa and Kubicek 2003; Villena and Gutiérrez-Correa 2007; Gamarra, Villena and Gutiérrez-Correa 2010). Likewise, biofilms of Saccharomyces cerevisiae, baker's yeast, serve multiple purposes in different industrial branches, including the production of bioethanol (Inloes et al.1983; Ciesarová et al.1998; Desimone et al.2002; Chen et al.2013), electricity (Rahimnejad et al.2012) and fermented beverages (Cortes et al.1998; Legras, Erny and Charpentier 2014; Marin-Menguiano et al.2017). pH plays an important role in many, if not all of the above named biofilm-related processes. It is intimately related to fungal virulence, and it has a dramatic impact on treatment strategies and the efficacy of antifungal drugs (Liu et al.2011; Danby et al.2012). Some pathologic conditions, such as periodontal disease (Pöllänen, Paino and Ihalin 2013) or chronic wound infections (Gethin 2007; Schneider et al.2007), are accompanied or favoured by a shift to alkaline pH, whereas other conditions, such as endodontic infections or denture stomatitis, are characterised by slight or strong pH drops (Samaranayake et al.1983; Nikawa et al.1994; Nekoofar et al.2009; Vasconcellos et al.2015). Likewise, up- or downregulation of acid generating processes have been shown to be important during infection with A. fumigatus and C. neoformans (Price et al.2011; Gibbons et al.2012). In industrial settings, the manipulation of pH has been shown to directly influence the yield during fungal production of bioethanol (Lin et al.2012; Mohd Azhar et al.2017) and the power density in microbial fuel cells (Ganguli and Dunn 2012). Despite the paramount importance of pH, it has, to the best of our knowledge, never been monitored directly inside the extracellular matrix of fungal biofilms, but only in the surrounding bulk fluid. Measurements of pH in the matrix of bacterial biofilms, using either microelectrodes (Schreiber et al.2010; Von Ohle et al.2010) or confocal microscopy based techniques (Schlafer and Meyer 2016), have demonstrated the presence of steep pH gradients and distinct microenvironments, contributing to a deeper understanding of biofilm metabolism (Xiao et al.2012; Dige et al.2016). We have recently developed a confocal microscopy based technique that exploits the staining properties and pH-dependent emission shift of the ratiometric dye C-SNARF-4 to map pH developments in bacterial biofilms (Schlafer et al.2015). Compared to other quantitative confocal microscopy based approaches, pH ratiometry with C-SNARF-4 offers certain advantages. Due to the pH-dependent emission shift of the dye, pH can be determined irrespective of the concentration of the dye (Marcotte and Brouwer 2005). Moreover, C-SNARF-4 penetrates biofilms easily and quickly, which has proven to be problematic when pH sensitive dyes are immobilised on nanoparticles (Hidalgo et al.2009). Finally, C-SNARF-4 can be employed in bacterial biofilms without the addition of a second dye, as it stains bacterial cells and the surrounding biofilm matrix with different fluorescence intensity levels. The latter property allows for an intensity threshold based identification of matrix areas in the confocal images, which is crucial, as the matrix pH differs from intracellular pH due to bacterial homeostasis (Schlafer and Meyer 2016). In summary, C-SNARF-4-based pH ratiometry allows monitoring pH developments in the matrix of bacterial biofilms in real-time, in all three dimensions, but it has never been applied to fungal biofilms. As the fungal cell wall structure differs fundamentally from the one of bacteria (Bowman and Free 2006), the aims of the present study were to (i) investigate the staining properties of C-SNARF-4 on fungi, using the well-studied A. fumigatus, C. albicans, C. dubliniensis, C. neoformans and S. cerevisiae; and to (ii) develop a procedure for reliable confocal microscopy based quantification of extracellular pH in biofilms formed by these species. MATERIALS AND METHODS Fungal strains Aspergillus fumigatus (DSM 790), Candida albicans (NCPF 3179), Candida dubliniensis (DSM 13 268), Cryptococcus neoformans (DSM 6972) and Saccharomyces cerevisiae (DSM 3799) were cultivated aerobically on Sabouraud dextrose agar containing penicillin and streptomycin (Thermo Fisher Scientific, Roskilde, Denmark) at 35°C. A. fumigatus was harvested from the agar plates by flooding the surface with 0.9% sterile NaCl containing 0.05% (v/v) Tween 80 (Sigma-Aldrich, Brøndby, Denmark) according to the protocol described by Shirazi et al. (2016), while all other strains were harvested with inoculation loops. Prior to experimental use, the strains were transferred to Brain Heart Infusion (Sigma-Aldrich, Brøndby, Denmark) and grown until late exponential phase (18–24 h) under aerobic conditions at 35°C. Visualisation of immobilised planktonic fungi with C-SNARF-4 To test if C-SNARF-4 visualised all employed fungal strains, the staining properties of C-SNARF-4 were compared to BacLight. Planktonic cultures were washed in 0.9% sterile NaCl and adjusted to an optical density of 0.2 (550 nm). HEPES buffer solutions (50 mM) were used to titrate the pH of the cultures to 4.5–8, in steps of 0.5 pH units. A. fumigatus, C. dubliniensis, C. neoformans and S. cerevisiae were set to settle for 1 h in optical-bottom 96-well plates (ibidi, Planegg/Martinsried, Germany) coated with porcine gelatin (Type A; Sigma-Aldrich, Brøndby, Denmark; 0.2% w V−1 in Milli-Q water). C. albicans did not attach to gelatin-coated surfaces and was therefore immobilised in Concanavalin A (Sigma-Aldrich, Brøndby, Denmark) coated 96-well plates. To remove loosely adherent cells, the wells were washed twice with HEPES buffer of the same pH. Then C-SNARF-4 (Life Technologies, Nærum, Denmark) was added to a concentration of 30 μM and the cells were imaged with confocal laser scanning microscopy (CLSM). Thereafter, the fungi were counterstained with BacLight (Thermo Fisher Scientific, Roskilde, Denmark) according to the manufacturer's instructions, and images were acquired in the same microscopic fields of view. The experiments were carried out in duplicate. Visualisation of fungi in biofilms with C-SNARF-4 Biofilm growth of A. fumigatus, C. albicans, C. dubliniensis, C. neoformans and S. cerevisiae was performed modifying published protocols (Ramage et al.2001a; Ramage et al.2001b; Ravi et al.2009; Shirazi et al.2016). Planktonic cultures of the employed strains were washed in phosphate-buffered saline (PBS; pH 7.4; Sigma-Aldrich¸ Brøndby, Denmark) and adjusted to optical densities (550 nm) of 0.05 (A. fumigatus, C. dubliniensis, C. neoformans and S. cerevisiae) or 0.5 (C. albicans and S. cerevisiae). Optical bottom 96-well plates (ibidi, Planegg/Martinsried, Germany) were coated for 30 min at 35°C with sterile saliva prepared according to the method of de Jong et al. (1984), and 100 μl of the planktonic suspensions was set to settle for 1.5 h at 35°C. Thereafter, the wells were washed twice with PBS to remove non-adherent cells, and 200 μl of Yeast Nitrogen Base (Sigma-Aldrich, Brøndby, Denmark) containing 100 mM glucose was added. Biofilm growth was carried out at 35°C under aerobic conditions for 24 h, after which stable biofilms had grown for all employed species except S. cerevisiae. We performed additional experiments with S. cerevisiae to test if biofilm growth occurred in the absence of a salivary coating or using other cell concentrations in the initial inoculum, but none of the setups yielded stable biofilms. Saccharomyces cerevisiae was therefore excluded from subsequent experiments. After biofilm growth, all wells were washed three times with HEPES buffer solutions (50 mM) titrated to pH 4.5–8 in steps of 0.5 pH units, and C-SNARF-4 was added to a concentration of 30 μM. CLSM images were acquired in different microscopic fields of view for each strain and pH, the wells were counterstained with BacLight, and identical fields of view were imaged again. Experiments were performed in duplicate for all strains and pH values. Monitoring of pH in fungal biofilms As a proof of concept, we monitored pH developments in monospecies biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans following exposure to glucose. For each strain, triplicate biofilms were grown, washed twice with sterile 0.9% NaCl and then incubated for 1 h with 0.9% NaCl containing C-SNARF-4 (30 μM) and glucose (0.4% w/v). In each biofilm, CLSM images were acquired 5 μm above the biofilm–substratum interface in three different microscopic fields of view. 5, 10, 15, 20, 30 and 60 min after the addition of glucose extracellular pH was determined as described below. Additionally, Z-stacks were recorded in selected fields of view to record vertical pH profiles of the biofilms. The experiments were performed in duplicate. Confocal laser scanning microscopy Confocal laser scanning microscopy (CSLM) All images were acquired with a Zeiss LSM 510 META (Zeiss, Jena, Germany) equipped with a 63× oil immersion objective (Plan-Apochromat; NA = 1.4). C-SNARF-4 was excited at 543 nm and detected from 576 to 608 nm (green channel) and 629 to 661 nm (red channel; Marcotte and Brouwer 2005). SYTO 9 and propidium iodide, the components of BacLight, were excited at 488 nm and 543 nm, respectively, and detected from 500 to 554 nm and 554 to 608 nm. The pinhole was set to 2 Airy Units (optical slice 1.6 μm), the images were 2048 × 2048 pixels in size (143 × 143 μm) and acquired with a pixel dwell time of 1.6 μs, line average 4. Calibration of C-SNARF-4 For pH analysis, the ratios of the fluorescence intensities of C-SNARF-4 in the two different detection windows are calculated and converted into pH values based on a calibration curve. The calibration was performed using HEPES buffer solutions (50 mM) containing C-SNARF-4 (30 μM), which were titrated to pH 3.0–8.5 and imaged with the same laser settings used for biofilm image acquisition. After each buffer image, an image with the laser turned off was taken to correct for detector offset. Calibration images were taken in triplicate in different microscopic fields of view; the means of the ratios were plotted against the pH of the buffer solutions and fitted to the following function, assuming a five-parameter logistic curve (MyCurveFit; MyAssays Ltd, Brighton, UK): \begin{eqnarray} pH = \left[ {{{\left( {{{\left( {\frac{{2.9371815}}{{Ratio - 0.1000935}}} \right)}^{\frac{1}{{5298906}}}} - 1} \right)}^{\frac{1}{{6.649332}}}}} \right] \times 61.95654 \end{eqnarray} (1) Calibration data is shown in Fig. S1, Supporting Information. Digital image analysis Both channels (green and red) of the CLSM images of the C-SNARF-4-stained biofilms were exported separately as TIF files into the software daime (Daims, Lücker and Wagner 2006). To remove all fungal cells from the biofilm images, the images of the brightest channel were segmented with both a lower and an upper brightness threshold. With the lower threshold, the intracellular space of viable cells was removed from the images. The higher threshold removed the fungal cell walls and the cytoplasm of membrane-compromised cells from the images, leaving only the extracellular space for subsequent pH analysis. The object layer of the segmented images was then transferred to the corresponding images of the other channel, and both image series were exported into ImageJ (Schneider, Rasband and Eliceiri 2012). In ImageJ, the mean filter (radius: 1 pixel) was applied for noise reduction, the green channel images were divided by the red channel images and the average ratios and standard deviations were calculated for all microscopic fields of view. Equation (1) was used to translate the ratios into pH values, and false colouring was applied for graphic representation. The digital image analysis procedure is presented in more detail in Schlafer and Dige (2016). RESULTS C-SNARF-4 stained all five employed fungal strains with an identical pattern. At all tested pH values, the dye bound strongly to the cell wall of immobilised viable cells but left the cytoplasm unstained. In membrane-compromised cells, C-SNARF-4 accumulated in the cytoplasm. Counterstaining with BacLight showed that all cells were targeted by C-SNARF-4 (Figs S2 and S3, Supporting Information). Likewise, C-SNARF-4 stained the walls of all cells, including hyphae, in single-species biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans, irrespective of the pH (Fig. 1). This staining pattern resulted in three fluorescence intensity levels in all acquired biofilm images, illustrated in Fig. 2 for C. dubliniensis (Figs 2A–E) and A. fumigatus (Figs 2F–J): In both channels (green and red), the cytoplasm of viable cells showed the lowest intensity, while the cell walls of all cells and the cytoplasm of membrane-compromised cells showed the highest intensity. The surrounding biofilm matrix displayed an intermediate level of fluorescence (Figs 2A–C, F–H). The staining pattern allowed for an intensity threshold based image analysis to identify and remove all fungal cells from the biofilm images. Images were segmented with both a lower threshold to remove the cytoplasm of viable cells and with a higher threshold to remove membrane-compromised cells and cell walls. Thereafter, only the biofilm matrix was left for pH analysis (Fig. 2D and I). Green/red fluorescence ratios were calculated and false colouring was applied for visualisation purposes (Fig. 2E and J). Figure 1. View largeDownload slide Visualisation of fungal biofilms with C-SNARF-4. Biofilms of Candida dubliniensis (A, B, E, F), Candida albicans (C, D, G, H), Cryptococcus neoformans (I, J, M, N) and Aspergillus fumigatus (K, L, O, P) were stained with C-SNARF-4 at different pH values, imaged with a confocal microscope (A–D and I–L), counterstained with BacLight and imaged again (E–H and M–P). At all investigated pH values, C-SNARF-4 stained the walls of all fungal cells in the samples, including hyphens, as shown by counterstaining with BacLight. The fungal cytoplasm remained unstained, except when cells were membrane-compromised (yellow/red cells in panels C, D, I, J and L). pH values are indicated in the C-SNARF-4 images. Bars = 20 μm. Figure 1. View largeDownload slide Visualisation of fungal biofilms with C-SNARF-4. Biofilms of Candida dubliniensis (A, B, E, F), Candida albicans (C, D, G, H), Cryptococcus neoformans (I, J, M, N) and Aspergillus fumigatus (K, L, O, P) were stained with C-SNARF-4 at different pH values, imaged with a confocal microscope (A–D and I–L), counterstained with BacLight and imaged again (E–H and M–P). At all investigated pH values, C-SNARF-4 stained the walls of all fungal cells in the samples, including hyphens, as shown by counterstaining with BacLight. The fungal cytoplasm remained unstained, except when cells were membrane-compromised (yellow/red cells in panels C, D, I, J and L). pH values are indicated in the C-SNARF-4 images. Bars = 20 μm. Figure 2. View largeDownload slide Digital image analysis procedure for ratiometric determination of extracellular pH in fungal biofilms. Biofilms of Candida dubliniensis (A–E) and Aspergillus fumigatus (F–J) were stained with C-SNARF-4 in the presence of glucose. In both detection channels (green: A and F; red: B and G), the cytoplasm of viable cells displays the lowest fluorescence intensity, while the cell walls as well as the cytoplasm of membrane compromised cells (bright cell in A–C) show the highest intensity. Fluorescence deriving from the extracellular matrix shows an intermediate level of intensity. Panels C and H show overlays of the green and red channels. Based on intensity thresholding with both a lower and a higher threshold, all cells can be removed from the images, leaving only the biofilm matrix for pH analysis (D and I). The ratios of fluorescence intensities are calculated, converted to pH values, and in E and J, false colours were applied to visualise pH in the matrix. After 30 min of exposure to glucose, average pH in the selected fields of view was 5.9 for C. dubliniensis and 6.9 for A. fumigatus. Bars = 20 μm. Figure 2. View largeDownload slide Digital image analysis procedure for ratiometric determination of extracellular pH in fungal biofilms. Biofilms of Candida dubliniensis (A–E) and Aspergillus fumigatus (F–J) were stained with C-SNARF-4 in the presence of glucose. In both detection channels (green: A and F; red: B and G), the cytoplasm of viable cells displays the lowest fluorescence intensity, while the cell walls as well as the cytoplasm of membrane compromised cells (bright cell in A–C) show the highest intensity. Fluorescence deriving from the extracellular matrix shows an intermediate level of intensity. Panels C and H show overlays of the green and red channels. Based on intensity thresholding with both a lower and a higher threshold, all cells can be removed from the images, leaving only the biofilm matrix for pH analysis (D and I). The ratios of fluorescence intensities are calculated, converted to pH values, and in E and J, false colours were applied to visualise pH in the matrix. After 30 min of exposure to glucose, average pH in the selected fields of view was 5.9 for C. dubliniensis and 6.9 for A. fumigatus. Bars = 20 μm. As a proof of concept, biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans were incubated with 0.4% glucose, and the extracellular pH at the bottom of the biofilms, 5 μm above the biofilm–substratum interface, was monitored for 1 h. The pH development in the biofilms differed markedly between the strains. C. albicans biofilms showed the lowest pH, typically around 5 (Fig. 3; Fig. S4A, Supporting Information), which corresponds to the pH levels reached in planktonic suspension upon exposure to glucose (data not shown). Interestingly, these low pH levels were reached much faster than in planktonic culture (pH 5 after 7 h of incubation), typically within 5 min after exposure to glucose (Fig. S5A, Supporting Information). Acid production in C. dubliniensis biofilms occurred more slowly, but after 1 h of exposure to glucose, pH levels approached 5–5.5, too (Fig. 3; Fig S4B, Supporting Information). Under the chosen experimental conditions, the pH in A. fumigatus and C. neoformans biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h (Fig. 3; Fig. S4C and D, Supporting Information). In general, pH differences between replicate biofilms were more pronounced than between different fields of view within the same biofilms (Fig. S4, Supporting Information). Under the chosen conditions, only slight differences between different areas within one biofilm could be observed. Likewise, vertical pH profiles showed a constant pH across the slim biofilms (ca. 20 μm) at a given time (see Fig. S5B, Supporting Information, for examples). Figure 3. View largeDownload slide Typical pH development in biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans after incubation with glucose. In each biofilm, extracellular pH was measured ratiometrically 5 μm above the biofilm–substratum interface in three different microscopic fields of view. Each line represents the pH development in one field of view in the course of 1 h. pH dropped rapidly (within 5 min) from 7 to 5 in C. albicans biofilms (blue lines) and remained stable afterwards. In C. dubliniensis biofilms (grey/black lines), pH dropped more slowly and approached levels of 5.5 at the end of the incubation period. pH in A. fumigatus (green lines) and C. neoformans (red lines) biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h. Error bars = standard deviations. Figure 3. View largeDownload slide Typical pH development in biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans after incubation with glucose. In each biofilm, extracellular pH was measured ratiometrically 5 μm above the biofilm–substratum interface in three different microscopic fields of view. Each line represents the pH development in one field of view in the course of 1 h. pH dropped rapidly (within 5 min) from 7 to 5 in C. albicans biofilms (blue lines) and remained stable afterwards. In C. dubliniensis biofilms (grey/black lines), pH dropped more slowly and approached levels of 5.5 at the end of the incubation period. pH in A. fumigatus (green lines) and C. neoformans (red lines) biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h. Error bars = standard deviations. DISCUSSION pH in fungal biofilms is of fundamental importance for both medical conditions and industrial applications. The present study is the first to describe a methodology to monitor pH in the matrix of fungal biofilms, using a combination of confocal microscopy with the ratiometric dye C-SNARF-4, and digital image post-processing. Unlike electrode measurements in bulk fluid, it provides not only a good temporal, but also a high spatial resolution. As a proof of concept, we monitored pH over time in monospecies biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans, exposed to glucose. Of the employed strains, C. albicans was by far the most acidogenic under the chosen conditions. pH inside the biofilms dropped from near neutral to values around five within less than 5 min (Fig. 3; Figs S4 and S5, Supporting Information), which is much faster than the pH drops reported in planktonic culture (Samaranayake et al.1986) and may be explained by the high local cell density inside the biofilms and the modulating effect of the biofilm matrix (Koo, Falsetta and Klein 2013). Rapid pH drops inside fungal biofilms may be of particular importance for antifungal therapy, as the efficacy of many antifungal drugs decreases at low pH (Liu et al.2011; Danby et al.2012). Acid production in biofilms of C. dubliniensis occurred much slower; the pH typically still dropped after half an hour of exposure to glucose, reaching levels of 5.2–6.2 (Fig. 3; Fig. S4, Supporting Information). Although most cells of C. dubliniensis were still viable after a short (<30 min) exposure to low pH (Figs S2 E–G), the low acidogenicity, compared to C. albicans, may in part explain the predominance of C. albicans in infections associated with low pH, such as denture stomatitis (Gendreau and Loewy 2011). In biofilms of A. fumigatus and C. neoformans, pH stayed in the near-neutral range under the chosen conditions, although, and this holds true for all investigated species, notable differences could be observed between replicate biofilms (Fig. 3; Fig. S4, Supporting Information). In contrast, only minor differences in pH were observed between different microscopic fields of view within the same biofilm, irrespective of the species. This homogenous behaviour may be attributed to the age of the biofilms (24 h), which were still in the maturation phase by the time pH was recorded, and, in particular, to the fact that the present study employed monospecies biofilms. Ratiometric pH measurements in the matrix of young dental biofilms, consisting of several different bacterial species, have revealed considerably more variation in pH within one biofilm specimen (Dige et al.2016). Studying pH developments in multispecies fungal biofilms is therefore of great interest, but beyond the scope of the present investigation. The staining pattern of C-SNARF-4 in biofilms was identical for all strains employed in this study: membrane-compromised cells and the walls of all cells showed a brighter fluorescence than the biofilm matrix, whereas the cytoplasm of viable cells remained unstained. These different levels of fluorescence intensity allowed for a reliable removal of fungal cells from the confocal images and thus the determination of extracellular pH only (Fig. 2). As bacterial cells have been shown to up-concentrate C-SNARF-4 irrespective of their viability status (Schlafer et al.2015), the same image analysis procedure will also be applicable to cross-kingdom biofilms. The strains employed in the present work all belong to the phyla Basidiomycota (C. neoformans) or Ascomycota (A. fumigatus, C. albicans, C. dubliniensis and Saccharomyces cerevisiae) and were selected because of their relevance in the medical field and/or industrial applications. Although S. cerevisiae did not form biofilms under the chosen conditions, it is likely that pH in S. cerevisiae biofilms can be monitored using C-SNARF-4, as the staining pattern of immobilised cells was identical to the one observed for the other species (Fig. S3C and D, Supporting Information). However, given the considerable differences in the fungal cell wall structure, it still remains to be investigated if the presented methodology is applicable to all biofilm forming fungi. The use of confocal microscopy imposes some limitations on C-SNARF-4-based pH ratiometry. While pH at the biofilm–substratum interface can be recorded irrespective of the dimensions of the biofilm, only thin biofilms with a thickness of up to 50 μm can be imaged entirely. If vertical gradients in thicker biofilms are to be assessed, motor-controlled pH microelectrodes may be used (Von Ohle et al.2010). While the image post-processing necessary for pH ratiometry takes time, image acquisition is rather fast. In the present study, an acquisition time of approximately 1 min was chosen, but if need be, scanning parameters can be adjusted accordingly and images can be acquired in less than 10 s, at the expense of detail. With a pka of ∼6.4, C-SNARF-4 is suitable for pH determination in both acidic and moderately alkaline environments. At very low (<4) and at high (>8) pH, the discriminative power gets lower, as can be seen from the resulting bigger error bars (Fig. 3; Fig. S5, Supporting Information). In conclusion, the present work provides a new tool for the confocal microscopy based determination of extracellular pH inside fungal biofilms. pH developments can be monitored in real-time, in all three dimensions, in the pH range between 4 and 8. pH ratiometry with C-SNARF-4 can contribute to a more thorough understanding of fungal biofilm metabolism, which may be important for the treatment of fungal diseases and the optimisation of fungal biofilm-based industrial processes. SUPPLEMENTARY DATA Supplementary data are available at FEMSYR online. Acknowledgements Mette Nikolajsen is acknowledged for excellent technical support. FUNDING This work was supported by AIAS, Aarhus Institute of Advanced Studies, Aarhus University and Aarhus University Research Foundation (AUFF). Conflicts of interest. None declared. 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Potential of biofilm-based biofuel production . Appl Microbiol Biot 2009 , DOI: 10.1007/s00253-009-1940-9 . Williams C , Ramage G . Fungal biofilms in human disease . Adv Exp Med Biol 2015 , DOI: 10.1007/978-3-319-09782-4_2 . Xiao J , Klein MI , Falsetta ML et al. The exopolysaccharide matrix modulates the interaction between 3D architecture and virulence of a mixed-species oral biofilm . PLoS Pathog 2012 , DOI: 10.1371/journal.ppat.1002623 . © FEMS 2018. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png FEMS Yeast Research Oxford University Press

A confocal microscopy based method to monitor extracellular pH in fungal biofilms

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Abstract

Abstract pH in fungal biofilms is important for a variety of fungal infections and industrial applications involving fungal biofilms, but to date, it has never been measured directly inside the biofilm matrix. In the present study, a new methodology was developed allowing for confocal microscopy based monitoring of extracellular pH inside fungal biofilms. Monospecies biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans were stained with the pH dependent ratiometric probe C-SNARF-4, imaged with a confocal microscope, and a digital image analysis procedure was developed to determine pH in the extracellular matrix. As a proof of concept, pH developments at the biofilm–substratum interface were monitored for 1 h after exposure to glucose. Observed pH drops differed considerably between the different species and also between replicate biofilms of the same species. Candida albicans biofilms showed the highest acidogenicity, with pH drops occurring much faster than in planktonic culture. pH ratiometry with C-SNARF-4 is a valuable tool to get insight into fungal biofilm metabolism and may shed new light on both disease-related and industrially relevant processes in fungal biofilms. Aspergillus, Candida, confocal laser scanning microscopy, Cryptococcus, C-SNARF-4, Saccharomyces INTRODUCTION In the past decade, the concept of fungal biofilm formation has gained increasing importance in mycology. Fungal biofilms are involved in a multitude of human diseases, especially in immune compromised patients, but also in connection with indwelling-device-related infections (Williams and Ramage 2015). On the other hand, fungal biofilms possess a huge potential for industrial use. The metabolic activity in surface-attached fungi is employed in microbial fuel cells, for purposes of bioremediation and in the food industry (Wang and Chen 2009; Gutiérrez-Correa et al.2012). In the medical field, Candida species, such as Candida albicans and Candida dubliniensis, are among the most important opportunistic fungal pathogens. In the oral cavity, they are frequently associated with endodontic infections (Siqueira and Sen 2004; Persoon, Crielaard and Ozok 2016), periodontal disease (Sardi et al.2010) and with denture stomatitis (Salerno et al.2011). Moreover, Candida biofilms have been implicated in vulvovaginal infections (Harriott et al.2010; Sherry et al.2017), in chronic wound infections (Dowd et al.2011) and in infections originating from intravascular and urinary catheters (Cauda 2009). In recent years, increasing evidence has accumulated linking biofilm formation to infections with Cryptococcus neoformans and Aspergillus spp., most notably Aspergillus fumigatus. Cryptococcus neoformans has been shown to form biofilms on indwelling devices, such as cardiac valves (Banerjee, Gupta and Venugopal 1997) and prosthetic joints (Johannsson and Callaghan 2009; Shah, Shoham and Nayak 2015), and is the main cause of cryptococcal meningoencephalitis (Sloan and Parris 2014). Aspergillus fumigatus biofilms are involved in severe infections of the upper and lower airways (Loussert et al.2010), and have been associated with cases of endocarditis (Escande et al.2011; McCormack and Pollard 2011). In sharp contrast to these detrimental effects, biofilms of Aspergillus spp. have also been exploited for industrial applications, such as the degradation of polymers (Upreti and Srivastava 2003; Sangeetha Devi et al.2015), the removal of heavy metals (Dias et al.2002) and the production of food ingredients and enzymes (Karaffa and Kubicek 2003; Villena and Gutiérrez-Correa 2007; Gamarra, Villena and Gutiérrez-Correa 2010). Likewise, biofilms of Saccharomyces cerevisiae, baker's yeast, serve multiple purposes in different industrial branches, including the production of bioethanol (Inloes et al.1983; Ciesarová et al.1998; Desimone et al.2002; Chen et al.2013), electricity (Rahimnejad et al.2012) and fermented beverages (Cortes et al.1998; Legras, Erny and Charpentier 2014; Marin-Menguiano et al.2017). pH plays an important role in many, if not all of the above named biofilm-related processes. It is intimately related to fungal virulence, and it has a dramatic impact on treatment strategies and the efficacy of antifungal drugs (Liu et al.2011; Danby et al.2012). Some pathologic conditions, such as periodontal disease (Pöllänen, Paino and Ihalin 2013) or chronic wound infections (Gethin 2007; Schneider et al.2007), are accompanied or favoured by a shift to alkaline pH, whereas other conditions, such as endodontic infections or denture stomatitis, are characterised by slight or strong pH drops (Samaranayake et al.1983; Nikawa et al.1994; Nekoofar et al.2009; Vasconcellos et al.2015). Likewise, up- or downregulation of acid generating processes have been shown to be important during infection with A. fumigatus and C. neoformans (Price et al.2011; Gibbons et al.2012). In industrial settings, the manipulation of pH has been shown to directly influence the yield during fungal production of bioethanol (Lin et al.2012; Mohd Azhar et al.2017) and the power density in microbial fuel cells (Ganguli and Dunn 2012). Despite the paramount importance of pH, it has, to the best of our knowledge, never been monitored directly inside the extracellular matrix of fungal biofilms, but only in the surrounding bulk fluid. Measurements of pH in the matrix of bacterial biofilms, using either microelectrodes (Schreiber et al.2010; Von Ohle et al.2010) or confocal microscopy based techniques (Schlafer and Meyer 2016), have demonstrated the presence of steep pH gradients and distinct microenvironments, contributing to a deeper understanding of biofilm metabolism (Xiao et al.2012; Dige et al.2016). We have recently developed a confocal microscopy based technique that exploits the staining properties and pH-dependent emission shift of the ratiometric dye C-SNARF-4 to map pH developments in bacterial biofilms (Schlafer et al.2015). Compared to other quantitative confocal microscopy based approaches, pH ratiometry with C-SNARF-4 offers certain advantages. Due to the pH-dependent emission shift of the dye, pH can be determined irrespective of the concentration of the dye (Marcotte and Brouwer 2005). Moreover, C-SNARF-4 penetrates biofilms easily and quickly, which has proven to be problematic when pH sensitive dyes are immobilised on nanoparticles (Hidalgo et al.2009). Finally, C-SNARF-4 can be employed in bacterial biofilms without the addition of a second dye, as it stains bacterial cells and the surrounding biofilm matrix with different fluorescence intensity levels. The latter property allows for an intensity threshold based identification of matrix areas in the confocal images, which is crucial, as the matrix pH differs from intracellular pH due to bacterial homeostasis (Schlafer and Meyer 2016). In summary, C-SNARF-4-based pH ratiometry allows monitoring pH developments in the matrix of bacterial biofilms in real-time, in all three dimensions, but it has never been applied to fungal biofilms. As the fungal cell wall structure differs fundamentally from the one of bacteria (Bowman and Free 2006), the aims of the present study were to (i) investigate the staining properties of C-SNARF-4 on fungi, using the well-studied A. fumigatus, C. albicans, C. dubliniensis, C. neoformans and S. cerevisiae; and to (ii) develop a procedure for reliable confocal microscopy based quantification of extracellular pH in biofilms formed by these species. MATERIALS AND METHODS Fungal strains Aspergillus fumigatus (DSM 790), Candida albicans (NCPF 3179), Candida dubliniensis (DSM 13 268), Cryptococcus neoformans (DSM 6972) and Saccharomyces cerevisiae (DSM 3799) were cultivated aerobically on Sabouraud dextrose agar containing penicillin and streptomycin (Thermo Fisher Scientific, Roskilde, Denmark) at 35°C. A. fumigatus was harvested from the agar plates by flooding the surface with 0.9% sterile NaCl containing 0.05% (v/v) Tween 80 (Sigma-Aldrich, Brøndby, Denmark) according to the protocol described by Shirazi et al. (2016), while all other strains were harvested with inoculation loops. Prior to experimental use, the strains were transferred to Brain Heart Infusion (Sigma-Aldrich, Brøndby, Denmark) and grown until late exponential phase (18–24 h) under aerobic conditions at 35°C. Visualisation of immobilised planktonic fungi with C-SNARF-4 To test if C-SNARF-4 visualised all employed fungal strains, the staining properties of C-SNARF-4 were compared to BacLight. Planktonic cultures were washed in 0.9% sterile NaCl and adjusted to an optical density of 0.2 (550 nm). HEPES buffer solutions (50 mM) were used to titrate the pH of the cultures to 4.5–8, in steps of 0.5 pH units. A. fumigatus, C. dubliniensis, C. neoformans and S. cerevisiae were set to settle for 1 h in optical-bottom 96-well plates (ibidi, Planegg/Martinsried, Germany) coated with porcine gelatin (Type A; Sigma-Aldrich, Brøndby, Denmark; 0.2% w V−1 in Milli-Q water). C. albicans did not attach to gelatin-coated surfaces and was therefore immobilised in Concanavalin A (Sigma-Aldrich, Brøndby, Denmark) coated 96-well plates. To remove loosely adherent cells, the wells were washed twice with HEPES buffer of the same pH. Then C-SNARF-4 (Life Technologies, Nærum, Denmark) was added to a concentration of 30 μM and the cells were imaged with confocal laser scanning microscopy (CLSM). Thereafter, the fungi were counterstained with BacLight (Thermo Fisher Scientific, Roskilde, Denmark) according to the manufacturer's instructions, and images were acquired in the same microscopic fields of view. The experiments were carried out in duplicate. Visualisation of fungi in biofilms with C-SNARF-4 Biofilm growth of A. fumigatus, C. albicans, C. dubliniensis, C. neoformans and S. cerevisiae was performed modifying published protocols (Ramage et al.2001a; Ramage et al.2001b; Ravi et al.2009; Shirazi et al.2016). Planktonic cultures of the employed strains were washed in phosphate-buffered saline (PBS; pH 7.4; Sigma-Aldrich¸ Brøndby, Denmark) and adjusted to optical densities (550 nm) of 0.05 (A. fumigatus, C. dubliniensis, C. neoformans and S. cerevisiae) or 0.5 (C. albicans and S. cerevisiae). Optical bottom 96-well plates (ibidi, Planegg/Martinsried, Germany) were coated for 30 min at 35°C with sterile saliva prepared according to the method of de Jong et al. (1984), and 100 μl of the planktonic suspensions was set to settle for 1.5 h at 35°C. Thereafter, the wells were washed twice with PBS to remove non-adherent cells, and 200 μl of Yeast Nitrogen Base (Sigma-Aldrich, Brøndby, Denmark) containing 100 mM glucose was added. Biofilm growth was carried out at 35°C under aerobic conditions for 24 h, after which stable biofilms had grown for all employed species except S. cerevisiae. We performed additional experiments with S. cerevisiae to test if biofilm growth occurred in the absence of a salivary coating or using other cell concentrations in the initial inoculum, but none of the setups yielded stable biofilms. Saccharomyces cerevisiae was therefore excluded from subsequent experiments. After biofilm growth, all wells were washed three times with HEPES buffer solutions (50 mM) titrated to pH 4.5–8 in steps of 0.5 pH units, and C-SNARF-4 was added to a concentration of 30 μM. CLSM images were acquired in different microscopic fields of view for each strain and pH, the wells were counterstained with BacLight, and identical fields of view were imaged again. Experiments were performed in duplicate for all strains and pH values. Monitoring of pH in fungal biofilms As a proof of concept, we monitored pH developments in monospecies biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans following exposure to glucose. For each strain, triplicate biofilms were grown, washed twice with sterile 0.9% NaCl and then incubated for 1 h with 0.9% NaCl containing C-SNARF-4 (30 μM) and glucose (0.4% w/v). In each biofilm, CLSM images were acquired 5 μm above the biofilm–substratum interface in three different microscopic fields of view. 5, 10, 15, 20, 30 and 60 min after the addition of glucose extracellular pH was determined as described below. Additionally, Z-stacks were recorded in selected fields of view to record vertical pH profiles of the biofilms. The experiments were performed in duplicate. Confocal laser scanning microscopy Confocal laser scanning microscopy (CSLM) All images were acquired with a Zeiss LSM 510 META (Zeiss, Jena, Germany) equipped with a 63× oil immersion objective (Plan-Apochromat; NA = 1.4). C-SNARF-4 was excited at 543 nm and detected from 576 to 608 nm (green channel) and 629 to 661 nm (red channel; Marcotte and Brouwer 2005). SYTO 9 and propidium iodide, the components of BacLight, were excited at 488 nm and 543 nm, respectively, and detected from 500 to 554 nm and 554 to 608 nm. The pinhole was set to 2 Airy Units (optical slice 1.6 μm), the images were 2048 × 2048 pixels in size (143 × 143 μm) and acquired with a pixel dwell time of 1.6 μs, line average 4. Calibration of C-SNARF-4 For pH analysis, the ratios of the fluorescence intensities of C-SNARF-4 in the two different detection windows are calculated and converted into pH values based on a calibration curve. The calibration was performed using HEPES buffer solutions (50 mM) containing C-SNARF-4 (30 μM), which were titrated to pH 3.0–8.5 and imaged with the same laser settings used for biofilm image acquisition. After each buffer image, an image with the laser turned off was taken to correct for detector offset. Calibration images were taken in triplicate in different microscopic fields of view; the means of the ratios were plotted against the pH of the buffer solutions and fitted to the following function, assuming a five-parameter logistic curve (MyCurveFit; MyAssays Ltd, Brighton, UK): \begin{eqnarray} pH = \left[ {{{\left( {{{\left( {\frac{{2.9371815}}{{Ratio - 0.1000935}}} \right)}^{\frac{1}{{5298906}}}} - 1} \right)}^{\frac{1}{{6.649332}}}}} \right] \times 61.95654 \end{eqnarray} (1) Calibration data is shown in Fig. S1, Supporting Information. Digital image analysis Both channels (green and red) of the CLSM images of the C-SNARF-4-stained biofilms were exported separately as TIF files into the software daime (Daims, Lücker and Wagner 2006). To remove all fungal cells from the biofilm images, the images of the brightest channel were segmented with both a lower and an upper brightness threshold. With the lower threshold, the intracellular space of viable cells was removed from the images. The higher threshold removed the fungal cell walls and the cytoplasm of membrane-compromised cells from the images, leaving only the extracellular space for subsequent pH analysis. The object layer of the segmented images was then transferred to the corresponding images of the other channel, and both image series were exported into ImageJ (Schneider, Rasband and Eliceiri 2012). In ImageJ, the mean filter (radius: 1 pixel) was applied for noise reduction, the green channel images were divided by the red channel images and the average ratios and standard deviations were calculated for all microscopic fields of view. Equation (1) was used to translate the ratios into pH values, and false colouring was applied for graphic representation. The digital image analysis procedure is presented in more detail in Schlafer and Dige (2016). RESULTS C-SNARF-4 stained all five employed fungal strains with an identical pattern. At all tested pH values, the dye bound strongly to the cell wall of immobilised viable cells but left the cytoplasm unstained. In membrane-compromised cells, C-SNARF-4 accumulated in the cytoplasm. Counterstaining with BacLight showed that all cells were targeted by C-SNARF-4 (Figs S2 and S3, Supporting Information). Likewise, C-SNARF-4 stained the walls of all cells, including hyphae, in single-species biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans, irrespective of the pH (Fig. 1). This staining pattern resulted in three fluorescence intensity levels in all acquired biofilm images, illustrated in Fig. 2 for C. dubliniensis (Figs 2A–E) and A. fumigatus (Figs 2F–J): In both channels (green and red), the cytoplasm of viable cells showed the lowest intensity, while the cell walls of all cells and the cytoplasm of membrane-compromised cells showed the highest intensity. The surrounding biofilm matrix displayed an intermediate level of fluorescence (Figs 2A–C, F–H). The staining pattern allowed for an intensity threshold based image analysis to identify and remove all fungal cells from the biofilm images. Images were segmented with both a lower threshold to remove the cytoplasm of viable cells and with a higher threshold to remove membrane-compromised cells and cell walls. Thereafter, only the biofilm matrix was left for pH analysis (Fig. 2D and I). Green/red fluorescence ratios were calculated and false colouring was applied for visualisation purposes (Fig. 2E and J). Figure 1. View largeDownload slide Visualisation of fungal biofilms with C-SNARF-4. Biofilms of Candida dubliniensis (A, B, E, F), Candida albicans (C, D, G, H), Cryptococcus neoformans (I, J, M, N) and Aspergillus fumigatus (K, L, O, P) were stained with C-SNARF-4 at different pH values, imaged with a confocal microscope (A–D and I–L), counterstained with BacLight and imaged again (E–H and M–P). At all investigated pH values, C-SNARF-4 stained the walls of all fungal cells in the samples, including hyphens, as shown by counterstaining with BacLight. The fungal cytoplasm remained unstained, except when cells were membrane-compromised (yellow/red cells in panels C, D, I, J and L). pH values are indicated in the C-SNARF-4 images. Bars = 20 μm. Figure 1. View largeDownload slide Visualisation of fungal biofilms with C-SNARF-4. Biofilms of Candida dubliniensis (A, B, E, F), Candida albicans (C, D, G, H), Cryptococcus neoformans (I, J, M, N) and Aspergillus fumigatus (K, L, O, P) were stained with C-SNARF-4 at different pH values, imaged with a confocal microscope (A–D and I–L), counterstained with BacLight and imaged again (E–H and M–P). At all investigated pH values, C-SNARF-4 stained the walls of all fungal cells in the samples, including hyphens, as shown by counterstaining with BacLight. The fungal cytoplasm remained unstained, except when cells were membrane-compromised (yellow/red cells in panels C, D, I, J and L). pH values are indicated in the C-SNARF-4 images. Bars = 20 μm. Figure 2. View largeDownload slide Digital image analysis procedure for ratiometric determination of extracellular pH in fungal biofilms. Biofilms of Candida dubliniensis (A–E) and Aspergillus fumigatus (F–J) were stained with C-SNARF-4 in the presence of glucose. In both detection channels (green: A and F; red: B and G), the cytoplasm of viable cells displays the lowest fluorescence intensity, while the cell walls as well as the cytoplasm of membrane compromised cells (bright cell in A–C) show the highest intensity. Fluorescence deriving from the extracellular matrix shows an intermediate level of intensity. Panels C and H show overlays of the green and red channels. Based on intensity thresholding with both a lower and a higher threshold, all cells can be removed from the images, leaving only the biofilm matrix for pH analysis (D and I). The ratios of fluorescence intensities are calculated, converted to pH values, and in E and J, false colours were applied to visualise pH in the matrix. After 30 min of exposure to glucose, average pH in the selected fields of view was 5.9 for C. dubliniensis and 6.9 for A. fumigatus. Bars = 20 μm. Figure 2. View largeDownload slide Digital image analysis procedure for ratiometric determination of extracellular pH in fungal biofilms. Biofilms of Candida dubliniensis (A–E) and Aspergillus fumigatus (F–J) were stained with C-SNARF-4 in the presence of glucose. In both detection channels (green: A and F; red: B and G), the cytoplasm of viable cells displays the lowest fluorescence intensity, while the cell walls as well as the cytoplasm of membrane compromised cells (bright cell in A–C) show the highest intensity. Fluorescence deriving from the extracellular matrix shows an intermediate level of intensity. Panels C and H show overlays of the green and red channels. Based on intensity thresholding with both a lower and a higher threshold, all cells can be removed from the images, leaving only the biofilm matrix for pH analysis (D and I). The ratios of fluorescence intensities are calculated, converted to pH values, and in E and J, false colours were applied to visualise pH in the matrix. After 30 min of exposure to glucose, average pH in the selected fields of view was 5.9 for C. dubliniensis and 6.9 for A. fumigatus. Bars = 20 μm. As a proof of concept, biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans were incubated with 0.4% glucose, and the extracellular pH at the bottom of the biofilms, 5 μm above the biofilm–substratum interface, was monitored for 1 h. The pH development in the biofilms differed markedly between the strains. C. albicans biofilms showed the lowest pH, typically around 5 (Fig. 3; Fig. S4A, Supporting Information), which corresponds to the pH levels reached in planktonic suspension upon exposure to glucose (data not shown). Interestingly, these low pH levels were reached much faster than in planktonic culture (pH 5 after 7 h of incubation), typically within 5 min after exposure to glucose (Fig. S5A, Supporting Information). Acid production in C. dubliniensis biofilms occurred more slowly, but after 1 h of exposure to glucose, pH levels approached 5–5.5, too (Fig. 3; Fig S4B, Supporting Information). Under the chosen experimental conditions, the pH in A. fumigatus and C. neoformans biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h (Fig. 3; Fig. S4C and D, Supporting Information). In general, pH differences between replicate biofilms were more pronounced than between different fields of view within the same biofilms (Fig. S4, Supporting Information). Under the chosen conditions, only slight differences between different areas within one biofilm could be observed. Likewise, vertical pH profiles showed a constant pH across the slim biofilms (ca. 20 μm) at a given time (see Fig. S5B, Supporting Information, for examples). Figure 3. View largeDownload slide Typical pH development in biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans after incubation with glucose. In each biofilm, extracellular pH was measured ratiometrically 5 μm above the biofilm–substratum interface in three different microscopic fields of view. Each line represents the pH development in one field of view in the course of 1 h. pH dropped rapidly (within 5 min) from 7 to 5 in C. albicans biofilms (blue lines) and remained stable afterwards. In C. dubliniensis biofilms (grey/black lines), pH dropped more slowly and approached levels of 5.5 at the end of the incubation period. pH in A. fumigatus (green lines) and C. neoformans (red lines) biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h. Error bars = standard deviations. Figure 3. View largeDownload slide Typical pH development in biofilms of Aspergillus fumigatus, Candida albicans, Candida dubliniensis and Cryptococcus neoformans after incubation with glucose. In each biofilm, extracellular pH was measured ratiometrically 5 μm above the biofilm–substratum interface in three different microscopic fields of view. Each line represents the pH development in one field of view in the course of 1 h. pH dropped rapidly (within 5 min) from 7 to 5 in C. albicans biofilms (blue lines) and remained stable afterwards. In C. dubliniensis biofilms (grey/black lines), pH dropped more slowly and approached levels of 5.5 at the end of the incubation period. pH in A. fumigatus (green lines) and C. neoformans (red lines) biofilms stayed near neutral, and only slight changes could be observed in the course of 1 h. Error bars = standard deviations. DISCUSSION pH in fungal biofilms is of fundamental importance for both medical conditions and industrial applications. The present study is the first to describe a methodology to monitor pH in the matrix of fungal biofilms, using a combination of confocal microscopy with the ratiometric dye C-SNARF-4, and digital image post-processing. Unlike electrode measurements in bulk fluid, it provides not only a good temporal, but also a high spatial resolution. As a proof of concept, we monitored pH over time in monospecies biofilms of A. fumigatus, C. albicans, C. dubliniensis and C. neoformans, exposed to glucose. Of the employed strains, C. albicans was by far the most acidogenic under the chosen conditions. pH inside the biofilms dropped from near neutral to values around five within less than 5 min (Fig. 3; Figs S4 and S5, Supporting Information), which is much faster than the pH drops reported in planktonic culture (Samaranayake et al.1986) and may be explained by the high local cell density inside the biofilms and the modulating effect of the biofilm matrix (Koo, Falsetta and Klein 2013). Rapid pH drops inside fungal biofilms may be of particular importance for antifungal therapy, as the efficacy of many antifungal drugs decreases at low pH (Liu et al.2011; Danby et al.2012). Acid production in biofilms of C. dubliniensis occurred much slower; the pH typically still dropped after half an hour of exposure to glucose, reaching levels of 5.2–6.2 (Fig. 3; Fig. S4, Supporting Information). Although most cells of C. dubliniensis were still viable after a short (<30 min) exposure to low pH (Figs S2 E–G), the low acidogenicity, compared to C. albicans, may in part explain the predominance of C. albicans in infections associated with low pH, such as denture stomatitis (Gendreau and Loewy 2011). In biofilms of A. fumigatus and C. neoformans, pH stayed in the near-neutral range under the chosen conditions, although, and this holds true for all investigated species, notable differences could be observed between replicate biofilms (Fig. 3; Fig. S4, Supporting Information). In contrast, only minor differences in pH were observed between different microscopic fields of view within the same biofilm, irrespective of the species. This homogenous behaviour may be attributed to the age of the biofilms (24 h), which were still in the maturation phase by the time pH was recorded, and, in particular, to the fact that the present study employed monospecies biofilms. Ratiometric pH measurements in the matrix of young dental biofilms, consisting of several different bacterial species, have revealed considerably more variation in pH within one biofilm specimen (Dige et al.2016). Studying pH developments in multispecies fungal biofilms is therefore of great interest, but beyond the scope of the present investigation. The staining pattern of C-SNARF-4 in biofilms was identical for all strains employed in this study: membrane-compromised cells and the walls of all cells showed a brighter fluorescence than the biofilm matrix, whereas the cytoplasm of viable cells remained unstained. These different levels of fluorescence intensity allowed for a reliable removal of fungal cells from the confocal images and thus the determination of extracellular pH only (Fig. 2). As bacterial cells have been shown to up-concentrate C-SNARF-4 irrespective of their viability status (Schlafer et al.2015), the same image analysis procedure will also be applicable to cross-kingdom biofilms. The strains employed in the present work all belong to the phyla Basidiomycota (C. neoformans) or Ascomycota (A. fumigatus, C. albicans, C. dubliniensis and Saccharomyces cerevisiae) and were selected because of their relevance in the medical field and/or industrial applications. Although S. cerevisiae did not form biofilms under the chosen conditions, it is likely that pH in S. cerevisiae biofilms can be monitored using C-SNARF-4, as the staining pattern of immobilised cells was identical to the one observed for the other species (Fig. S3C and D, Supporting Information). However, given the considerable differences in the fungal cell wall structure, it still remains to be investigated if the presented methodology is applicable to all biofilm forming fungi. The use of confocal microscopy imposes some limitations on C-SNARF-4-based pH ratiometry. While pH at the biofilm–substratum interface can be recorded irrespective of the dimensions of the biofilm, only thin biofilms with a thickness of up to 50 μm can be imaged entirely. If vertical gradients in thicker biofilms are to be assessed, motor-controlled pH microelectrodes may be used (Von Ohle et al.2010). While the image post-processing necessary for pH ratiometry takes time, image acquisition is rather fast. In the present study, an acquisition time of approximately 1 min was chosen, but if need be, scanning parameters can be adjusted accordingly and images can be acquired in less than 10 s, at the expense of detail. With a pka of ∼6.4, C-SNARF-4 is suitable for pH determination in both acidic and moderately alkaline environments. At very low (<4) and at high (>8) pH, the discriminative power gets lower, as can be seen from the resulting bigger error bars (Fig. 3; Fig. S5, Supporting Information). In conclusion, the present work provides a new tool for the confocal microscopy based determination of extracellular pH inside fungal biofilms. pH developments can be monitored in real-time, in all three dimensions, in the pH range between 4 and 8. pH ratiometry with C-SNARF-4 can contribute to a more thorough understanding of fungal biofilm metabolism, which may be important for the treatment of fungal diseases and the optimisation of fungal biofilm-based industrial processes. SUPPLEMENTARY DATA Supplementary data are available at FEMSYR online. Acknowledgements Mette Nikolajsen is acknowledged for excellent technical support. FUNDING This work was supported by AIAS, Aarhus Institute of Advanced Studies, Aarhus University and Aarhus University Research Foundation (AUFF). Conflicts of interest. None declared. 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FEMS Yeast ResearchOxford University Press

Published: Apr 19, 2018

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