Visualizing biomaterial degradation by Candida albicans using

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Visualizing biomaterial degradation by Candida albicans using embedded luminescent molecules to report on substrate digestion and cellular uptake of hydrolysate Bryan Robert Coad, Thomas Danny Michl, Christie Ann Bader, Joris Baranger, Carla Giles, Giovanna Cufaro Goncalves, Pratiti Nath, Stephanie Jade Lamont-Friedrich, Malin Johnsson, Hans J Griesser, and Sally E Plush ACS Appl. Bio Mater., Just Accepted Manuscript • DOI: 10.1021/acsabm.9b00520 • Publication Date (Web): 15 Aug 2019 Downloaded from pubs.acs.org on August 20, 2019

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ACS Applied Bio Materials

Visualizing biomaterial degradation by Candida albicans using embedded luminescent molecules to report on substrate digestion and cellular uptake of hydrolysate Bryan R. Coad,1,2 * Thomas D. Michl,1 Christie A. Bader,3 Joris Baranger,1 Carla Giles,1,4 Giovanna Cufaro Gonçalves,1 Pratiti Nath,1 Stephanie J. Lamont-Friedrich,1 Malin Johnsson,1 Hans J. Griesser,1 Sally E. Plush3

1. Future Industries Institute, University of South Australia, Mawson Lakes, South Australia, Australia 5095

2. School of Agriculture, Food & Wine. University of Adelaide. Adelaide, Australia 5000

3. School of Pharmacy and Medical Sciences, University of South Australia, Adelaide, South Australia, Australia 5000

ACS Paragon Plus Environment

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ACS Applied Bio Materials 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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4. Centre for Aquatic Animal Health & Vaccines. Tasmania Department of Primary Industries Parks Water & Environment, 165 Westbury Road, Prospect, Tasmania, Australia, 7250

Corresponding Author * [email protected]

Keywords: Surface coating; biodegradable; Candida albicans; fungus; hydrolysis; pathogen

Abstract

Microbial pathogens use hydrolases as a virulence strategy to spread disease through tissues and colonize medical device surfaces; however, visualizing this process is a


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technically challenging problem. In order to understand better the role of secreted fungal hydrolases and their role in Candida albicans virulence, we developed an in situ model system using luminescent Re(I) and Ir(III) containing probe molecules embedded in a biodegradable (poly(lactic-co-glycolic acid), PLGA) polymer, and tracked their uptake using epifluorescent imaging. We found that secretion of esterases can explain how physically embedded probes are acquired by fungal cells through the degradation of PLGA since embedded probes could not be liberated from non-biodegradable polystyrene (PS). It was important to verify that epifluorescent imaging captured the fate of probe molecules rather than naturally occurring fungal autofluorescence. For this, we exploited the intense luminescent signals and long spectral relaxation times of the Re and Ir containing probe molecules, resolved in time using a gated imaging system. Results provide a visual demonstration of a key virulence trait of C. albicans: the use of hydrolases as a means to degrade materials and acquire hydrolysis products during fungal growth and hyphal development. These results help to explain the role of non-specific hydrolases using a degradable material that is relevant to the study of fungal pathogenesis on biotic (tissues) surfaces. Additionally, understanding how fungal pathogens condition surfaces


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by using non-specific hydrolases is important to the study of fungal attachment on abiotic surfaces, the first step in biofilm formation on medical devices.


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Introduction Pathogen colonization of medical device surfaces presents a serious health concern.13

While bacterial and viral pathogens are generally recognized for their immense adverse

effects on human health, there is increasing concern about fungal pathogens (yeasts and molds). Fungi possess remarkable survival strategies such as being able to switch to forms that are more virulent and the secretion of degradative enzymes.4-7 Within and on the body, this latter trait allows the fungus to degrade biopolymers such as host tissues in order to invade deeper and thus disseminate infection, and subsequently acquire hydrolysis products as nutrients to sustain fungal growth.8-9 On abiotic materials such as those used to fabricate medical implant devices (especially plastics), enzymatic attack can lead to their degradation10 and an increase in nano-scale roughness.11 Medical device surfaces which have been physically etched by the action of secreted proteases provide an opportunity for strong surface adherence by pathogens followed by biofilm production – conditions which contribute to recalcitrant infections that necessitate the removal of contaminated devices.12 Thus, understanding how fungi use degradative


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enzymes to colonize surfaces (both biotic and abiotic) is fundamental to the understanding of pathogens, and their spread and colonization of surfaces. Model materials can be used in the study of how pathogens use secreted enzymes to degrade biopolymers.13 This is illustrated by reports on so-called self-defensive materials where antibiotics are absorbed into a biodegradable matrix that is then subjected to degradation and release caused by the action of microbial enzymes.14-21 While bacterial inhibition on such surfaces has been demonstrated, it is not clear whether their release was caused by the action of matrix-degrading enzymes or at least partly contributed by elution of the antibiotic from the substrate. Other approaches have used covalent bond formation to link antibiotics to the matrix on enzyme-labile linkers. Cado et al. created a polyelectrolyte multilayer substrate coating based on hyaluronic acid and chitosan where the antimicrobial peptide cateslytin was covalently bound to the matrix.22 Here, release of the antibiotic was expected to be caused by secreted hyaluronidase and/or chitosanase from the yeast Candida albicans. While these enzymes are substrate specific, an interesting question is what role do non-substrate specific enzymes like esterases play in microbial hydrolysis. Thus a study that uses biodegradable ester-linked substrates could


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be more widely relevant both as a model of polyester medical devices degradation and to model microbial hydrolysis of ester bonds which would be necessary in a host for lipid hydrolysis and colonization of fatty tissues. While hydrolysis causes a partial breakdown of substrate material, few studies have investigated the fate of the hydrolyzed matrix. Acquisition of digested materials is a driver of microbial virulence: it provides a source of small molecule building blocks that sustain pathogen growth.23-24 Thus, we are interested in visualizing the fate of the liberated molecules and their eventual uptake by the organism as a way of further understanding microbial pathogenesis on surfaces. In this study, we tested the hypothesis whether fungal pathogens can use nonspecific esterases to degrade a polymeric matrix and take up the hydrolysate. To prove this point, it was necessary to 1) prove that luminescent probes molecules embedded within a digestible matrix could only be liberated by the presence of fungi (i.e. not through spontaneous elution), 2) prove that emission from luminescent probe molecules could not be confused with natural autofluorescence produced by the organism itself and 3)


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demonstrate that the hydrolysate is concentrated within the cell compared to its surroundings. The general schematic for the experiment is shown in Figure 1.

Figure 1. Schematic illustration of the materials and biological experiments. A) Spin casting was used to incorporate Re or Ir probe molecules in a polymer matrix that was non-biodegradable (polystyrene) or biodegradable (poly(lactic-co-glycolic acid)). B)

Candida albicans grown on biodegradable surface coatings was able to digest the matrix and acquire the probe molecules intracellularly.

To prove this concept we produced robust biodegradable polymer films using spin casting of biodegradable poly(lactic-co-glycolic acid) (PLGA) and used surface analysis


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to confirm its physical and chemical properties. The non-biodegradable control was polystyrene (PS). We used luminescent Re(I) or Ir(III) containing organometallic dyes to probe the degradation of the matrix and visualize their uptake by the pathogen. These probes were chosen for their long-lived emission and low levels of photobleaching that provided a means by which we could discount fungal autofluorescence. Materials and methods

Reagents and supplies Polystyrene (Mw = 280,000 Da, PS), poly(DL-lactide-co-glycolide, Mw = 50,000 75,000 Da, PLGA), phosphate-buffered saline tablets, NANG (no amino acid, no glucose) yeast nitrogen base medium, nutrient broth, d-mannitol, K2HPO4 and cleaning surfactant RBS 25% were obtained from Sigma-Aldrich. ReZolve-ER and IraZolve-L1 were provided by ReZolve Scientific Pty Ltd (Adelaide, Australia). Ethyl acetate AR (99.5 %), ethanol undenatured 100 %, hydrochloric acid (36 %) and acetone AR were purchased from Chem-Supply Pty Ltd. Spider medium was prepared according to previous reports and titrated to pH 7 prior to autoclaving.25 All other microbiological solutions were likewise autoclaved prior to use. Candida albicans SC5314 was provided by the National


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Mycology Reference Centre, SA Pathology, Adelaide. Silicon wafers were purchased from MMRC. Coverglass was obtained from ProSciTech (Ø12 mm). All water used was purified (Milli-Q).

Substrate preparation Glass coverslips were used for the epifluorescence microscopy studies, whereas silicon wafers were used for surface analysis (XPS, TOF-SIMS and ellipsometry). Glass coverslips were used as is, whereas silicon wafers were cleaned prior to use. Silicon wafers were cut into 1 cm squares using a Disco DAD 321 automatic dicing saw, and then washed to remove any surface residues. Wafers were soaked in a 5% RBS solution for 30 min. at 70°C. Then, wafers were washed three times with distilled water and soaked in a 1% hydrochloric acid (HCl) solution. After one hr., the wafers were washed three times with distilled water. Finally, the wafers were sonicated in ethanol for five minutes and then sonicated in acetone for five minutes. The clean silicon substrates were stored in acetone prior to use to keep them clean. Spin coating


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PLGA and PS solutions were made by stirring the polymers (PS or PLGA) in ethyl acetate at 25°C to make up a concentration of 1.75 % (w/v). Fluorescent dyes ReZolveER, and IraZolve-L1 were incorporated into the PLGA and PS solutions at a concentration of 100 µM. Polymer coatings were deposited on glass coverslips or silicon wafers using a WS650MZ-23NPPB spin coating device (Laurell Technologies Corporation). Aliquots of PLGA or PS solutions (100µL) were deposited on the center of the substrate, and then the substrate was accelerated at 500 rpm to a final speed of 2000 rpm and held for 1 min, permitting the coatings to be deposited uniformly.

Surface analysis The thicknesses of deposited films was determined using a variable angle spectroscopic ellipsometer (J.A. Woollam M-2000DI). Analyses were conducted over the wavelength range 400 - 1000 nm at angles of 60°, 65°, 70° and 75° each for 10 seconds. The analysis was performed assuming the native oxide layer to be 1.49 nm and fitting the plasma polymers with a Cauchy layer with Urbach extension.

ICP-MS


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ICP-MS was used to detect whether Re or Ir was leaching from probe-embedded coatings. Spin coated films were made on glass coverslips using the two model materials (PLGA and PS) containing either probes. Then, the samples were soaked in 2 mL of phosphate buffer saline (PBS). After the pre-determined soaking time, the supernatant was removed and kept in a sealed vial until analysis. This supernatant was then analyzed in triplicate by ICP-MS to determine the Re or Ir concentration. This was measured using an 8800 Triple Quadrupole ICP-MS. Before the experiment, calibration was conducted with known 1 ppb standards of Re, Ir, Li, Y, Tl, Ce and Co in 2 % HNO3.

XPS A Kratos Axis Ultra with Delay Line Detector (DLD) was used to record XP spectra with a monochromatic Al Kα radiation source operating at 225 W. The elements present on the surface were identified from a survey spectrum recorded over the binding energy range from 0-1000 eV at a pass energy of 160 eV and energy steps of 0.5 eV. Binding energies were referenced to the aliphatic C 1s carbon peak at 285 eV to compensate for surface charging. The analysis area was circular and 3 mm in diameter. Processing and


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component fitting of the high-resolution spectra was performed using CasaXPS (Neal Fairley, U.K.).

Time of Flight Secondary Ion Mass Spectrometry (ToF SIMS) ToF SIMS measurements were performed with a TRIFT V NanoToF instrument (Physical Electronics Inc., Chanhassen, MN, USA). The analysis gun was a liquid metal 197Au+

Ion Gun (LMIG) operated at 30 kV to sputter and ionize species from each sample

surface. The LMIG was operated in Au1 “bunched” mode to optimize mass resolution. A dual beam charge neutralization system using low energy argon ions (up to 10 eV) and electrons (up to 25 eV) was employed to provide charge neutralization performance. Experiments were carried out under a vacuum of 5 x 10-6 Pa. Data were collected in the positive SIMS mode at least three different locations, typically using a 100 x 100 µm raster area. Positive mass axis calibration was conducted with CH3+, C2H5+, and C3H7+. Each sample was characterized by six independently acquired positive ions mass spectra. All data were collected and interpreted with WinCadenceN software (ULVAC-PHI Inc.).

Microbiology methods


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Candida albicans SC5314 was prepared by streaking the fungi from a -80°C stock onto a Sabouraud agar (SAB) plate and incubated overnight at 37°C. Single colonies were picked and diluted to the same optical density as a 3.0 McFarland standard solution (approx. 9.0 x 108 colony forming units/mL). Isolates were the sub-cultured in Spider medium overnight. The solution was diluted ten-fold into NANG. Additional details are provided in the Supporting Information. To evaluate coatings, coated glass coverslips were placed into individual wells on a 24 well plate, then 1 mL of the fungal suspension was added to each well. Plates were then covered by aluminum foil and incubated at 37°C. Subsequently, each sample was imaged. Before taking images with the microscope, the supernatant was firstly removed, and then samples were washed gently three times with PBS.

Microscopy Spin coated substrates were initially imaged by confocal microscopy using a Nikon A1+ confocal microscope, fitted with a LU-N4/LU-N4S 4-laser unit (403, 488, 561 and 640 nm), an A1-DUG GaAsP Multi Detector Unit (2GaAsP PMTs + standard PMTs), and an A1-DUS Spectral Detector (Nikon). Spectral profiles were obtained using 403 nm


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excitation with emission collected across 32 channels between 410 nm and 750 nm. NIS Elements software (Nikon) was then used to generate spectral profiles of the substrates. Epifluorescent images of yeast grown on coated substrates were collected on an Olympus BH2 RFC microscope equipped with a 365 nm and a 565 nm visible light emitting diode (LED) (Thorlabs M365L2 LED and M565L3 LED) and broad band pass emission filter. Images were captured on a Kingfisher Scientific CCD 6MP cooled digital B/W camera (Raptor Photonics). To allow time gated imaging, this microscope was customized by Lastek with an optical chopper system (MC2000B-EC, Thorlabs) and LED drive with pulse modulation (DC2200, Thorlabs).26 Fluorescent images were acquired by exciting probes with a 365 nm LED and emission collection using a DAPI long band pass filter and a 200 ms exposure time. Time gated images were collected using pulse excitation with the 365 nm LED pulsing at 1 ms on 9 ms off and the chopper speed set to 20 Hz that allowed elimination of autofluorescence while maintaining probe fluorescence detection. Image processing was performed in FIJI software.27

Results and discussion


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Understanding how microbes colonize surfaces (biotic or abiotic) is fundamental to understanding the causes of disease. After adhesion, the secretion of hydrolases is important to the survival of pathogens as a way to acquire nutrients and disseminating infection throughout tissues through degradation of host-biopolymers.28-29 Hydrolysis of abiotic polymers (such as the materials comprising medical device surfaces) is also an effective strategy in infection because this process promotes surface conditioning which aids in attachment: the first step in biofilm formation.30-32 While the importance of hydrolases is recognized, studies of the interaction between pathogens and surfaces are few. One reason could be the technical challenge of finding suitable model materials that act as hydrolysable substrates, developing methods that convincingly report on their degradation and provide a means of their measurement. A powerful and informative approach is to visualize the microbial degradation of substrates containing reporter probes with the organism in-situ. This in-situ approach, however, is confounded by a major technical obstacle in fluorescence microscopy where it is challenging to distinguish probe signal from the organism’s own autofluorescence.


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We here report a strategy that has overcome these challenges, and allows for the study of in-situ degradation of a model material through a luminescent reporter molecule without interference from the microorganism’s native autofluorescence. Our study focused on the secretion of fungal lipases (esterases), an important virulence factor for the biomedically relevant Candida albicans.33-36 On animal tissues, this pathogen can penetrate tissues and become invasive, leading to high mortality.37 Furthermore, on implanted medical devices, C. albicans has been implicated in biofilm development and recalcitrant infections.38 We chose PLGA as the substrate material as a simple polymer linked with ester groups. It serves as a straightforward mimetic of other ester-linked (bio)polymers39 including polyesters, such as those used in medical device manufacture for sutures, vascular grafts and stents, just to name a few. PLGA also serves as a simple model substrate for studying hydrolysis of organic biomolecules such as lipids, by which we mean their ester-linked triglyceride bonds. Since C. albicans readily hydrolyses lipids and fatty acids and utilizes the resulting hydrolysis products as carbon sources, we anticipated the same fate for PLGA. In contrast, we wanted to challenge C. albicans with a control surface that is known to be de-facto non-biodegradable, polystyrene (PS).40


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Robust polymer films were fabricated and characterized using instrumental analysis Spin-cast PLGA and PS polymer films were reliably manufactured 100 to 140 nm thick as measured by spectroscopic ellipsometry (Figure S1). XPS analysis of PLGA and PS coatings produced survey spectra containing 66% C and 34% O for the former and 100% C for the latter (Figure S2), which is typical for a homogenous layer that exceeds the sampling depth of XPS. For samples containing organometallic probe molecules, XPS was not sensitive enough to observe trace amounts of metals within the coatings due to the low concentration of dyes used. We performed full ToF-SIMS characterization on the Re(I) embedded material and also evaluated the spectral properties of the film. For the two polymeric coatings without probes, spectra show characteristic C7H7+ peaks observed for the PS sample, and C2H3O+ and C3H4O+ peaks observed for PLGA (Figures S4a and S4b). For the Re(I) embedded material, ToF-SIMS revealed two peaks at m/z = 449 amu and 451 amu that were not present in the undoped polymer coating. This mass corresponds to a large Re-containing structural fragment, and further confirmed by the 2 amu peak splitting and peak heights which match the relative isotopic abundance of 185Re and 187Re atoms (Figure S4d).


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Experimentation with different probe concentrations showed acceptable spectral intensity was obtained from polymer films when 100 µM of the ReZolve-ER probe was used in the spin casting solution (Figure S4). Our initial experiments with yeasts used materials with the Re-based probe; however, we found that better images were produced but substituting the Ir(III) probe (IraZolve-L1) at the same concentration. It is clear from data that will be shown below, that using either the Re or Ir doped materials makes no difference in their physicocchemical properties, but only improves the resolution of yeasts in imaging experiments when using the Ir probe. For comparison sake, the emission intensity for the ReZolve-ER and IraZolve-L1 probes is presented in Figure S5.

The films and embedded probes were stable when soaked in aqueous solution To confirm the stability and non-leaching properties of the polymer films, samples of dye-loaded films were soaked in PBS buffer for 4 days. The soaking solution did not significantly change the thickness of the films, showing that they were physically unchanged (Figure S1). The soaking solution was analyzed by ICP-MS to determine whether any Re(I) or Ir(III) could be detected (Figure 2). The ICP-MS assay has a detection limit in the ppb range. For both PS and PLGA matrix, and regardless of the


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probe molecule, the detectable amounts were all less than 1.7 ppb. The data show no increasing trend, as might be expected if ions were accumulating in the soaking solution with time. This and the large relative errors and low absolute concentrations indicate that any leaching was at the limit of detection of this technique. These values show that the probes are not leached out of the coating in any appreciable concentration, and by this utility, were diagnostic for release only in the presence of biological organisms as shown below.


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Figure 2. Total concentration of embedded Re or Ir ions that could be potentially leached from PS or PLGA thin films over the course of 4 days’ soaking. Values are averages and error bars represent the standard deviation (n=3).

Time gated fluorescence microscopy eliminated the confounding effect of autofluorescence Fluorescent microscopy was used to visualize acquisition of polymer-embedded dye molecules by the fungus. However, when collecting fluorescent images, interference is


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caused by naturally occurring fluorescent molecules in C. albicans cells that can make definitive identification of the embedded dyes potentially obscure. To overcome this challenge, we used group 6 transition metal organometallic probes that possess a longlived emission spectrum on the order of 30 ms; much longer than natural fluorescence lifetimes in the nano/picosecond range. This difference in emission time can be used to distinguish between the two. A time gating system was used to block out signal from short-lived emissions and collect only the longer-lived signal stemming from the probes as schematically illustrated in Figure 3A.26 Figure 3B shows a weak green signal from naturally occurring autofluorescence can be blocked when time gating is “on”. In contrast, Figure 3C and D show that when Re or Ir-containing dyes were incorporated in the polymeric matrix and with the gate off, the images appeared bright, which was due to both the presence of the dye and a small contribution from autofluorescence. Finally, for these samples with the gate on, autofluorescence was eliminated showing only the signal originating from the probes. While we experimented with the Re-based ReZolve-ER probe (Figure 3C), and these provided evidence of uptake from biodegradable polymers, the most visually striking results were obtained using time gating with the Ir-based IraZolve-


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L1 probe (Figure 3D) which were both brighter and longer-lived. By combining the evidence that the dyes are not leaching out of the substrate (Figure 2) and that the microscope only acquires the dyes’ emitted light, not autofluorescence (Figure 3), we have demonstrated a method where any fluorescence observed within the organism must be due to the dyes that were taken up from the substrate. However, a consequence of blocking the most intense emission and collecting a much weaker, time-resolved and decaying signal from the probe is that images tend to have a lower signal to noise ratio and can appear grainy.

Figure 3. (A) Schematic illustrating the imaging method in which samples were excited by a 365 nm pulse LED for 1 ms and no excitation was performed for 9 ms. When time gating


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was on, a chopper was used to prevent initial natural fluorescence reaching the camera. (B) Micrographs illustrate that the yeast and PLGA polymer have a faint autofluorescent signal when gating was not used (gate off), but could be removed by gating this emission (gate on). Using materials embedded with ReZolve-ER dye (C) the IraZolve-L1 dye (D) provided long-lived emission. Thus, when gating was on, the fluorescent signal was still bright, and not interfered by autofluorescence. Note that the probes are initially distributed throughout the polymer film substrate, which is why both the organisms and the background produce a fluorescent signal.

Microbial growth on surfaces led to increased substrate degradation Next, we studied the ability of C. albicans to degrade the coatings over the course of several days’ growth. C. albicans is capable of using a wide variety of organic molecules as a source of carbon and can therefore use hydrolysable materials as a potential source of nutrition.41 When provided with minimal growth medium (NANG: no amino acids, no glucose) and exposed to biodegradable polymers such as PLGA, we observed that C.

albicans adheres strongly to the material’s surface and attempts to digest it. This process


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was visualized as the uptake of the luminescent dye after growth on the surface after 1, 2, 4 and 6 days. However, the earliest time point when we could clearly identify a luminescent signal in the yeast was after 2 days growth (Figure 4). At shorter time points, an insufficient amount of dye was present in the cells meaning that outlines of the yeast could not be clearly distinguished from signal originating from the polymeric matrix itself. The rate of accumulation is illustrated by comparing images at days 2 and 4, where the latter time point shows that the cells become brighter owing to increased growth on the surface. After 2 days’ growth, dye accumulation was sufficient to show concentration in the periphery of fungal colonies. By day 4, this was shown to be evenly distributed in the interior of the yeast cells. Thus, as the fungus grew and digested the substrate over time, the accumulated dye concentrated within the cell and increased in brightness, providing a signal that could gradually be distinguished from the background.


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Figure 4. C. albicans grown for 2 days (upper) or 4 days (bottom) in NANG medium on PLGA substrate with embedded IraZolve L1. Representative micrographs using the time gating procedure show many yeast forms, and individual colonies preferring to cluster


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together in transmitted light images (left). Few hyphal filaments are present. Scale bars = 10 µm.

C. albicans was able to degrade the PLGA polymers but not polystyrene After 6 days’ growth on PLGA and PS plastic surfaces, C. albicans continued to grow and began to develop some pseudohyphal filaments as shown by transmitted light images (Figure 5). When the Ir-based dye was incorporated in the PLGA sample, fluorescent imaging and time gating showed the dye to be taken up by the fungal cells. However, no fluorescent signal was observed from the PS coating where the Ir dye was present (as confirmed in the substrate background) indicating that the yeast was not able to liberate and accumulate the dye from this substrate (Figure 5). This result can be explained by the difference in chemical composition of the PLGA and PS substrates where the former is an ester-linked polymeric matrix susceptible to degradation to hydrolases whereas the latter cannot be degraded.


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Figure 5. C. albicans grown for 6 days in NANG medium on PLGA and PS substrate with or without embedded Ir-based dye. On the left epifluorescence imaging using transmitted light image shows predominantly yeast forms with some hyphae/pseudohyphae present. On the right, the same image but using 365 nm excitation, and resolved using time gating. For the top and bottom images on the right, fluorescent signal from the embedded IraZolve-L1 dye is visible as a green background. But only in the case where fungi were exposed to the biodegradable PLGA substrate was the dye acquired by the organisms and accumulated intracellularly. Note that the method of time gating has eliminated autofluorescence from C. albicans grown on the PLGA substrate with no probe. Scale bars = 10 µm.

Hyphal development on surfaces and probe uptake Candida albicans is a polymorphic fungus, meaning that it is capable of transitioning from yeast form to produce hyphae.7 In this study, we observed that over the course of 6 days’ growth, an increase of filamentation on both PS and PLGA polymeric surfaces (Figure 5). On the degradable surface, we observed that dye molecules could be


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accumulated within the developing filaments of C. albicans. Hyphal growth is associated with increased protease activity23 which, in the context of animal tissues, contributes to tissue damage as the fungus become invasive.42 Here, we speculate that this increased protease activity contributes to increased substrate digestion and probe uptake, however it is unclear in this case because the supplied minimal growth medium induced morphogenesis on both degradable and non-degradable surfaces. Nevertheless, the fact that the dye is evident in fungal structures such as pseudohyphae is encouraging for future studies aimed at understanding the mechanism by which filamentation contributes to surface degradation of materials. Since our system has produced a model polyester material, findings are also relevant to investigations of how pathogens could degrade abiotic polyester polymers such as those used in medical device manufacture. While microbial degradation of medical grade polyesters is known,43 the general focus of research has been on the consequence to the device itself and its potential failure. We have produced a new insight into fungal microbial/surface interactions by visualizing a consequence of its pathogenicity: the ability to biochemically degrade materials and acquire embedded molecules, such as the


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luminescent probes contained within a polymer matrix. While it is known from previous studies that in solution C. albicans can sustain itself using ester-containing hydrocarbons as the sole source of carbon44 our study suggests that these hydrocarbons could be obtained from surfaces of abiotic materials, if internalized in the same way as the tracer probes. As a model material for ester hydrolysis, it is interesting to note that the rate of PLGA degradation by enzymes is influenced by the proportion of lactic and glycolic acid residues in the polymer. Cai et al. reported that the rate of PLGA degradation by a trypsin solution was faster on PLGA (50/50) than on PLGA (70/30).45 Thus it would be interesting in future work to investigate whether the rate of PLGA hydrolysis by an organism (Candida) could be influenced by the polymer composition. Additionally, C. albicans shows a remarkable ability to survive on and adapt to different sources of carbon as a way of colonizing different host environments where nutrition may be available in different forms.24 Our results show the ability of C. albicans to survive on surfaces when available nutrition is poor presumably by digesting plastics and accumulating hydrolysis products. This has implications to better understanding the role of non-specific secreted hydrolases as a virulence factor in fungal pathogenesis.


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Conclusion We have investigated the role of hydrolysis in fungal colonization of surfaces. In minimal medium, the yeast C. albicans grew on and degraded the ester-linked polymer PLGA. Transition-metal containing fluorescent dyes embedded in this matrix could only be liberated by the degradation-action of the organism and could not be passively released by soaking. The dyes were then taken up intracellularly and distributed throughout the cell. Hydrolyzed fragments of the PLGA polymer are probably acquired in a similar way based on the observation that even simple lipids can sustain C. albicans growth as the sole source of carbon;44 however, proving this assumption with PLGA will require further experimentation. By selecting appropriate dyes and acquisition parameters, we demonstrated unambiguous discrimination of the signal produced by accumulated dye and potentially confounding signals caused by naturally occurring autofluorescence. This study helps to understand the role of non-specific enzymatic degradation of surfaces, how hydrolysis products are trafficked in the pathogen, and the implications for colonization and biodegradation of polymeric materials relevant to host tissues and medical devices.


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SUPPORTING INFORMATION

Figure S1. Thickness of polymer coatings before and after soaking.

Figure S2. XPS characterization of PS and PLGA.

Figure S3. ToF SIMS characterization of materials.

Figure S4. Spectral intensity versus wavelength for the ReZolve-ER luminescent probes embedded in materials.

Figure S5. Comparison of emission intensities for IraZolve-L1 and ReZolve-ER probes.

Detailed method for culturing C. albicans on biodegradable surfaces.

Funding Sources This work was funded by the University of South Australia’s Research Themes Investment Scheme.


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Acknowledgements We acknowledge use of equipment from the South Australian Node of the Australian National Fabrication Facility (ANFF-SA).

We acknowledge the National Mycology Reference Centre, SA Pathology, Adelaide in providing organisms used in biological experiments.

Disclosures SEP is a shareholder in ReZolve Scientific.

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Table of contents figure:


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Figure 1. Schematic illustration of the materials and biological experiments. A) Spin casting was used to incorporate Re or Ir probe molecules in a polymer matrix that was non-biodegradable (polystyrene) or biodegradable (poly(lactic-co-glycolic acid)). B) Candida albicans grown on biodegradable surface coatings were able to digest the matrix and acquire the probe molecules intracellularly. 194x87mm (150 x 150 DPI)


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Figure 2. Total concentration of embedded Re or Ir ions that could be potentially leached from PS or PLGA thin films over the course of 4 days’ soaking. Values are averages and error bars represent the standard deviation (n=3). 177x115mm (150 x 150 DPI)



Figure 3. (A) Schematic illustrating the imaging method in which samples were excited by a 365 nm pulse LED for 1 ms and no excitation was performed for 9 ms. When time gating was on, a chopper was used to prevent initial natural fluorescence reaching the camera. (B) Micrographs illustrate that the yeast and PLGA polymer have a faint autofluorescent signal when gating was not used (gate off), but could be removed by gating this emission (gate on). Using materials embedded with ReZolve-ER dye (C) the IraZolve-L1 dye (D) provided long-lived emission. Thus, when gating was on, the fluorescent signal was still bright, and not interfered by autofluorescence. Note that the probes are initially distributed throughout the polymer film substrate, which is why both the organisms and the background produce a fluorescent signal. 159x63mm (220 x 220 DPI)


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Figure 4. C. albicans grown for 2 days (upper) or 4 days (bottom) in NANG medium on PLGA substrate with embedded IraZolve L1. Representative micrographs using the time gating procedure show many yeast forms, and individual colonies preferring to cluster together in transmitted light images (left). Few hyphal filaments are present. Scale bars = 10 µm. 159x166mm (220 x 220 DPI)



Figure 5. C. albicans grown for 6 days in NANG medium on PLGA and PS substrate with or without embedded Ir-based dye. On the left epifluorescence imaging using transmitted light image shows predominantly yeast forms with some hyphae/pseudohyphae present. On the right, the same image but using 365 nm excitation, and resolved using time gating. For the top and bottom images on the right, fluorescent signal from the embedded IraZolve-L1 dye is visible as a green background. But only in the case where fungi were exposed to the biodegradable PLGA substrate was the dye acquired by the organisms and accumulated intracellularly. Note that the method of time gating has eliminated autofluorescence from C. albicans grown on the PLGA substrate with no probe. Scale bars = 10 µm. 148x228mm (220 x 220 DPI)


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Table of contents figure 84x46mm (150 x 150 DPI)


Visualizing biomaterial degradation by Candida albicans using

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