Skip to main content
NCBI home page
ACS Omega logoLink to ACS Omega
. 2021 Jan 4;6(2):1361–1369. doi: 10.1021/acsomega.0c04980

Cell Rupture and Morphogenesis Control of the Dimorphic Yeast Candida albicans by Nanostructured Surfaces

Naga Venkatesh Kollu 1, Dennis R LaJeunesse 1,*
PMCID: PMC7818643  PMID: 33490795

Abstract

graphic file with name ao0c04980_0010.jpg

Nanostructured surfaces control microbial biofilm formation by killing mechanically via surface architecture. However, the interactions between nanostructured surfaces (NSS) and cellular fungi have not been thoroughly investigated and the application of NSS as a means of controlling fungal biofilms is uncertain. Cellular yeast such as Candida albicans are structurally and biologically distinct from prokaryotic microbes and therefore are predicted to react differently to nanostructured surfaces. The dimorphic opportunistic fungal pathogen, C. albicans, is responsible for most cases of invasive candidiasis and is a serious health concern due to the rapid increase of drug resistance strains. In this paper, we show that the nanostructured surfaces from a cicada wing alter C. albicans’ viability, biofilm formation, adhesion, and morphogenesis through physical contact. However, the fungal cell response to the NSS suggests that nanoscale mechanical interactions impact C. albicans differently than prokaryotic microbes. This study informs on the use of nanoscale architecture for the control of eukaryotic biofilm formation and illustrates some potential caveats with the application of NSS as an antimicrobial means.

1. Introduction

Candida albicans (C. albicans) is a dimorphic opportunistic fungal pathogen and a commensal member of the human microbiome.1−6C. albicans infections in tissue or on biomedical devices are a serious problem in immunodeficient patients and implant recipients.7−10 Candida infections result in invasive candidiasis, which often leads to fungal sepsis and 100 000 deaths worldwide per year.1,11−15 Moreover, mature Candida biofilms are structurally complex and often serve as a reservoir for more serious bacterial infections via biofilm encapsulation.12,16−21 Candida biofilms often host pathogenic microbes like Methicillin-resistant Staphylococcus aureus and as such are a major source of secondary bacterial infections.22−26 Candida biofilm controls are diverse and have involved a wide range of material interventions including polymeric composites that contain metal and metal oxide nanomaterials, biopolymer thin films, and even structural low aspect ratio colloidal monolayers;27−32 however, little work has been done on nanoscale two-dimensional (2D) materials like the high aspect ratio nanostructured surfaces (NSS) derived from insect wings.

The structure and composition of the Candida biofilm depend on the properties of the contact surface, environmental factors, fungal cell morphology, and the fungal species.12,28,33−35 Antimicrobial NSS have been proposed as a stand-alone treatment or as a potential additive treatment with existing antimicrobial agents.36−41 NSS were initially identified as structurally defining components of insect wings, but more recently these have been fabricated in materials like polymers or metal oxides.42−49 NSS inhibit microbial growth and viability via mechanical interrogation of a microbial cell.38,50−54 Physical parameters like cell–substrate adhesion and cell wall rigidity appear to play a role in NSS-induced cell rupture.55−58 Biophysical models suggest that the bactericidal mechanism is a result of physical incompatibility between the cell membrane and the surface.41,59−65

In this manuscript, we characterized the interaction of the cellular fungi C. albicans with NSS found on the wing of the dog day cicada, Neotibicen tibicen. We define critical timing events associated with specific physiological and morphological responses of the C. albicans cell to the NSS. C. albicans reacts to the NSS differently than other microbes exhibiting a slower rupture rate and evidence of mechanically induced cell wall stress. We also have identified a novel example of thigmotropism, a mechanosensory response of a cell or organism to a surface, regarding hyphal morphogenesis of C. albicans, which may provide a novel means of controlling C. albicans pathogenesis.

2. Results and Discussion

2.1. Surface Characterization

We examined C. albicans/surface interactions on a NSS derived from the wings of the dog day cicada (N. tibicen); this NSS contains an ordered array of nanoscale cones (Figure 1), 200 nm tall, 200 nm wide with 30 nm tips, and spaced 200 nm apart.38 We used a flat glass coverslip for the controls. To control for surface composition effects, all experiments used uncoated surfaces and surfaces glow-coated with a 7 nm layer of gold. We observed a slight increase in hydrophobicity with the deposition of 7 nm gold later to the flat glass and a light reduction in contact of the Au-coated NSS when compared to the native NSS (Table 1). However, all of the surfaces used in the experiments displayed a similar range of contact angles/surface energy.

Figure 1.

Figure 1

Open in a new tab

SEM micrographs of surfaces used in these experiments. (A) SEM of the flat, glass coverslip that was used as a control. (B) Image of the glass coverslip. (C) SEM micrograph of the NSS from the dog day cicada wing, N. tibicen. (D) Graphical representation of the NSS.

Table 1. Contact Angle for Control and NSS Used in This Study.

surface native/uncoated gold coated
glass coverslip 79.8 ± 01° 93.1 ± 0.04°
NSS/cicada wing 114.6 ± 0.1° 87.0 ± 0.1°
Open in a new tab

2.2. C. albicans Adhesion to NSS

The formation of a stable adhesion between a microorganism and a surface is the first step of biofilm formation.66−70 To determine the role that NSS roughness had on C. albicans cell–surface adhesion, we cultured both cellular and hyphal forms of C. albicans on flat and NSS and counted the number of cells bound to the surface. On control surfaces, the number of C. albicans cells/FOV increased as a function of the time of incubation on flat surfaces, both Au coated and uncoated (Figures 2 and S1), and during the first 8 h, the number of cells/FOV on the NSS mirrored those on control surfaces (Figures 2 and S1). After 16 h the rate of cell deposition on NSS slowed relative to control surfaces, but eventually reached the same surface density after 24 h.71,72

Figure 2.

Figure 2

Open in a new tab

Surface–cell adhesion of C. albicans with control and NSS. (A) Confocal micrographs of CFW labeled over time. First column shows C. albicans cells cultured on flat control surfaces; the second column shows C. albicans cells cultured on NSS. (B) Average number of cells per field of view with time for each condition.

2.3. C. albicans Cell Wall Composition Response to NSS Interaction

The polysaccharide chitin provides mechanical stability to the yeast cell wall, and its expression level is controlled by mechanical stress via the cell wall integrity signaling pathway.73−75 During the first 4 h of contact with an NSS We observe a significant increase in the fluorescence of chitin binding dye Calcofluor white (CFW) in C. albicans cells (Figures 3 and S2). In contrast, C. albicans cells cultured on control surfaces show no change in levels of CFW fluorescence over the time course (Figures 3 and S2). However, after 4 h, the CFW fluorescent levels of C. albicanson NSS dropped to the level observed on control surfaces. The transient chitin increase/stress response suggests that either the yeast cells recover from NSS mechanical stress or the NSS stops presenting a mechanical challenge to the C. albicans cell. NSS activity is complex; some model NSS as a passive mechanical device, in which microbes impale themselves onto the nanostructures, while other models evoke a more active, “nano-snare” device in which cellular interactions with the surface trigger lateral mechanical reactions.41,76−79 In either case, it is unclear how a cell would compensate for either activity. However, microbial activity such as the deposition of extracellular matrix and dead cells has been shown to alter surface properties such as super hydrophobicity,80 and it is possible that C. albicans biofilm foul the NSS during culture.

Figure 3.

Figure 3

Open in a new tab

NSS induces an increase in cell wall chitin. The CFW intensity in the C. albicans cell walls is represented as a grayscale value (y-axis) and is examined over six time points (x-axis) on gold-coated flat and NSS substrates.

2.4. C. albicans Viability and Rupture on NSS

NSS with a high aspect ratio nanoscale architecture rupture Gram-negative bacteria within moments after contact.53,54,79,81,82 We examined changes to C. albicans cell morphology on control and NSS materials using scanning electron microscopy. In these experiments, we observed ruptured C. albicans cells on the NSS after 8 h of incubation on the NSS. These ruptured cells resembled the ruptured Saccharomyces cerevisiae yeast cells from previous work and appeared as deflated cells and the deposition of cell wall remnants and other materials (Figure 4, second row, second and third columns, arrows).38

Figure 4.

Figure 4

Open in a new tab

Scanning electron micrographs of the interaction of C. albicans with flat control surfaces and NSS. First column C. albicans cells cultured on a flat surface: top row, 0 h, an ovoid/intact cell; middle row at 8 h of contact, a cluster of ovoid/intact cells; bottom row 16 h of contact, a large cluster of intact cells. Second column, top row, two ovoid intact cells, the arrow denotes a common feature of NSS interactions with yeast cells, a trough of crushed nanoscale cones; the middle row 8 h of contact, a deflated yeast cell, arrow; and the bottom row at 16 h of contact, a field of ruptured cells and cellular debris that is identified by altered contrast (arrows).

To complement these experiments, we labeled C. albicans cells with the vital dye FUN-1 and monitored FUN-1 fluorescence on control and experimental surfaces with confocal microscopy. FUN-1 is a ratio metric dye that is taken up and processed by living, metabolically active fungal cells; upon cellular uptake, the FUN-1 dye fluoresces red and in metabolically active cells the FUN-1 dye will be enzymatically processed to a product with green fluorescence.83 We observed a reduction in viability in C. albicans cells that have been exposed to NSS, as indicated by loss/reduction of green FUN-1 fluorescence (Figures 5A, arrows, and S3). The maximal loss of green FUN-1 fluorescence in cells that have been exposed to the NSS for 8 h and increased as incubation time increased (Figure 5B).

Figure 5.

Figure 5

Open in a new tab

Cell viability of C. albicans stained with fluorescent dye FUN-1. Merged images showing cells labeled with CFW (blue) and cells labeled with FUN-1. Cells labeling with green fluorescence live cells and those only containing red cells (arrows) are the dead or dying cells. (A) Representative confocal micrographs of cells on either Flat control surface (first column) or NSS (second column). (B) Graphical summary of the percentage of dead/dying cells on each surface over the time recorded. **p values < 0.001.

To confirm these results, we measured lactate dehydrogenase (LDH) release and metabolic activity in C. albicans cells cultured on control and NSS. Release of lactate dehydrogenase (LDH) is indicative of cell rupture.84C. albicans cells exposed to NSS exhibited increased LDH release relative to controls (Figure 6A). We also observed a reduction of total cellular metabolic activity of C. albicans cells exposed to NSS, when compared to the control surfaces using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay (Figure 6B). The timing of C. albicans loss of viability/rupture was preceded by NSS-induced increase in cell/wall chitin, suggesting that NSS rupture may result from an early NSS-induced cell wall remodeling response.

Figure 6.

Figure 6

Open in a new tab

LDH and MTT assays of C. albicans exposed to control and NSS. (A) Normalized LDH release by C. albicans on NSS; (B) MTT assay of C. albicans on flat/control surfaces (gray columns), NSS (white columns), and a negative control treatment with antifungal drug voriconazole (black columns); y-axis shows absorbance at 570 nm and x-axis shows time.

When compared to the rates that have been demonstrated for bacterial cell rupture via NSS, C. albicans ruptures on NSS was slower, measured in hours versus minutes, and to a lesser extent than bacterial cells.85 Cellular fungi such as C. albicans are larger (i.e., 5 μm versus 1–2 μm), have a different shape (i.e., ovoid versus cylindrical), and are structurally different from bacterial cells with cell walls that are an order of magnitude stronger than most bacterial cell walls.86,87 The lower amount of NSS-induced rupture of C. albicans cells when compared to what has been described in the literature for bacteria may reflect these structural/physical differences.

2.5. Surface Structure and Biofilm Formation

Many microbes including cellular yeast respond to contact with surfaces through the deposition of extracellular matrix.34,88,89 We examined C. albicans biofilm production in response to control and NSS; we examined biofilm production using a crystal violet assay.90 In control experiments, we observed an increase in biofilm deposition as the culture ages, and after 24 h, we observed a thick and mature biofilm (Figure 7A); however, C. albicans cells cultured on NSS showed a significantly reduced biofilm when compared to the control surfaces (Figure 7B).

Figure 7.

Figure 7

Open in a new tab

Biofilm formation of C. albicans on control and NSS. (A) Scanning electron micrograph of biofilm formation of C. albicans on a flat surface. (B) Crystal violet biofilm assay of C. albicans cultured on NSS and other surfaces. Quantification of crystal violet at 550 nm absorbance collected in a microtiter plate at different incubation time periods.

2.6. NSS Interactions with C. albicans Hyphae

C. albicans is a polymorphic fungus with hyphal morph that is associated with pathogenesis and virulence.11,17 Hyphal C. albicans exhibited no alteration in adhesion, viability, or loss of cell integrity on NSS when compared to control surfaces, which suggests that this morphological form of C. albicans responded differently to the NSS. However, NSS contact inhibits C. albicans ability to form a hyphal morphology. In control experiments, we observed 80 ± 0.81% (n = 30) hyphal differentiation of the C. albicans microcolonies when cultured with a hyphal-induction medium. However, microcolonies of C. albicans cultured with this same medium on NSS never resulted in hyphal formation (Figure 8B, second column). Other materials and conditions have been shown to inhibit C. albicans hyphal formation including natural and synthetic peptides, plant-derived compounds, polymeric/oligomeric materials, and metal oxide-based nanomaterials.91−94 In some of these cases, a reduction of biofilm was associated with the inhibition of hyphal formation. Many of these conditions or treatments appear to interfere with the chemical signals and quorum sensing pathways. The NSS results may be the first example of a purely structural inhibition of hyphal morphogenesis.

Figure 8.

Figure 8

Open in a new tab

NSS controls C. albicans morphogenesis. (A) Schematic of the microcolony assay; (B) first column, C. albicans cultured on flat control surfaces in differentiation media. Top image, confocal micrograph showing hyphal formation (arrow); bottom image, SEM showing hyphal growth; second column, C. albicans cultured in differentiation medium on NSS. (Top) A representative confocal micrograph shows cellular forms of C. albicans (arrow); bottom image, an SEM showing a cluster of C. albicans cells, note the lack of any hyphae.

Contact-dependent/mechanical response or thigmotropism has been observed in plant root tips and fungal hyphal growth.95−100 Perturbation of these structures results in a stretching of the cell wall and/or plasma membrane, thereby activating an intercellular signaling cascade.95 While microscale structures have been shown to redirect the growth of fungal hyphae, this is among the first nanoscale thigmotropic response that controls a genetic process, i.e., fungal cell morphology switch. Many different environmental conditions result in the switch from a cellular of C. albicans to a hyphal morphology including nutrient levels, pH, temperature, and cell density92,97,98 and is inhibited by the quorum sensing through the activation of the cAMP-PKA pathway.92,101,102 It is unclear how an NSS, an external mechanical challenge, inhibits hyphal morphogenesis; however, in some fungal species such as the fission yeast Schizosaccharomyces pombe there is some cross-talk between the cell wall integrity and cAMP-PKA pathways,103 which suggests that mechanical cues may impinge pathways needed for hyphal morphogenesis. Future work using genetic knockouts of components of these pathways will be necessary to determine how interaction with NSS controls intracellular signaling pathways to inhibit hyphal morphogenesis.

3. Conclusions

The application of NSS for the control of bacterial biofilms has made them an attractive alternative to or complement to traditional antibiotics. The rupture of bacterial cells on native and synthetic NSS is well documented and has been ascribed to physical incompatibility between the bacterial cell and the surface that depends on cell wall mechanics and NSS structure. However, a role of NSS for controlling pathogenic fungal biofilms is not as clear. As an antifungal surface, NSS lags significantly in both the timing and the amount of C. albicans rupture when compared to NSS bactericidal effects. In medically relevant biofilms where fungal and bacterial microbes coexist, this may lead to preferential selection for fungal growth. The interaction between the cellular yeast cell and NSS may be a more complex than that between bacteria and the NSS, perhaps involving other surface properties and/or evoking different biological responses. Understanding and controlling these differences will be critical for NSS controls of microbial biofilm formation in medically relevant situations.

4. Experimental Section

4.1. Yeast Strains and Cultures

A wild type strain of C. albicans (ATCC 90028) was grown in Sabouraud dextrose broth (SDB) at 25 °C in 50 mL conical flasks. Depending on the experimental needs, cultures were grown to the mid-log phase at an optical density at 600 nm (OD600) ∼0.6. OD600 measurements were made using a Thermo Scientific Nanodrop 2000C spectrophotometer. To induce hyphal growth, C. albicans was cultured in a modified SDB media containing 10% bovine serum albumin (81066, Sigma-Aldrich) in 50 mL conical flasks incubated overnight in a shaking incubator at 37 °C and 200 rpm. Hyphal growth was indicated by the formation of granular sediments in the growth media.

4.2. Surface Preparation and Characterization

We investigated the interaction of C. albicans cells with flat control coverslips and compared to NSS from the wing of the cicada, N. tibicen (BioQuip Products, Inc., CA). Wings from whole cicadas were dissected from the organism. Isolated wings were sonicated in 70% ethanol for 10 min to remove any contaminants, air-dried at room temperature, and carefully cut into smaller pieces of size 5 mm × 5 mm. Glass coverslips were cleaned using 70% ethanol. The cleaned sections of the cicada wings were mounted on cleaned coverslips using silicone glue (Silicone VC6-1/2). All of the surfaces (wings and coverslips) used in the experiments were glow-coated with a 7 nm layer of gold using a Leica EM ACE200; this was an additional control for differences in surface composition. The surface energy was measured via contact angle goniometry using a Rame-Hart 260-F4 goniometer and DROPIMAGE advanced software. A volume of 3 μL of deionized water was used on all surfaces, and the averages of four repetitions on similar surfaces were calculated.

4.3. Culturing Conditions for Imaging C. albicans Response to NSS

To examine the interactions of C. albicans cells with a different surface, the surfaces with cells were placed in a poly(ethylene glycol) (PEG) 8000 treated (Sigma, Cat # 1546605) well of a 24-well plate.; PEG treatment prevented nonspecific interactions of yeast cells with nonexperimental surfaces. Cells were imaged using a Zeiss Z1 Spinning disk confocal with a setting appropriate for each dye set at six time points: 0, 2, 4, 8, 16, and 24 h.

4.4. Determining Cell Viability

The vital dye, FUN-1 (ex ∼470/560–610 em1, em2 510–560), measures cell viability. Cells were labeled with a 5 mM solution of FUN-1 stain (Molecular Probes F-7030). FUN-1-labeled C. albicans cells were imaged using a Zeiss Z1 spinning disk confocal with excitation with the 488 laser and imaging with simultaneously with the 515 nm filter and 561 nm filters settings and the 100× objective. A CyQUANT MTT assay kit (Life Technologies, Catalog#: V13154) was used to measure viabilty and a CyQUANT LDH Cytotoxicity 2001 assay kit (Life Technologies, Catalog#: C20300) to measure LDH. For these experiments, C. albicans yeast cultures were grown to an OD600 of 0.6/∼1 × 106 cells/mL. Negative controls for these experiments were replicated C. albicans grown parallelly and treated with an antifungal drug (7 μg/mL) voriconazole.

4.5. Densitometric Analysis of C. albicans Cell Wall Chitin

The fluorescent dye Calcofluor white (CFW; ex, 370 nm/em 440 nm) binds chitin. C. albicans cells labeled with 1 μg/mL solution of calcofluor white (CFW) (Sigma-Aldrich, 18909). The intensity values were captured using Zen Blue Software from 100 images per sample/condition per time point. All data were statistically analyzed using the ANOVA: single-factor analysis, and statistical significance is denoted by (*) for p ≤ 0.05 and (**) for p ≤ 0.001.

4.6. Biofilm Quantification

Biofilms formed on different nanostructured surfaces were quantified by a crystal violet microtiter dish biofilm formation assay as previously described.90 The absorbance intensity of the crystal violet dye is measured using a BioTek Synergy Mx plate reader at 550 nm.

4.7. Scanning Electron Microscopy (SEM)

C. albicans cells samples were prepared on both the flat surfaces and NSS; cells were fixed overnight with 2.5% glutaraldehyde/2% formaldehyde solution in 0.1 M cacodylate buffer. The samples were dehydrated by an ethanol series (e.g., 30, 50, 70, 95, and 100%) with 10 min washes/exchanges at each concentration. The samples were air-dried, mounted onto aluminum SEM stubs, and coated with a 5 nm gold layer using Leica EM ACE200. The samples were then observed using a Zeiss Auriga scanning electron microscope.

4.8. Microcolony Assay

To determine the effect of NSS on the morphogenesis of C. albicans from its cellular to its hyphal morphology, we performed a microcolony assay; 100 μL of C. albicans yeast was cultured in a modified SBD medium that contained 10% bovine serum albumin and was placed on control and experimental surfaces and incubated at 37 °C overnight. C. albicans cellular forms that had been cultured in SBD media and C. albicans hyphal forms that had been induced on glass coverslips served as negative and positive controls. To evaluate the microcolony assay, microcolony C. albicans cultures on control and NSS were washed twice with PBS, labeled with CFW, and imaged using a confocal microscope as described above.

Acknowledgments

This work was supported by an NIH BIB Grant to D.R.L. (1R15EB024921-01). This work was performed as a member of Southeastern Nanotechnology Infrastructure Corridor (SENIC) and National Nanotechnology Coordinated Infrastructure (NNCI), which is supported by the National Science Foundation (ECCS-1542174).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.0c04980.

  • S1: Surface–cell adhesion of C. albicans with uncoated control and NSS surfaces. S2: NSS induces the increase in cell wall chitin. S3: Cell viability of C. albicans stained with fluorescent dye FUN-1 in different channels (PDF)

The authors declare no competing financial interest.

Supplementary Material

ao0c04980_si_001.pdf (687.3KB, pdf)

References

  1. Neville B. A.; d’Enfert C.; Bougnoux M. E. Candida albicans commensalism in the gastrointestinal tract. FEMS Yeast Res. 2015, 15, fov081 10.1093/femsyr/fov081. [DOI] [PubMed] [Google Scholar]
  2. Pellon A.; Nasab S. D. S.; Moyes D. L. New Insights in Candida albicans Innate Immunity at the Mucosa: Toxins, Epithelium, Metabolism, and Beyond. Front. Cell. Infect. Microbiol. 2020, 10, 81 10.3389/fcimb.2020.00081. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Nobile C. J.; Johnson A. D. Candida albicans Biofilms and Human Disease. Annu. Rev. Microbiol. 2015, 69, 71. 10.1146/annurev-micro-091014-104330. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Forbes J. D.; Bernstein C. N.; Tremlett H.; Van Domselaar G.; Knox N. C. A Fungal World: Could the Gut Mycobiome Be Involved in Neurological Disease?. Front. Microbiol. 2019, 9, 3249 10.3389/fmicb.2018.03249. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Nash A. K.; Auchtung T. A.; Wong M. C.; Smith D. P.; Gesell J. R.; Ross M. C.; Stewart C. J.; Metcalf G. A.; Muzny D. M.; Gibbs R. A.; et al. The gut mycobiome of the Human Microbiome Project healthy cohort. Microbiome 2017, 5, 153 10.1186/s40168-017-0373-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bernard C.; Girardot M.; Imbert C. Candida albicans interaction with Gram-positive bacteria within interkingdom biofilms. J. Mycol. Med. 2020, 30, 100909 10.1016/j.mycmed.2019.100909. [DOI] [PubMed] [Google Scholar]
  7. Sardi J. C.; Scorzoni L.; Bernardi T.; Fusco-Almeida A. M.; Giannini M. J. M. Candida species: current epidemiology, pathogenicity, biofilm formation, natural antifungal products and new therapeutic options. J. Med. Microbiol. 2013, 62, 10. 10.1099/jmm.0.045054-0. [DOI] [PubMed] [Google Scholar]
  8. Cavalheiro M.; Teixeira M. C. Candida Biofilms: Threats, Challenges, and Promising Strategies. Front. Med. 2018, 5, 28 10.3389/fmed.2018.00028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Kumamoto C. A.; Gresnigt M. S.; Hube B. The gut, the bad and the harmless: Candida albicans as a commensal and opportunistic pathogen in the intestine. Curr. Opin. Microbiol. 2020, 56, 7–15. 10.1016/j.mib.2020.05.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Van Thiel D. H.; George M.; Moore C. M. Fungal Infections: Their Diagnosis and Treatment in Transplant Recipients. Int. J. Hepatol. 2012, 2012, 106923 10.1155/2012/106923. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Ahmad Khan M. S.; Alshehrei F.; Al-Ghamdi S. B.; Bamaga M. A.; Al-Thubiani A. S.; Alam M. Z. Virulence and biofilms as promising targets in developing antipathogenic drugs against candidiasis. Future Sci. OA 2020, 6, FSO440 10.2144/fsoa-2019-0027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. de Mello T. P.; de Souza Ramos L.; Braga-Silva L. A.; Branquinha M. H.; Dos Santos A. L. Fungal Biofilm - A Real Obstacle against an Efficient Therapy: Lessons from Candida. Curr. Top. Med. Chem. 2017, 17, 1987–2004. 10.2174/1568026617666170105145227. [DOI] [PubMed] [Google Scholar]
  13. Neves R. P.; de Carvalho Parahym A. M. R.; da Silva C. M.; Macêdo D. P. C.; Leal A. F. G.; Neves H. J.; Lima-Neto R. G.. Fungal Infections in Neonatal Intensive Care, Selected Topics in Neonatal Care; IntechOpen, 2017. [Google Scholar]
  14. Dale A. P.; Dedicoat M. J.; Saleem T.; Moran E. Percutaneous breast implant herniation: a rare complication of miliary TB. BMJ Case Rep. 2015, 2015, bcr2014207546 10.1136/bcr-2014-207546. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Ooi A.; Song D. H. Reducing infection risk in implant-based breast-reconstruction surgery: challenges and solutions. Breast Cancer 2016, 8, 161. 10.2147/BCTT.S97764. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Carlisle P. L.; Banerjee M.; Lazzell A.; Monteagudo C.; Lopez-Ribot J. L.; Kadosh D. Expression levels of a filament-specific transcriptional regulator are sufficient to determine Candida albicans morphology and virulence. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 599. 10.1073/pnas.0804061106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Kadosh D.; Mundodi V. A Re-Evaluation of the Relationship between Morphology and Pathogenicity in Candida Species. J. Fungi 2020, 6, 13 10.3390/jof6010013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Zago C. E.; Silva S.; Sanita P. V.; Barbugli P. A.; Dias C. M.; Lordello V. B.; Vergani C. E. Dynamics of biofilm formation and the interaction between Candida albicans and methicillin-susceptible (MSSA) and -resistant Staphylococcus aureus (MRSA). PLoS One 2015, 10, e0123206 10.1371/journal.pone.0123206. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Dos Santos A. L. S.; Galdino A. C. M.; de Mello T. P.; de Souza Ramos L..; Branquinha M. H.; Bolognese A. M.; Neto J. C.; Roudbary M. What are the advantages of living in a community? A microbial biofilm perspective!. Mem. Inst. Oswaldo Cruz 2018, 113, e180212 10.1590/0074-02760180212. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Armbruster C. R.; Parsek M. R. New insight into the early stages of biofilm formation. Proc. Natl. Acad. Sci. U.S.A. 2018, 115, 4317. 10.1073/pnas.1804084115. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Tsui C.; Kong E. F.; Jabra-Rizk M. A. Pathogenesis of Candida albicans biofilm. Pathog. Dis. 2016, 74, ftw018 10.1093/femspd/ftw018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Eix E. F.; Nett J. E. How Biofilm Growth Affects Candida-Host Interactions. Front. Microbiol. 2020, 11, 1437 10.3389/fmicb.2020.01437. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Boisvert A.-A.; Cheng M. P.; Sheppard D. C.; Nguyen D. Microbial Biofilms in Pulmonary and Critical Care Diseases. Ann. Am. Thorac. Soc. 2016, 13, 1615. 10.1513/AnnalsATS.201603-194FR. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Vertes A.; Hitchins V.; Phillips K. S. Analytical Challenges of Microbial Biofilms on Medical Devices. Anal. Chem. 2012, 84, 3858. 10.1021/ac2029997. [DOI] [PubMed] [Google Scholar]
  25. Wu H.; Moser C.; Wang H.-Z.; Høiby N.; Song Z.-J. Strategies for combating bacterial biofilm infections. Int. J. Oral Sci. 2015, 7, 1. 10.1038/ijos.2014.65. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Hickok N. J.; Shapiro I. M.; Chen A. F. The Impact of Incorporating Antimicrobials into Implant Surfaces. J. Dent. Res. 2018, 97, 14. 10.1177/0022034517731768. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Takamiya A. S.; Monteiro D. R.; Gorup L. F.; Silva E. A.; de Camargo E. R.; Gomes-Filho J. E.; de Oliveira S. H. P.; Barbosa D. B. Biocompatible silver nanoparticles incorporated in acrylic resin for dental application inhibit Candida albicans biofilm. Mater. Sci. Eng., C 2021, 118, 111341 10.1016/j.msec.2020.111341. [DOI] [PubMed] [Google Scholar]
  28. Lagree K.; Mon H. H.; Mitchell A. P.; Ducker W. A. Impact of surface topography on biofilm formation by Candida albicans. PLoS One 2018, 13, e0197925 10.1371/journal.pone.0197925. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kumari S.; Lang G.; DeSimone E.; Spengler C.; Trossmann V. T.; Lücker S.; Hudel M.; Jacobs K.; Krämer N.; Scheibel T. Data for microbe resistant engineered recombinant spider silk protein based 2D and 3D materials. Data Brief 2020, 32, 106305 10.1016/j.dib.2020.106305. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Cousins B. G.; Allison H. E.; Doherty P. J.; Edwards C.; Garvey M. J.; Martin D. S.; Williams R. L. Effects of a nanoparticulate silica substrate on cell attachment of Candida albicans. J. Appl. Microbiol. 2007, 102, 757–765. 10.1111/j.1365-2672.2006.03124.x. [DOI] [PubMed] [Google Scholar]
  31. Rosenberg M.; Visnapuu M.; Vija H.; Kisand V.; Kasemets K.; Kahru A.; Ivask A. Selective antibiofilm properties and biocompatibility of nano-ZnO and nano-ZnO/Ag coated surfaces. Sci. Rep. 2020, 10, 13478 10.1038/s41598-020-70169-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Lara H. H.; Lopez-Ribot J. L. Inhibition of mixed biofilms of Candida albicans and methicillin-resistant Staphylococcus aureus by positively charged silver nanoparticles and functionalized silicone elastomers. Pathogens 2020, 9, 784 10.3390/pathogens9100784. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. McCall A. D.; Pathirana R. U.; Prabhakar A.; Cullen P. J.; Edgerton M. Candida albicans biofilm development is governed by cooperative attachment and adhesion maintenance proteins. NPJ Biofilms Microbiomes 2019, 5, 21 10.1038/s41522-019-0094-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Cavalheiro M.; Teixeira M. C. Candida Biofilms: Threats, Challenges, and Promising Strategies. Front. Med. 2018, 5, 28 10.3389/fmed.2018.00028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Pais P.; Galocha M.; Viana R.; Cavalheiro M.; Pereira D.; Teixeira M. C. Microevolution of the pathogenic yeasts Candida albicans and Candida glabrata during antifungal therapy and host infection. Microb. Cell 2019, 6, 142. 10.15698/mic2019.03.670. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Luan Y.; Liu S.; Pihl M.; van der Mei H. C.; Liu J.; Hizal F.; Choi C.-H.; Chen H.; Ren Y.; Busscher H. J. Bacterial interactions with nanostructured surfaces. Curr. Opin. Colloid Interface Sci. 2018, 38, 170. 10.1016/j.cocis.2018.10.007. [DOI] [Google Scholar]
  37. Ivanova K.; Ramon E.; Hoyo J.; Tzanov T. Innovative Approaches for Controlling Clinically Relevant Biofilms: Current Trends and Future Prospects. Curr. Top. Med. Chem. 2017, 17, 1889–1914. 10.2174/1568026617666170105143315. [DOI] [PubMed] [Google Scholar]
  38. Nowlin K.; Boseman A.; Covell A.; LaJeunesse D. Adhesion-dependent rupturing of Saccharomyces cerevisiae on biological antimicrobial nanostructured surfaces. J. R. Soc. Interface 2015, 12, 20140999 10.1098/rsif.2014.0999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Modaresifar K.; Azizian S.; Ganjian M.; Fratila-Apachitei L. E.; Zadpoor A. A. Bactericidal effects of nanopatterns: A systematic review. Acta Biomater. 2019, 83, 29. 10.1016/j.actbio.2018.09.059. [DOI] [PubMed] [Google Scholar]
  40. Zhang X.; Zhang G.; Chai M.; Yao X.; Chen W.; Chu P. K. Synergistic antibacterial activity of physical-chemical multi-mechanism by TiO2 nanorod arrays for safe biofilm eradication on implant. Bioact. Mater. 2021, 6, 12. 10.1016/j.bioactmat.2020.07.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Linklater D. P.; Baulin V. A.; Juodkazis S.; Crawford R. J.; Stoodley P.; Ivanova E. P. Mechano-bactericidal actions of nanostructured surfaces. Nat. Rev. Microbiol. 2021, 19, 8–22. 10.1038/s41579-020-0414-z. [DOI] [PubMed] [Google Scholar]
  42. Rigo S.; Cai C.; Gunkel-Grabole G.; Maurizi L.; Zhang X.; Xu J.; Palivan C. G. Nanoscience-Based Strategies to Engineer Antimicrobial Surfaces. Adv. Sci. 2018, 5, 1700892 10.1002/advs.201700892. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Jaggessar A.; Shahali H.; Mathew A.; Yarlagadda P. Bio-mimicking nano and micro-structured surface fabrication for antibacterial properties in medical implants. J. Nanobiotechnol. 2017, 15, 64 10.1186/s12951-017-0306-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Nowlin K.; LaJeunesse D. R. Fabrication of hierarchical biomimetic polymeric nanostructured surfaces. Mol. Syst. Des. Eng. 2017, 2, 201–213. 10.1039/C7ME00009J. [DOI] [Google Scholar]
  45. Wang Y.; Gao C.; Zhao W.; Zheng G.; Ji Y.; Dai K.; Mi L.; Zhang D.; Liu C.; Shen C. Large-area fabrication and applications of patterned surface with anisotropic superhydrophobicity. Appl. Surf. Sci. 2020, 529, 147027 10.1016/j.apsusc.2020.147027. [DOI] [Google Scholar]
  46. Jiang R.; Hao L.; Song L.; Tian L.; Fan Y.; Zhao J.; Liu C.; Ming W.; Ren L. Lotus-leaf-inspired hierarchical structured surface with non-fouling and mechanical bactericidal performances. Chem. Eng. J. 2020, 398, 125609 10.1016/j.cej.2020.125609. [DOI] [Google Scholar]
  47. Arciola C. R.; Campoccia D.; Montanaro L. Implant infections: adhesion, biofilm formation and immune evasion. Nat. Rev. Microbiol. 2018, 16, 397. 10.1038/s41579-018-0019-y. [DOI] [PubMed] [Google Scholar]
  48. Khodaei M.; Shadmani S. Superhydrophobicity on aluminum through reactive-etching and TEOS/GPTMS/nano-Al2O3 silane-based nanocomposite coating. Surf. Coat. Technol. 2019, 374, 1078. 10.1016/j.surfcoat.2019.06.074. [DOI] [Google Scholar]
  49. Higgins S. G.; Becce M.; Belessiotis-Richards A.; Seong H.; Sero J. E.; Stevens M. M. High-Aspect-Ratio Nanostructured Surfaces as Biological Metamaterials. Adv. Mater. 2020, 32, 1903862 10.1002/adma.201903862. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Wu S.; Zuber F.; Brugger J.; Maniura-Weber K.; Ren Q. Antibacterial Au nanostructured surfaces. Nanoscale 2016, 8, 2620. 10.1039/C5NR06157A. [DOI] [PubMed] [Google Scholar]
  51. Olewnik-Kruszkowska E.; Gierszewska M.; Jakubowska E.; Tarach I.; Sedlarik V.; Pummerova M. Antibacterial Films Based on PVA and PVA–Chitosan Modified with Poly(Hexamethylene Guanidine). Polymers 2019, 11, 2093. 10.3390/polym11122093. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Persat A.; Nadell C. D.; Kim M. K.; Ingremeau F.; Siryaporn A.; Drescher K.; Wingreen N. S.; Bassler B. L.; Gitai Z.; Stone H. A. The mechanical world of bacteria. Cell 2015, 161, 988. 10.1016/j.cell.2015.05.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Bandara C. D.; Singh S.; Afara I. O.; Wolff A.; Tesfamichael T.; Ostrikov K.; Oloyede A. Bactericidal Effects of Natural Nanotopography of Dragonfly Wing on Escherichia coli. ACS Appl. Mater. Interfaces 2017, 9, 6746. 10.1021/acsami.6b13666. [DOI] [PubMed] [Google Scholar]
  54. Hasan J.; Webb H. K.; Truong V. K.; Pogodin S.; Baulin V. A.; Watson G. S.; Watson J. A.; Crawford R. J.; Ivanova E. P. Selective bactericidal activity of nanopatterned superhydrophobic cicada Psaltoda claripennis wing surfaces. Appl. Microbiol. Biotechnol. 2013, 97, 9257. 10.1007/s00253-012-4628-5. [DOI] [PubMed] [Google Scholar]
  55. Höfte H. The Yin and Yang of Cell Wall Integrity Control: Brassinosteroid and FERONIA Signaling. Plant Cell Physiol. 2015, 56, 224. 10.1093/pcp/pcu182. [DOI] [PubMed] [Google Scholar]
  56. Vasudevan R.; Kennedy A. J.; Merritt M.; Crocker F. H.; Baney R. H. Microscale patterned surfaces reduce bacterial fouling-microscopic and theoretical analysis. Colloids Surf., B 2014, 117, 225. 10.1016/j.colsurfb.2014.02.037. [DOI] [PubMed] [Google Scholar]
  57. Xie K.; Yang Y.; Jiang H. Controlling Cellular Volume via Mechanical and Physical Properties of Substrate. Biophys. J. 2018, 114, 675. 10.1016/j.bpj.2017.11.3785. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Odermatt P. D.; Hannebelle M. T. M.; Eskandarian H. A.; Nievergelt A. P.; McKinney J. D.; Fantner G. E. Overlapping and essential roles for molecular and mechanical mechanisms in mycobacterial cell division. Nat. Phys. 2020, 16, 57 10.1038/s41567-019-0679-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Pogodin S.; Hasan J.; Baulin V. A.; Webb H. K.; Truong V. K.; Phong Nguyen T. H.; Boshkovikj V.; Fluke C. J.; Watson G. S.; Watson J. A.; et al. Biophysical model of bacterial cell interactions with nanopatterned cicada wing surfaces. Biophys. J. 2013, 104, 835. 10.1016/j.bpj.2012.12.046. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Hasan J.; Chatterjee K. Recent advances in engineering topography mediated antibacterial surfaces. Nanoscale 2015, 7, 15568. 10.1039/C5NR04156B. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Marquette A.; Bechinger B. Biophysical Investigations Elucidating the Mechanisms of Action of Antimicrobial Peptides and Their Synergism. Biomolecules 2018, 8, 18 10.3390/biom8020018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Glossop H. D.; De Zoysa G. H.; Hemar Y.; Cardoso P.; Wang K.; Lu J.; Valéry C.; Sarojini V. Battacin-Inspired Ultrashort Peptides: Nanostructure Analysis and Antimicrobial Activity. Biomacromolecules 2019, 20, 2515. 10.1021/acs.biomac.9b00291. [DOI] [PubMed] [Google Scholar]
  63. Wu S.; Zuber F.; Maniura-Weber K.; Brugger J.; Ren Q. Nanostructured surface topographies have an effect on bactericidal activity. J. Nanobiotechnol. 2018, 16, 20 10.1186/s12951-018-0347-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Vadillo-Rodríguez V.; Guerra-García-Mora A. I.; Perera-Costa D.; Gónzalez-Martín M. L.; Fernández-Calderón M. C. Bacterial response to spatially organized microtopographic surface patterns with nanometer scale roughness. Colloids Surf., B 2018, 169, 340. 10.1016/j.colsurfb.2018.05.038. [DOI] [PubMed] [Google Scholar]
  65. Berne C.; Ellison C. K.; Ducret A.; Brun Y. V. Bacterial adhesion at the single-cell level. Nat. Rev. Microbiol. 2018, 16, 616–627. 10.1038/s41579-018-0057-5. [DOI] [PubMed] [Google Scholar]
  66. Oh J. K.; Yegin Y.; Yang F.; Zhang M.; Li J.; Huang S.; Verkhoturov S. V.; Schweikert E. A.; Perez-Lewis K.; Scholar E. A.; et al. The influence of surface chemistry on the kinetics and thermodynamics of bacterial adhesion. Sci. Rep. 2018, 8, 17247 10.1038/s41598-018-35343-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Dang H.; Lovell C. R. Microbial Surface Colonization and Biofilm Development in Marine Environments. Microbiol. Mol. Biol. Rev. 2016, 80, 91. 10.1128/MMBR.00037-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Song F.; Koo H.; Ren D. Effects of Material Properties on Bacterial Adhesion and Biofilm Formation. J. Dent. Res. 2015, 94, 1027. 10.1177/0022034515587690. [DOI] [PubMed] [Google Scholar]
  69. Kock C.; Dufrêne Y. F.; Heinisch J. J. Up against the Wall: Is Yeast Cell Wall Integrity Ensured by Mechanosensing in Plasma Membrane Microdomains?. Appl. Environ. Microbiol. 2015, 81, 806. 10.1128/AEM.03273-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Lagree K.; Desai J.; Finkel J.; Lanni F. Microscopy of fungal biofilms. Curr. Opin. Microbiol. 2018, 43, 100. 10.1016/j.mib.2017.12.008. [DOI] [PubMed] [Google Scholar]
  71. Charoenbhakdi S.; Dokpikul T.; Burphan T.; Techo T.; Auesukaree C. Vacuolar H+-ATPase Protects Saccharomyces cerevisiae Cells against Ethanol-Induced Oxidative and Cell Wall Stresses. Appl. Environ. Microbiol. 2016, 82, 3121. 10.1128/AEM.00376-16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Sanz A. B.; García R.; Rodríguez-Peña J. M.; Arroyo J. The CWI Pathway: Regulation of the Transcriptional Adaptive Response to Cell Wall Stress in Yeast. J. Fungi 2018, 4, 1. 10.3390/jof4010001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Dichtl K.; Samantaray S.; Wagener J. Cell wall integrity signaling in human pathogenic fungi. Cell. Microbiol. 2016, 18, 1228–1238. 10.1111/cmi.12612. [DOI] [PubMed] [Google Scholar]
  74. Banavar S. P.; Trogdon M.; Drawert B.; Yi T.-M.; Petzold L. R.; Campàs O.. Coordinating cell polarization and morphogenesis through mechanical feedback. 2020. 10.1101/2020.05.21.108076. [DOI] [PMC free article] [PubMed]
  75. Khatoon Z.; McTiernan C. D.; Suuronen E. J.; Mah T.-F.; Alarcon E. I. Bacterial biofilm formation on implantable devices and approaches to its treatment and prevention. Heliyon 2018, 4, e01067 10.1016/j.heliyon.2018.e01067. [DOI] [PMC free article] [PubMed] [Google Scholar]
  76. Wang L.; Hu C.; Shao L. The antimicrobial activity of nanoparticles: present situation and prospects for the future. Int. J. Nanomed. 2017, 12, 1227. 10.2147/IJN.S121956. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Imani S. M.; Maclachlan R.; Rachwalski K.; Chan Y.; Lee B.; McInnes M.; Grandfield K.; Brown E. D.; Didar T. F.; Soleymani L. Flexible Hierarchical Wraps Repel Drug-Resistant Gram-Negative and Positive Bacteria. ACS Nano 2020, 14, 454–465. 10.1021/acsnano.9b06287. [DOI] [PubMed] [Google Scholar]
  78. Ivanova E. P.; Linklater D. P.; Werner M.; Baulin V. A.; Xu X.; Vrancken N.; Rubanov S.; Hanssen E.; Wandiyanto J.; Truong V. K.; Elbourne A.; Maclaughlin S.; Juodkazis S.; Crawford R. J. The multi-faceted mechano-bactericidal mechanism of nanostructured surfaces. Proc. Natl. Acad. Sci. U.S.A. 2020, 117, 12598–12605. 10.1073/pnas.1916680117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Hwang G. B.; Page K.; Patir A.; Nair S. P.; Allan E.; Parkin I. P. The Anti-Biofouling Properties of Superhydrophobic Surfaces are Short-Lived. ACS Nano 2018, 12, 6050. 10.1021/acsnano.8b02293. [DOI] [PubMed] [Google Scholar]
  80. Hasan J.; Xu Y.; Yarlagadda T.; Schuetz M.; Spann K.; Yarlagadda P. K. D. V. Antiviral and Antibacterial Nanostructured Surfaces with Excellent Mechanical Properties for Hospital Applications. ACS Biomater. Sci. Eng. 2020, 6, 3608. 10.1021/acsbiomaterials.0c00348. [DOI] [PubMed] [Google Scholar]
  81. Li X.; Tsui K.-H.; Tsoi J. K. H.; Green D. W.; Jin X.-z.; Deng Y. Q.; Zhu Y. M.; Li X. G.; Fan Z.; Cheung G. S.-p. A nanostructured anti-biofilm surface widens the efficacy against spindle-shaped and chain-forming rod-like bacteria. Nanoscale 2020, 12, 18864. 10.1039/D0NR03809A. [DOI] [PubMed] [Google Scholar]
  82. Henry-Stanley M. J.; Garni R. M.; Wells C. L. Adaptation of FUN-1 and Calcofluor white stains to assess the ability of viable and nonviable yeast to adhere to and be internalized by cultured mammalian cells. J. Microbiol. Methods 2004, 59, 289. 10.1016/j.mimet.2004.07.001. [DOI] [PubMed] [Google Scholar]
  83. Haque F.; Verma N. K.; Alfatah M.; Bijlani S.; Bhattacharyya M. S. Sophorolipid exhibits antifungal activity by ROS mediated endoplasmic reticulum stress and mitochondrial dysfunction pathways in Candida albicans. RSC Adv. 2019, 9, 41639. 10.1039/C9RA07599B. [DOI] [PMC free article] [PubMed] [Google Scholar]
  84. Nowlin K.; Boseman A.; Covell A.; LaJeunesse D. Adhesion-dependent rupturing of Saccharomyces cerevisiae on biological antimicrobial nanostructured surfaces. J. R. Soc. Interface 2015, 12, 20140999 10.1098/rsif.2014.0999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  85. Pham D. Q.; Bryant S. J.; Cheeseman S.; Huang L. Z. Y.; Bryant G.; Dupont M. F.; Chapman J.; Berndt C. C.; Vongsvivut J.; Crawford R. J.; Truong V. K.; Ang A. S. M.; Elbourne A. Micro- To nano-scale chemical and mechanical mapping of antimicrobial-resistant fungal biofilms. Nanoscale 2020, 12, 19888–19904. 10.1039/D0NR05617K. [DOI] [PubMed] [Google Scholar]
  86. Davì V.; Chevalier L.; Guo H.; Tanimoto H.; Barrett K.; Couturier E.; Boudaoud A.; Minc N. Systematic mapping of cell wall mechanics in the regulation of cell morphogenesis. Proc. Natl. Acad. Sci. U.S.A. 2019, 116, 13833–13838. 10.1073/pnas.1820455116. [DOI] [PMC free article] [PubMed] [Google Scholar]
  87. Zarnowski R.; Sanchez H.; Covelli A. S.; Dominguez E.; Jaromin A.; Bernhardt J.; Mitchell K. F.; Heiss C.; Azadi P.; Mitchell A.; Andes D. R. Candida albicans biofilm-induced vesicles confer drug resistance through matrix biogenesis. PLoS Biol. 2018, 16, e2006872 10.1371/journal.pbio.2006872. [DOI] [PMC free article] [PubMed] [Google Scholar]
  88. Gulati M.; Nobile C. J. Candida albicans biofilms: development, regulation, and molecular mechanisms. Microbes Infect. 2016, 18, 310–321. 10.1016/j.micinf.2016.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  89. O’Toole G. A. Microtiter dish biofilm formation assay. J. Visualized Exp. 2011, 30, 2437 10.3791/2437. [DOI] [PMC free article] [PubMed] [Google Scholar]
  90. Graham C. E.; Cruz M. R.; Garsin D. A.; Lorenz M. C. Enterococcus faecalis bacteriocin EntV inhibits hyphal morphogenesis, biofilm formation, and virulence of Candida albicans. Proc. Natl. Acad. Sci. U.S.A. 2017, 114, 4507–4512. 10.1073/pnas.1620432114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  91. Lee J. H.; Kim Y. G.; Khadke S. K.; Yamano A.; Watanabe A.; Lee J. Inhibition of Biofilm Formation by Candida albicans and Polymicrobial Microorganisms by Nepodin via Hyphal-Growth Suppression. ACS Infect. Dis. 2019, 5, 1177–1187. 10.1021/acsinfecdis.9b00033. [DOI] [PubMed] [Google Scholar]
  92. Lee J. H.; Kim Y. G.; Lee J. Inhibition of Candida albicans biofilm and hyphae formation by biocompatible oligomers. Lett. Appl. Microbiol. 2018, 67, 123–129. 10.1111/lam.13016. [DOI] [PubMed] [Google Scholar]
  93. Valdez-Salas B.; Beltrán-Partida E.; Nedev N.; Ibarra-Wiley R.; Salinas R.; Curiel-Álvarez M.; Valenzuela-Ontiveros Y.; Pérez G. Controlled antifungal behavior on Ti6Al4V nanostructured by chemical nanopatterning. Mater. Sci. Eng., C 2019, 96, 677–683. 10.1016/j.msec.2018.11.086. [DOI] [PubMed] [Google Scholar]
  94. Almeida M. C.; Brand A. C.. Thigmo Responses: The Fungal Sense of Touch. In The Fungal Kingdom; 2017; Chapter 22, pp 487−507. 10.1128/microbiolspec.FUNK-0040-2016. [DOI] [PMC free article] [PubMed] [Google Scholar]
  95. Stephenson K. S.; Gow N. A. R.; Davidson F. A.; Gadd G. M. Regulation of vectorial supply of vesicles to the hyphal tip determines thigmotropism in Neurospora crassa. Fungal Biol. 2014, 118, 287. 10.1016/j.funbio.2013.12.007. [DOI] [PubMed] [Google Scholar]
  96. Fukuda S.; Yamamoto R.; Yanagisawa N.; Takaya N.; Sato Y.; Riquelme M.; Takeshita N.. Trade-off between plasticity and velocity in mycelial growth. 2020. 10.1101/2020.10.25.354373. [DOI] [PMC free article] [PubMed]
  97. Lee H.-J.; Kim H.-S.; Park J. M.; Cho H. S.; Jeon J. H. PIN-mediated polar auxin transport facilitates root–obstacle avoidance. New Phytol. 2020, 225, 1285–1296. 10.1111/nph.16076. [DOI] [PubMed] [Google Scholar]
  98. Vylkova S.; Carman A. J.; Danhof H. A.; Collette J. R.; Zhou H.; Lorenz M. C. The fungal pathogen Candida albicans autoinduces hyphal morphogenesis by raising extracellular pH. mBio 2011, 2, e00055 10.1128/mBio.00055-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  99. Boyce K. J.; Andrianopoulos A. Fungal dimorphism: the switch from hyphae to yeast is a specialized morphogenetic adaptation allowing colonization of a host. FEMS Microbiol. Rev. 2015, 39, 797. 10.1093/femsre/fuv035. [DOI] [PubMed] [Google Scholar]
  100. Lv Q.-Z.; Yan L.; Jiang Y.-Y. The synthesis, regulation, and functions of sterols in Candida albicans: Well-known but still lots to learn. Virulence 2016, 7, 649. 10.1080/21505594.2016.1188236. [DOI] [PMC free article] [PubMed] [Google Scholar]
  101. Rodrigues C. F.; Černáková L. Farnesol and Tyrosol: Secondary Metabolites with a Crucial quorum-sensing Role in Candida Biofilm Development. Genes 2020, 11, 444 10.3390/genes11040444. [DOI] [PMC free article] [PubMed] [Google Scholar]
  102. Schadick K.; Fourcade H. M.; Boumenot P.; Seitz J. J.; Morrell J. L.; Chang L.; Gould K. L.; Partridge J. F.; Allshire R. C.; Kitagawa K.; et al. Schizosaccharomyces pombe Git7p, a member of the Saccharomyces cerevisiae Sgtlp family, is required for glucose and cyclic AMP signaling, cell wall integrity, and septation. Eukaryotic Cell 2002, 1, 558. 10.1128/EC.1.4.558-567.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

ao0c04980_si_001.pdf (687.3KB, pdf)

Articles from ACS Omega are provided here courtesy of American Chemical Society

RESOURCES