Ratiometric Imaging of the in situ pH Distribution of Biofilms by use of

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Biological and Medical Applications of Materials and Interfaces

Ratiometric Imaging of the in situ pH Distribution of Biofilms by use of Fluorescent Mesoporous Silica Nanosensors Stephanie Fulaz, Dishon Wayne Hiebner, Caio Barros, Henry Devlin, Stefania Vitale, Laura Quinn, and Eoin Casey ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.9b09978 • Publication Date (Web): 16 Aug 2019 Downloaded from pubs.acs.org on August 17, 2019

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ACS Applied Materials & Interfaces

Ratiometric Imaging of the in situ pH Distribution of Biofilms by use of Fluorescent Mesoporous Silica Nanosensors

Stephanie Fulaz‡, Dishon Hiebner‡, Caio H. N. Barros, Henry Devlin, Stefania Vitale, Laura Quinn and Eoin Casey*

UCD School of Chemical and Bioprocess Engineering, University College Dublin, Belfield

Dublin 4, Dublin, Ireland

ACS Paragon Plus Environment

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KEYWORDS: pH sensor, mesoporous silica nanoparticles, ratiometric imaging, bacterial biofilm,

Pseudomonas fluorescens

ABSTRACT

Biofilms are communities of microorganisms enclosed in a self-generated matrix of extracellular polymeric substances (EPS). While biofilm recalcitrance and persistence are caused by several factors, a reduction in antimicrobial susceptibility has been closely associated with the generation of pH gradients within the biofilm structure. Cells embedded within the biofilm create a localized acidic microenvironment which is unaffected by the external pH. Therefore, pH monitoring is a promising approach for understanding the complexities of a three-dimensional heterogeneous biofilm. A fluorescent pH nanosensor was designed through the synthesis of mesoporous silica nanoparticles (47 ± 5 nm diameter) conjugated to a pH-sensitive dye (fluorescein) and a pH insensitive dye (rhodamine B) as


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an internal standard (dye-MSNs). The fluorescence intensity of fluorescein (IF) reduced significantly as the pH was decreased from 8.5 to 3.5. In contrast, the fluorescence intensity of rhodamine B (IR) remained constant at any pH. The ratio of IF/IR produced a sigmoidal curve with respect to the pH, in a working pH range of between 4.5 and 7.5. Dye-MSNs enabled the measurement of pH gradients within Pseudomonas fluorescens WCS 365 biofilm microcolonies. The biofilms showed spatially distinct low-pH regions that were enclosed into large clusters corresponding to high cell-density areas. Also present were small lowpH areas spread indistinctly throughout the microcolony, caused by mass transfer effect. The lowest detected pH within the inner core of the microcolonies was 5.1, gradually increasing to a neutral pH towards the exterior of the microcolonies. The dye-MSNs were able to fully penetrate the biofilm matrix and allowed a quantitative ratiometric analysis of pH gradients and distribution throughout the biofilm which was independent of nanoparticle concentration.

INTRODUCTION Biofilms are ubiquitous, dynamic and structurally complex communities of microorganisms embedded in a self-produced matrix of extracellular polymeric


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substances (EPS). The EPS is predominantly made up of polysaccharides, proteins and nucleic acids.1–3 The EPS matrix, in terms of its weight and volume, is a key structural component within interfacial bacterial communities.4 It is a highly ordered structure that provides mechanical support and also offers protection from antibacterial agents and harsh environmental stresses.5–7 A fully developed biofilm is typified by its threedimensional (3D) structure, including both live and dead bacterial cells, a bacteriallyderived polymer matrix and interstitial water channels or pores that can facilitate the exchange of solutes from the surrounding environment.8 An important feature of bacterial biofilms is the development of various gradients of the key biochemical parameters (i.e., pH, oxygen, redox potential, virulence factors and ions).9 The 3D architecture of the EPS matrix can control the differential diffusion of nutrients, metabolic products and oxygen to the interior of the biofilm while also trapping acids inside the microcolonies.10 These gradients have been recognized and implicated in a number of natural, clinical and industrial environments. For example, the heterogenous distribution of charged polymers within the matrix has been shown to increase bacterial tolerance to various antimicrobials through sorption or deactivation of


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these chemically reactive molecules.11,12 Due to bacterial metabolism and the subsequent production of acidic by-products, bacterial cells embedded in the biofilm create a localized acidic microenvironment. The pH of these microenvironments is often unaffected by and can be markedly different from the external pH due to the presence of the EPS matrix.13,14 The occurrence of an acidic environment within the biofilm is a result of low oxygen tension inducing anaerobic fermentation which causes the production of organic acids such as lactic acid and acetic acid in certain bacteria.15,16 Although much research has sought to chemically eliminate biofilms, eradicating unwanted biofilms using smart responsive technologies which are activated by environmental cues are a strategy gaining much traction.17 Bacterial biofilms should be observed as a three-dimensional structure and biofilm analysis needs to be investigated spatially.18 Therefore, the quantitative pH mapping in all dimensions of the biofilms is of great significance to enhance the understanding of the processes leading to the diverse gradients within the biofilm. Even though the inherent variability within biofilms has been well documented, the direct implication of these distinct microenvironments (notably pH and dissolved gas


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concentrations) has been difficult to measure in situ with a high spatiotemporal resolution.19 pH monitoring is a promising approach for understanding the complexities of biofilms with potential applications in diagnostics; however, pH measurement is not a trivial task in a 3D and highly heterogeneous system such as a biofilm. For example, microelectrodes with tip diameters of 10-15 μm have been implemented to measure the pH profiles within dental biofilms.20 Nonetheless, the use of these physical methods disrupts the biofilm structure and cannot evaluate the horizontal pH profiles in real time. Sensors based on analyte-specific effects of fluorophore emission, such as quantum yield, lifetime or wavelength have also been described.21–23 In any case they are only effective as qualitative sensors, as the fluorescence is dependent on the concentration of both analyte and sensor.24 They may also undergo quenching, photobleaching, solvatochromic effect and have a limited brightness.25 Several approaches for dye-related pH sensing exist, including the use of lipid-bilayer vesicles with a reference and a sensor dye.26 Polymer based pH sensors, such as the dextran-based sensors, are commercially available, however, they do not offer the dye protection against bleaching or quenching and may decrease the overall fluorescence


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yield.27,28 The use of a nanoparticle-based system with an internal standard allows the measurement of local pH independent of sensor concentration.29 Sensor dyes covalently bound to a silica matrix can form a robust system for which sensing in a biocompatible and functionalizable surface is achievable. This approach offers reduced photobleaching and solvatochromic effect and increased fluorescence efficiency.30 It also allows the combination of multiple fluorophores in the same nanoparticle, enabling in-situ analysis of the pH gradient inside biofilms, without altering its natural structure. Fluorescein dye is widely applied for pH sensing due to the existence of several protonation states with varying pH.27,29 Changes in the protonation state lead to an alteration of the emission intensity of fluorescein. For instance the monoanion species has a low quantum yield of Ф = 0.36, while the dianion has a quantum yield of Ф = 0.9327 (Figure 1). On the other hand, rhodamine has a near constant emission intensity regardless of the pH. Rhodamine has been used as an internal standard for pH determination, as an indicator of nanoparticle concentration and also to identify the distribution of nanoparticles throughout the biofilm.29


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Figure 1. Structure and fluorescence quantum yield of fluorescein in different pHs.

This study assesses the potential of mesoporous silica nanoparticles (MSNs) sensors as a quantitative indicator of pH within the unique microenvironment inside the microcolonies of bacterial biofilms. We have used fluorescent core/shell MSNs based on fluorescein (pH-sensitive) and rhodamine (reference) dyes (dye-MSNs) in conjunction with confocal laser scanning microscopy (CLSM) to study a single-species Pseudomonas

fluorescens WC365 biofilm and the pH of their microenvironments. Novel techniques for pH monitoring are required to fully understand how changes in the pH gradients within the biofilm not only contributes to biofilm-specific processes but can be used for the development and design of smart pH-responsive biofilm removal technologies. EXPERIMENTAL SECTION Materials


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Hexadecyltrimethylammonium bromide (CTAB, 98%), tetraethylortosilicate (TEOS, 99%), (3-aminopropyl)triethoxysilane (APTES, 99%), fluorescein isothiocyanate (FITC, 90%), rhodamine B isothiocyanate (RITC, mixed isomers), triethanolamine (TEA, 99%), Mowiol 4-88 were purchased from Sigma-Aldrich. Ethanol (99.5%) and hydrochloric acid (HCl, 37%) were obtained from Merck. All reagents were used as received without further purification. Synthesis of dye-MSNs Both FITC and RITC dyes used for MSNs synthesis were previously conjugated with APTES (reaction of 50 µL of APTES with 10 mg of FITC, or RITC, dissolved in 5 mL of ethanol, under stirring at room temperature for 4 hours).31 The synthesis of MSNs was adapted from Moller et al. (2007).32 Briefly, a stock solution of 28% wt CTAB was prepared and sonicated until complete dissolution. The reaction mixture was prepared adding 32 mL of H2O, 5.25 mL of ethanol and 5 mL of CTAB 28 % wt and stirring for 10 min at room temperature. 2.063 mL of TEA was added, and the reaction was heated up to 60 oC. 3.25 mL of TEOS was added dropwise followed by 1 mL of FITC-APTES. The mixture was kept under vigorous stirring at 60 oC for 2 h and protected from the light. 500 µL of TEOS


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and 500 µL of RITC-APTES were added. The reaction was kept under vigorous stirring at 60 oC for 1 h and at room temperature overnight. The suspension was centrifuged at 9000 rpm for 20 min, and the solid washed three times with H2O and twice with ethanol. To remove the CTAB, the solid was suspended in 50 mL of ethanol and heated up to 60 oC.

40 µL of HCl concentrate was added and the reaction was kept under vigorous stirring

at 60 oC for 3 h. The solid was removed by centrifugation, washed with H2O and ethanol and kept in an ethanol suspension protected from light. MSNs characterization Zeta potential of nanoparticles suspension in H2O (1 mg mL-1) and in different McIlvaine buffers (pH 3.0 – 8.0) was determined using a Zetasizer Nano-ZS apparatus (Malvern Instruments). All experiements were done in triplicate, with 10 scans each. Transmission electron microscopy (TEM) analysis of samples dispersed in ethanol and deposited on carbon-coated copper grids were performed using a FEI Tecnai G2. Average size and size distribution of nanoparticles were determined by Fiji software33 using TEM images, by analyzing a minimum of 100 nanoparticles per sample. N2 adsorption-desorption isotherms were recorded in a BET Nova 2400e (Quantachrome, UK) apparatus at 77 K


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under continuous adsorption conditions. Samples were out-gassed at 110 oC for 16 h prior to the experiments. The surface areas were obtained by BET (Brunauer-EmmettTeller) analysis.34 The pore volume and pore size were obtained using the BJH (BarretJoyner-Halenda) model on adsorption. pH calibration curve To evaluate the effect of the pH on fluorescence intensity of the MSNs, 11 different solutions of McIlvaine buffer from pH 3.5 to pH 8.5 were prepared by mixing different volumes of the stock solutions of 0.2 mol L-1 Na2HPO4 and 0.1 mol L-1 citric acid. The dye-MSNs were suspended in H2O (0.5 mg mL-1), added to the wells of optically clear bottom 96-well tissue culture and incubated at 25 oC for 45 min. The supernatant was removed, and the nanoparticles were gently washed with MilliQ H2O to remove unattached MSNs. 100 µL of the corresponding buffer was added to each well, in triplicate. The images of the MSNs attached to the plate substratum were acquired with an Olympus FluorView FV1000 CLSM attached to an inverted Olympus IX81 microscope with a 60x/1.35 NA UPL SAPO oil immersion objective (Olympus Optical, Tokyo, Japan) using two separate laser lines, 488 nm for FITC and 559 nm for RITC. Fluorescence


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emission was collected sequentially. The images were analysed using Fiji image processing software,33 obtaining mean grey values for each region of interest (ROI), with a minimum of 100 ROIs per image. The mean fluorescence intensity (I) was calculated by averaging all results for the replicates. The ratio IF/IR was plotted as a function of the pH values. In order to investigate the influence of dye-MSNs concentration in the ratio IF/IR, 0.5 and 1.0 mg mL-1 H2O suspensions of dye-MSNs were analysed as described previously in different pHs (4.0, 5.0, 6.0 and 7.0). To evaluate the reversibility of the fluorescence measurement, a 1.0 mg mL-1 H2O suspensions of dye-MSNs were incubated with buffered solution pH 3.0 for 15 min. The fluorescence spectra was acquired in a plate reader (SpectraMax iD3, Molecular devices). The dye-MSNs were then washed and incubated in buffered solution of pH 8.0 and the fluorescence spectra was again measured. The process was repeated by incubating the dye-MSNs consecutively in buffered solutions of pH 3.0 and 8.0. Bacterial culture and maintenance The bacterial strain used in this investigation was P. fluorescens WCS365. Bacterial cultures were stored at –80 °C in MilliQ H2O supplemented with 25 % glycerol. For


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cultivation, thawed aliquots were streaked onto King B agar plates and incubated for 24 h at 30 °C. Prior to pH probe exposure experiments, a single bacterial colony was used to inoculate 50 mL of sterile King B medium in a 250 mL Erlenmeyer flask and incubated at 30 °C with shaking at 200 rpm overnight (14 – 16 h) to an approximate OD600 of between 2.3 – 2.6. The overnight cultures were then diluted to a final OD600 of 1.0 using fresh sterile King B medium. Biofilm growth For a single centrifuge tube biofilm, biofilms were prepared as per Safari et al. (2014) with minor modifications.35 A 5 mL volume of the OD600 of 1 diluted culture, supplemented with CaCl2 to a final concentration of 1.5 mM, was added to a sterile 50 mL centrifuge tube containing a glass coverslip (24 mm X 50 mm) and plugged with cotton wool. Tubes were then incubated for 72 h at 30 °C with shaking at 100 rpm. Sample preparation and confocal microscopy MSN suspensions were prepared by diluting NP stock solutions to a final concentration of 0.5 mg mL-1 using H2O. The working solutions were sonicated for 1 h prior to use. For biofilm imaging, each biofilm-coated glass coverslip was carefully removed from the


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centrifuge tubes with sterile forceps. The coverslip was then placed on a sample holder and 150 μL of dye-MSNs (0.5 mg mL-1) was placed on top of the biofilm and incubated at room temperature for 15 min in the dark. The biofilms were then gently rinsed 3 times in H2O and prepared for imaging. Each biofilm-coated coverslip was mounted onto a glass microscope slide (25 mm x 75 mm x 1 mm) as described in the supporting information Figure S1. The coverslip was mounted using Tris-buffered Mowiol 4-88 (pH 8.5) mounting medium to maintain the fully hydrated 3D native state of the biofilm. Horizontal plane z-stack images were acquired with an Olympus FluorView FV1000 CLSM attached to an inverted Olympus IX81 microscope with a 60x/1.35 NA UPL SAPO oil immersion objective (Olympus Optical, Tokyo, Japan). The microscope system was equipped with laser excitation sources at 405, 488, 543 and 633 nm for the excitation of all fluorophores used. All images were acquired equally; 1x digital zoom, a scanning speed of 10.0 μs/pixel, with 3x Kalman line averaging and sequential acquisition. Image stacks were acquired in at least 3 random areas per biofilm experiment with a step size of 0.5 μm. Each 3D stack was rendered using Fiji’s 3D viewer plugin.33


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Image processing and analysis All image files were analysed in Fiji Image Processing Software.36 All images were background subtracted and median filtered prior to all analysis. For 3D stack rendering, Fiji’s 3D viewer plugin was used.33 For ratiometric imaging, the fluorescence intensities of all sensor (FITC) and reference (RITC) images were divided pixelwise using Fiji’s Ratio Plus plugin (https://imagej.nih.gov/ij/plugins/ratio-plus.html) to generate IF/IR values throughout various ROIs in the biofilm which were considered to have enough signal intensity to not be deemed noise. The 1024 x 1024 pixel arrays were used in conjunction with the pH calibration curve to determine the IF/IR values at each region. Output ratio image files were scaled to 8-bit grayscale and a modified rainbow look-up-table (LUT) was used to false colour each ratiometric image and amend the pH scale bar. Unexposed biofilms were imaged to determine the baseline background or autofluorescence at the wavelengths and laser intensity used. No autofluorescence or background signal at the selected excitation and emission were detected in any control biofilm images. To assess cell density in the biofilm, mean grey values of at least 3 ROIs (100 x 100 pixels) per CLSM image single slice (x,y) were measured in Fiji Image Processing Software.


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Dye-MSNs sorption into biofilm To assess dye-MSNs sorption into P. fluorescens biofilms, a method developed by Nevius et al.37 was used with minor modifications. Briefly, biofilm samples were cultured in black 96-well plates at 30 °C for 24 h with orbital shaking at 125 rpm. Dye-MSNs suspensions (0.5 and 1.0 mg mL-1) were exposed to the biofilm for 15 minutes. The fluorescence intensity (λex= 550, λem= 585 nm) from the dye-MSNs was measured using a plate reader (SpectraMax iD3, Molecular devices). The sorption of dye-MSNs into the biofilms was calculated by using the reduction of dye-MSNs fluorescence intensities between the biofilm exposed particles and that of a control.

RESULTS The core/shell MSNs based on fluorescein (pH-sensitive) and rhodamine (reference) dyes (dye-MSNs) were synthesized using an adapted Stöber method38 by cocondensation of APTES-conjugated FITC and RITC. The synthesis in two consecutive steps allows the formation of a core containing the sensor dye and a shell comprised of the reference dye (Figure 2 a).


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The TEM image (Figure 2 b) represents the spherical monodisperse MSNs with a diameter of 47 ± 5 nm. The charge measured by zeta potential was -16.2 ± 0.5 mV in water, the zeta potential values in the buffers with different pHs are represented in Table S1. The dye-MSNs presented a typical type IV adsorption-desorption isotherm (Figure S2 a) and a high surface area (789 m2 g-1). The presence of mesoporous channels was confirmed by the capillary condensation observed at a relative pressure (P/P0) between 0.4 and 0.5. The pore size distribution was determined by the Barret-Joyner-Halenda (BJH) method (Figure S2 b), with a mean pore size of 3.1 nm. The summary of the physical properties is represented in Table 1.


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Figure 2. (a) Schematic structure of dye-MSNs with FITC in the core and RITC in the shell; (b) TEM image of dye-MSNs. Scale bar represents 100 μm.

Table 1. Physical properties of dye-MSNs

Sample

Particle

Pore

sizea (nm)

(nm)

sizeb Surface

Pore volume Zeta

area (m2 g-1) (cm3 g-1)

Potentialc (mV)

Dye-MSNs a

47 ± 5

3.1

789

0.590

-16.2 ± 0.5

Particle sizes were calculated from TEM images, the mean ± standard deviation is

presented.

b

Pore sizes were determined from the adsorption isotherms using Barret-

Joyner-Halenda (BJH) method. c Measured in water. The dye-MSNs were exposed to different pH values in a McIlvaine buffer ranging from pH 3.5 to 8.5. The fluorescence imaging in CLSM was obtained using two laser lines, 488 nm (green channel) for the sensor dye fluorescein and 559 nm (red channel) for the reference dye rhodamine, as represented in Figure 3.


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Figure 3. CLSM image of fluorescently labelled (FITC and RITC) dye-MSNs in presence of different pHs (McIlvaine buffer). (a) Green (FITC) channel and (b) red (RITC) channel. Scale bar represents 100 µm.

Figure 3 shows the direct relationship between pH and the intensity of the FITC. The fluorescence intensity of fluorescein (IF - green) decreases drastically when the pH is decreased from 8 to 4 while the fluorescence intensity of rhodamine (IR - red) is constant regardless of the pH. A ratio between the fluorescence intensity (I) of the two channels (IF/IR) was calculated and plotted against the pH, as shown in Figure 4.


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Figure 4. Curve representing the ratio between the fluorescence intensity in the FITC (green channel) and fluorescence intensity in RITC (red channel) in relation to the pH. Means ± standard deviation is presented.

The sigmoidal calibration curve describes a system in equilibrium between two different states, generally monoanionic and dianionic states with a pKa of 6.4.27 The pH-response range from the calibration curve is between 4.5 and 7.5. The sensing response of dye-MSNs was evaluated regarding the nanoparticle concentration as represented in Figure S3. The two dye-MSNs suspensions, 0.5 and 1


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mg mL-1 had the same performance indicating that dye-MSNs sensing ability is independent of the nanoparticle concentration. To evaluate if the sensing response would not change when exposed sequentially to different pH values, a suspension of dye-MSNs was incubated sequentially in a solution with pH 3.0, followed by pH 8.0, back to pH 3.0 and then pH 8.0. The fluorescence emission in these conditions can be seen in Figure S4, which indicates that the pH response of the dye-MSNs is reversible.

P. fluorescens forms heterogeneous yet well-defined 3D microcolonies comprising of both live and dead bacterial cells as well as an EPS matrix rich in polysaccharides, proteins and eDNA.39,40 After 72 hours of cultivation, P. fluorescens communities were mature enough to form distinct structural features typified by microcolonies organised, on average, into cell clusters of 60 – 100 µm in diameter and 20 – 45 µm in height separated by areas with smaller and less dense cell clusters. The dye-MSNs penetration is a critical step for the ratiometric quantitative measure of the pH of the biofilm microcolonies. The as-synthesized dye-MSNs were able to fully penetrate the biofilm microcolonies and show an even distribution throughout the EPS matrix (Figure S5). The sorption of nanoparticles by biofilm samples was assessed using a method developed by Nevius et al37 with some


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modifications. The results show that regardless of dye-MSNs concentration (0.5 or 1 mg mL-1) a high percentage of dye-MSNs sorption occurred (25%) (Figure S6). A representative 2D schematic diagram of a P. fluorescens biofilm microcolony structure and resultant pH gradient in various segments is depicted in Fig. 5 a. Representative CLSM image slices from individual z-stacks are shown in Fig. 5 b; divided into bottom, middle and top layers of three different microcolonies. The ratiometric in situ mapping of the microcolony pH shows the formation of various spatially heterogeneous pH microenvironments, despite the presence of a slightly alkaline buffered pH in the external environment provided by the mounting medium (pH 8.5). A separated view of the biofilm shows the various spatially distinct pH profiles as follows: (1) regions of low pH are confined to large dense clusters close to the substratum at the deepest regions of the biofilm (Figure 5 b1-bottom), (2) a range of acidic “pockets” of 2 – 5 µm distributed evenly throughout the microcolonies bottom surface (Figure 5 b2-bottom) or (3) a combination of both large dense clusters surrounded by acidic pockets (Figure 5 b3-bottom). The pH within the clusters or pockets is shown to increase as it approaches the microcolony periphery in the lateral direction and is consistent throughout the middle and top layers.


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Figure 5. Lateral pH distribution within an intact microcolony. (a) 2D schematic of distribution of the biofilm microcolonies pH gradient in the bottom, middle and top layers. (b). Ratiometric CLSM x,y cross-section images showing pH gradients within the microcolonies of 3 individual 72 h P. fluorescens biofilms (I-III) in the bottom, middle and top layers, respectively. The pH was analysed ratiometrically based on the FITC (sensor) and RITC (reference) images and is expressed as a false-coloured rainbow spectrum. Scale bars represent 50 µm.


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The development of pH gradients within bacterial biofilms occurs in both the lateral and axial direction. Therefore, cross-sectional y,z images of the biofilm from the glass surface to the biofilm exterior were acquired to illustrate the pH distribution within surfaceattached microcolonies and their corresponding pH values. Figure 6 a. shows a representative 3D reconstruction CLSM z-stack image separated into 5 discrete regions (I – V) surrounding a dense microcolony (visualized in Figure 6 b. III). The development of spatially distinct pH gradients in the axial direction is apparent; with an acidic pH calculated within the deepest region of the microcolony, which increases gradually towards a neutral pH as the top of the microcolony is approached. Well-defined spatially distinct pH profiles within various regions are displayed. In Figure 6 b. I, regions of relatively low cell biomass include acidic pockets which are distributed throughout the biofilm and interestingly, also in close proximity to the top of the biofilm. Figure 6 b. III shows the distinct acidic clusters within the deepest regions of the biofilm and Figure 6 b. IV displaying a combination of both these pH profiles.


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Figure 6. Axial pH distribution within an intact microcolony. (a) Representative 3D CLSM reconstruction and (b) corresponding ratiometric orthogonal slices showing the pH


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gradients in the y,z direction within microcolonies of 72 h P. fluorescens biofilms. Scale bars represent 20 µm.

The ratiometric quantification of pH gradients within the P. fluorescens biofilms is calculated independently from the bacterial cell biomass but rather as part of the EPS matrix biomass. Even so, the presence of bacterial cells in high numbers and also in close proximity to one another inside the biofilm has been shown to dramatically affect bacterial cells both phenotypically and genotypically.41 Therefore, spatially distinct pH gradients must be viewed in the context of bacterial cell density which is relative to each region’s pH profile. Figure S7 exemplify what is considered an area of low cell-density and an area of high cell density based on the overall fluorescence emission. As can be noted areas of high cell density have a fluorescence emission of approximately 4-fold higher than the ones from low cell density. Figure 7 shows the relationship between cell density and lowpH regions. The areas of high cell density, shown in Figure 7. within the white dotted square, correspond to areas with an acidic pH in the cluster-like spatial region.


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Interestingly, areas with low cell density (Figure 7. white arrows) also display the presence of low pH regions.

Figure 7. Representative CLSM images showing the development of low-pH regions within both high cell density (white dotted square) and low cell density areas (white arrows) in the microcolony interior. (a) Bacteria and (b) Ratiometric pH image.

DISCUSSION The current sensor design employed, with a core/shell system composed of a sensitive dye (FITC) and a reference dye (RITC), enabled the quantitative ratiometric analysis of pH throughout the biofilm which is independent of nanoparticle concentration (Figure S3). These dye-MSNs have the potential to be used as “single-particle laboratories” in a myriad of applications including drug-


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delivery with a nanoparticle tracking mechanism. The application of dye-MSNs in the highthroughput screening of molecules and/or drugs for treatment of infectious diseases is just one example of their potential uses when applied to medical, industrial and environmental systems.29,42 The synthesized dye-MSNs were a monodisperse suspension of 47 nm dye-rich core/shell particles. The silica shell acts as a filter, allowing analyte molecules to diffuse to and from the sensor dyes, while protecting the dyes from interactions with larger molecules such as proteins or organic quenchers that could interfere with the measurements.29 The presence of mesoporous channels was confirmed by BET and BJH (Figure S2), indicating the possibility of further functionalization or molecule-encapsulation into the structure. The pH response was verified by fluorescence measurements in MclIvaine buffer solutions from pH 3.5 to 8.5 (data not shown). While FITC fluorescence intensity dropped when the solution’s pH was reduced, the RITC fluorescence intensity remained constant. The measurements were repeated in the CLSM and the same results were obtained as depicted in Figure 3. The ratio between the two confocal channels in a variety of pH conditions yielded the sigmoidal curve observed in Figure 4, as expected. FITC has a pKa of 6.4 and the sigmoidal curve is attributed to the equilibrium of the dianion and monoanion species27 represented in Figure 1. The pH response observed is reversible, i.e. the dye-MSNs have the same fluorescence emission once they reach an acidic pH regardless of being exposed previously to a basic pH (Figure S4). Although the applied methodology is independent of the nanoparticle concentration, the ability of dye-MSNs to penetrate throughout the entire biofilm is paramount. The penetration of nanoparticles within bacterial biofilms is mainly regulated by nanoparticles’ size and charge.43–45 Based on previous studies, the dye-MSNs are expected to interact with the EPS matrix,46 attaching


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to virtually any component, such as cell membranes, polysaccharides, proteins or stay in the biofilm pores or water channels.47,48 This allows for the pH in all types of microenvironments present in the biofilm to be successfully quantified. As can be observed in Figure S5 the dye-MSNs were able to fully penetrate the biofilm and were even encountered in the bottom layer of dense microcolonies. Approximately 25% of the dye-MSNs suspension was sorbed into the biofilm (Figure S6) Biofilms consisting of multiple species of bacteria with different metabolisms (e.g. aerobic, fermentative and anaerobic), will produce more acidic by-products under the oxygen-limiting conditions found in the deepest regions of the biofilm than single-species biofilm consisting of obligate aerobic bacteria.49,50 Therefore, the lowest detected pH for P. fluorescens biofilms was found to be 5.1, this compared to a pH of between 4 – 5 detected for mixed-culture in situ grown oral biofilms.49 As the P. fluorescens biofilm matures, the production of the EPS matrix assisted in the creation of spatial heterogeneities leads to the development of these physicochemical gradients (Figure 6). Xiao et al. (2012) demonstrated the presence of neutral pH regions close to the biofilm periphery, acidic pockets within the microcolony and low pH (4.5 – 5.5) regions at the microcolony/substratum interface in a mixed-species oral biofilm.14 Similarly, P. fluorescens biofilms show spatially distinct pH gradients that are confined and partitioned into clusters or pockets throughout the microcolony and microcolony surface/fluid interface (Figure 6). The presence of discrete pockets of low pH may be attributed to the 3D architecture and porous nature of the biofilm structure51 and indicates the accumulation of acidic metabolites or waste-products produced by the bacteria.14


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This effect occurs simultaneously in both the lateral (Figure 5) and axial (Figure 6) directions. In areas with high cell-densities and therefore an increased local rate of metabolic processing within a defined area inside microcolony core resulted in an increased accumulation of acidic byproducts (Figure 7. white dotted square). Acidic by-products may become trapped and pooled within the confines of the EPS matrix52 leading to the development of low pH cluster regions. In contrast, the development of spatially separated acidic pockets (Figure 7. white arrows) may not be due the result of the accumulation of acidic by-products because of high cell densities but rather due to the effect of mass transfer of solutes through the EPS matrix. Within heterogeneous bacterial biofilms, the rates of mass transfer vary from one region to the next in both the diffusion and distribution of molecular species both into and out of the biofilm,53 explaining the disparity between the relative frequency, distribution and size of the acidic pockets within the microcolony superstructure of all analysed biofilms (Figure 5 b. I-III). Additionally, it has also been suggested that the EPS matrix could serve as an energy reserve in which glucans and fructans could be degraded by extracellular enzymes to release monosaccharides that can be metabolized to produce acidic by-products, especially relevant in mature biofilms.54 The presence of a slightly alkaline-buffered bulk fluid (pH 8.5) on the exterior of the P. fluorescens biofilms did not result in the neutralization of the microcolony interior. Again, due to the presence of the EPS matrix limiting access and diffusion into the biofilm thereby helping to confine the acidic regions within the biofilm structure. Using an EPS-degrading dextranase, Hwang et al. 2016,10 degraded and therefore compromised the ability of the EPS matrix in Streptococcus mutans biofilms to limit diffusion of solutes. Strikingly, most of the biofilm interior, even in large microcolonies, was neutralized after treatment with dextranase, demonstrating the role of the EPS matrix in facilitating the development of these acidic regions. However, the precise


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mechanism by which the biofilm produces and maintains their pH microenvironments remains unclear, it appears to be a highly complex and multifaceted process. While biofilm recalcitrance and persistence is caused by several factors, the reduced antimicrobial susceptibility has also been associated with the generation of pH gradients in the biofilm.55–57 The EPS matrix physicochemical properties are also affected by the pH, with changes in “polymer-polymer, ion-polymer, and macromolecule-polymer interactions, and even polymer motion, gyration, and polymer-polymer entwining”.19 These changes lead to the emergence of a physical barrier that hinders antimicrobials penetration.58 Furthermore the antimicrobials can be protonated/deprotonated thereby inhibiting their action. Acidic moieties produced by bacteria which are present at the attached substratum are well documented virulence factors in many biofilm-related infections.14 With a more acidic pH, there is a higher mineral dissolution and a lower uptake of metal by biofilms, impacting biogeochemistry and bioremediation applications.59,60 Therefore, there is a requirement for a better understanding of biofilm architecture, in relation to its physicochemical gradients, mainly pH, redox potential and oxygen, as well as their impact in biofilm eradication strategies. However, the study of these local microenvironments without disturbing the biofilm natural structure is still a challenge.

CONCLUSIONS

The relevance of the heterogeneity of P. fluorescens biofilms’ 3D architecture and composition on the production and localization of low pH microenvironments is highlighted by our pH mapping


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approach which is based on the ratiometric quantification using dye-MSNs. Our study provides new insights into the correlation between the EPS matrix, the local external environment and the bacteria’s metabolic activity at the single microcolony level. Using a facile and powerful potential analytical tool for biofilm research, the analysis revealed spatially distinct pH gradients in both the axial (xz) and lateral (xy) directions. The low pH regions and subsequent gradients, which approached a neutral pH towards the biofilm periphery, were manifested as either large acidic clusters at the microcolony core or smaller dispersed acidic pockets found in the interior of the biofilm microcolonies. These observations have both clinical and industrial relevance where the presence of increasing concentrations of acids at/near the biofilm substratum favours the local demineralization of the surface, while the presence of acidic pockets may then also influence the penetration and activity of antimicrobial agents. Understanding of the complex physicochemical properties that the EPS matrix provides to biofilms, and the gradients therein, will allow for the development of exciting antibiofilm strategies that take advantage of these properties. We envision that this ratiometric methodology can be exploited in the future for analysis of various biofilms. ASSOCIATED INFORMATION The supporting information is available free of charge on the ACS Publications website at DOI XX. Biofilm preparation for CLSM, Zeta potentials table, BET and BJH results, IF/IR for different concentrations of dye-MSNs, reversibility of sensing capacity, dye-MSNs penetration into the biofilm and dye-MSNs sorption into the biofilm.


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AUTHOR INFORMATION

Corresponding Author *e-mail: [email protected]

ORCID

Stephanie Fulaz: 0000-0002-0152-2754

Dishon Hiebner: 0000-0001-6764-6170

Caio H. N. Barros: 0000-0002-8151-8703

Henry Devlin: 0000-0003-4467-232X

Stefania Vitale: 0000-0002-7039-9470

Laura Quinn: 0000-0001-6563-3800

Eoin Casey: 0000-0002-6471-7356

Author Contributions ‡These authors contributed equally. All authors have given approval to the final version of the manuscript.


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Funding Sources

This research was supported by Science Foundation Ireland (SFI) under grant number 15/IA/3008.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENT

The authors would like to thank Prof. Dr. Kenneth Dawson from Centre for BioNano Interactions (CBNI) UCD for the Zetasizer equipment.

ABBREVIATIONS Dye-MSNs - mesoporous silica nanoparticles with fluorescein and rhodamine B

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TOC graphic 338x190mm (300 x 300 DPI)



Figure 1 81x23mm (300 x 300 DPI)


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Figure 2 338x190mm (300 x 300 DPI)



figure 3 357x144mm (300 x 300 DPI)


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Figure 4 219x180mm (300 x 300 DPI)



figure 5 366x275mm (300 x 300 DPI)


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Figure 6 219x180mm (300 x 300 DPI)



Figure 7 209x109mm (300 x 300 DPI)


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Ratiometric Imaging of the in situ pH Distribution of Biofilms by use of

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