Î²-CD-Functionalized Microdevice for Rapid Capture and Release of

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β‑CD-Functionalized Microdevice for Rapid Capture and Release of Bacteria Alexandra Perez-Anes,‡,§ Anna Szarpak-Jankowska,‡ Dorothée Jary,§ and Rachel Auzély-Velty*,‡,⊥ ‡

Grenoble Alpes University and Centre de Recherches sur les Macromolécules Végétales, 601, rue de la Chimie, BP 53, 38041 Grenoble Cedex 9, France § Grenoble Alpes University and CEA LETI MlNATEC Campus, 17, avenue des Martyrs, 38054 Grenoble, France ⊥ CNRS and Centre de Recherches sur les Macromolécules Végétales, 601, rue de la Chimie, BP 53, 38041 Grenoble Cedex 9, France S Supporting Information *

ABSTRACT: Most procedures for detecting pathogens in liquid media require an initial concentration step. In this regard, carbohydrates have proven to be attractive affinity ligands for the solid-phase capture of bacteria that use lectins for adhesion to host cell membranes. However, the use of cyclodextrin-immobilized substrates to selectively trap bacteria has not been explored before. Here, using quartz-crystal microbalance with dissipation monitoring experiments, we demonstrate that functionalization of surfaces by β-cyclodextrin (β-CD) can not only allow for rapid and efficient capture of bacterial cells in liquid but also their facile elution with an aqueous solution of a selectively methylated β-CD derivative as a competitive molecule. This capture/ elution strategy, which is based on host−guest interactions between membrane components of the bacterial cell and the CD cavities, is performed in physiological conditions and can be integrated in a microchip. Indeed, proof-of-concept studies showed the potential of β-CD-modified micropillar-integrated microfluidic devices for concentration of bacteria. The results obtained with Escherichia coli suggest that this approach could be broadly applicable among Gram-negative bacteria, which share common cell membrane structures. KEYWORDS: cyclodextrin, bacterial capture, quartz crystal microbalance, microfluidic chip, host−guest interaction

1. INTRODUCTION Rapid and sensitive detection of pathogenic organisms at very low concentrations is crucial to safeguard public health from food- and water-borne pathogens as well as from infectious diseases. Among the rapid pathogen detection technologies, nucleic acid amplification methods such as polymerase chain reaction (PCR) are widely reported. PCR detection is highly specific, can facilitate the identification of microorganisms that are difficult to culture, and has the potential to reduce the overall cost of testing.1 Although a sample (pre)-preparation step is not always necessary for successful PCR amplification,2 it is often required when using environmental or food samples due to the mismatch between high volume sample (>25 mL) and small amplification volume (10−50 μL). Consequently, rapid preconcentration of pathogens is essential to take full advantage of PCR for detecting bacteria. One of the most popular enrichment approaches is based on pathogen capture using antibody-labeled magnetic beads.3 Although very power© XXXX American Chemical Society

ful, this method has the drawbacks of limited lifetime and high cost associated with the antibody, which makes it unsuitable for routine screening of environmental or food samples. Thus, there is a need for alternative platforms for the rapid, simple, and efficient capture of bacteria. It is known that surfaceexposed proteins and lipopolysaccharides in bacteria are responsible for important functions, including adhesion and virulence. This fact, together with the ability of cyclodextrins to selectively interact with cellular membranes by virtue of either their complexation ability and/or their surface activity,4 prompted us to use these cage molecules as synthetic receptors to design a simple and cost-effective bacterial capture platform. Here, we demonstrate that functionalization of a micropillarintegrated microfluidic device by β-cyclodextrin (β-CD) can Received: February 14, 2017 Accepted: April 10, 2017 Published: April 10, 2017 A

DOI: 10.1021/acsami.7b02194 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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Figure 1. (A) Experimental setup for real-time qRT-PCR detection of bacterial cells from aqueous samples. (B) Scheme of bacterial capture/elution using β-CD-modified substrates and a methylated β-CD derivative. (EDC), (3-aminopropyl)triethoxysilane (APTS), cysteamine, succinic anhydride, and dNTP (deoxynucleotide set, 100 mM) were purchased from Sigma-Aldrich-Fluka and were used without further purification. Bovine serum albumin (Ultrapure BSA 50 mg/mL) and TaqGold polymerase and buffer (AmpliTaq Gold DNA Polymerase, ABI) were from Life Technologies. Taqman primers and probe were purchased from Eurofins MWG Operon. Zirconia/Silica Beads were from BioSpec Products. Bacterial strains were purchased from LGC Standards. The water used in all experiments was purified by a Millipore Milli-Q Plus purification system, with a resistivity of 18.2 MΩ cm. 2.2. Synthesis of Mono-6-amidosuccinyl-6-deoxy-cyclomaltoheptaose (β-CD-COOH). To a vigorously stirred solution of 6monodeoxy-6-monoamino-β-cyclodextrin hydrochloride (300 mg, 0.2560 mmol) in water (30 mL), 1 M NaOH (282 μL, 0.2820 mmol) was dropped at 0 °C. The mixture was kept at 0 °C and stirred for 15 min. After solvent removal (lyophilization), the resulting white powder βCD-NH2 was used in the next step without further purification (290 mg, 99%). To a vigorously stirred solution of βCD-NH2 (290 mg, 0.2560 mmol) in dry dimethylformamide (DMF; 7 mL) at room temperature a solution of succinic anhydride (28 mg, 0.2820 mmol) in dry DMF (2 mL) was added. The mixture was stirred for 45 min at room temperature. After evaporation of most of the solvent, the residual syrup was poured into acetone (100 mL). The white precipitate was collected by filtration, washed with acetone (3 × 100 mL), and dried to afford β-CD-COOH as a white powder (282 mg, 97%). 1H NMR (400 MHz, D2O, δ): 5.10 (m, 7H, H1 of β-CD), 4.03−3.90 (m, 28H, H2, H3, H4, and H5 of β-CD), 3.69−3.62 (m, 12 H, H6 of β-CD), 3.45 (t, J = 8 Hz, 1H, H6 of the modified glucose unit of β-CD), 3.35−3.29 (dd, J = 16 Hz, J = 8 Hz, 1H, H6 of the modified glucose unit of β-CD), 2.61−2.54 (m, 4H, CH2−COOH, NH−CO−CH2). 13C NMR (100 MHz, D2O, δ): 177.9 (CO), 174.9 (CO), 101.7 (C1 of β-CD), 83.05, 81.1, 72.9, 72.2, 71.7 (C2,

not only allow for rapid and efficient capture of bacterial pathogens in liquid but also their facile elution in mild conditions, which does not inhibit the subsequent PCR amplification. Moreover, since the capture is mainly governed by reversible binding with CD molecules, the microfluidic chip may be used for detection of different bacteria and reused. Microfluidic technology was used because of its applicability to sorting and enrichment of cells under continuous flow.5 The setup for the bacterial capture and elution developed here (Figure 1) relies on selective interactions between the bacterial cell envelope and β-CD molecules immobilized on the micropillar-structured surface. The latter interactions play a key role in regulating the surface attachment/detachment of bacterial cells. The large surface/volume ratio of the micropillar array offers more contact area for grafting CD molecules, thereby enhancing capture. The geometrical design of the micropillar array (200 μm height by 20 μm in diagonals; space between the pillars: 20 μm) was previously optimized to reduce clustering of the cells.

2. MATERIALS AND METHODS 2.1. Materials. β-Cyclodextrin was kindly provided by ROQUETTE (Lestrem, France). 6-Monodeoxy-6-monoamino-β-cyclodextrin (β-CD-NH2) was purchased from AraChem (Netherlands). NHydroxysulfosuccinimide sodium salt (sulfo-NHS) was purchased from Chemrio (Ningbo Zhejiang, P.R. CHINA). Heptakis (2,6-di-Omethyl)-β-cyclodextrin (DIMEB), randomly methylated β-CD (RAMEB, degree of methylation = 1.6−2), tris(hydroxymethyl)aminomethane hydrochloride, (TRIS-HCl), 2-(N-morpholino)ethanesulfonic acid (MES), phosphate buffer saline (PBS, pH 7.4), N-ethyl-N′-(3-(dimethylamino)propyl)carbodiimide hydrochloride B


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ACS Applied Materials & Interfaces C3, C4, C5 of β-CD), 60.2 (CH2−OH of β-CD), 40.2 (CH2−NH of β-CD), 30.6 (CH2−COOH), 30.4 (CH2−CONH). 2.3. Bacteria and Culture Conditions. Bacterial strains used in this study, namely, the Escherichia coli (ATCC 9637), Staphylococcus epidermidis (ATCC 14990), Serratia marcescens (ATCC 27137), and Bacillus subtilis (ATCC 23857) strains, were obtained from LGC Standards. E. coli and S. epidermidis were grown in liquid LB Broth (Miller) (Sigma-Aldrich, ref L2542) at 37 °C overnight. S. marcescens was grown in liquid medium Tryptic Soy Broth (Sigma-Aldrich, ref 22092) at 26 °C overnight. B. subtilis was first grown in liquid Tryptic Soy Broth at 30 °C overnight. Then spores were prepared by spreading 0.2 mL of culture samples on agar plate and incubating at 30 °C for one week in slightly modified fortified nutrient agar (FNA)6 containing beef extract, 3 g; tryptone, 5 g; NaCl, 3 g; glucose, 0.1 g; agar, 20 g; CaCl2, 0.06 g; modified Gb mineral solution (containing MnSO4·H2O, 0.05 g; MnSO4·H2O, 0.05 g; (NH4)2SO4, 0.08 g; MnCl2· 4 H2O, 0.008 g; CuSO4·5 H2O, 0.005g; ZnSO4·7 H2O, 0.005g); MilliQ water, and NaOH for a final volume of 1 L and a pH adjusted to 7). Spores were suspended to appropriate concentration after they were washed three times with DNase, RNase, Protease free water (W4502, Sigma-Aldrich) and counting by using Quick-Read-Ten Chamber Microscope Slide (Globe Scientific Inc.) with an optical microscope (Axioplan 2 imaging Zeiss). The absence of vegetative cells is checked by direct observation with optical microscopy and also by comparing the threshold Cycle (Ct) obtained by qPCR of untreated spores suspension and of mechanically lysed spores suspension at the same concentration using Precellys24 (Bertin Technologies). The number of Ct difference between spores and lysed spores must be at least of five cycles indicating a contamination of less than 3% by vegetative cells or free DNA. The suspension of each type of bacteria is then prepared freshly prior to experiments (QCM-D or concentration with microdevice) by washing twice the bacteria obtained after overnight culture with Milli-Q water. The bacteria concentration in final suspension is then determined by microscope counting method using disposable slides for cell counting (Fast Read 102, BioSigma). 2.4. Quartz Crystal Microbalance with Dissipation Monitoring Measurements. QCM-D measurements were performed using Q-Sense E1 or E4 instruments (Biolin Scientific, Sweden) equipped with one or four flow modules, respectively. Besides the measurement of bound mass related to changes in the resonance frequency f of the sensor crystal, the QCM-D technique also provides structural information on biomolecular films via changes in the energy dissipation D of the sensor crystal. Values of f and D were measured at the fundamental resonance frequency (5 MHz) as well as at the 3rd, 5th, 7th, 9th, 11th, and 13th overtones (n = 3, 5, 7, 9, 11, and 13). Experiments were conducted either in a continuous flow of buffer with a flow rate of 50 μL/min by using a peristaltic pump (ISM935C, Ismatec, Zurich, Switzerland) or in batch mode. All buffers were previously degassed to avoid bubble formation in the QCM-D measuring chamber. Prior to use, the gold-coated crystal sensors (Q-Sense QSX 301) were exposed to a UV−ozone treatment for 10 min using a UV− ozone cleaner (Jelight Company) and immersed in ethanol under stirring for 20 min. The surfaces were then dried under nitrogen before dipping overnight in the mixture comprising 1 mM ethanol solution of cysteamine, then carefully rinsed with ethanol and dried under nitrogen. The functionalized gold quartz crystal was then mounted in the QCM-D flow module, and the formation of the β-CD layer was monitored in the presence of MES buffer (pH 4.75) as the running buffer. The transducer surface was then exposed to 1 mL of mixture solution of β-CD-COOH (2, 10, or 20 g/L), EDC (50 g/L), and sulfoNHS (30 g/L) for 4 h in batch mode. The resulting functionalized surface was rinsed under flow until reaching the stabilization of the QCM-D signals. For bacteria-adhesion assays, the flow module surfaces were passivated by using BSA at 10 g/L for 30 min in batch mode. After they were rinsed with 10 mM TRIS (pH 7), the quartz crystal sensors were equilibrated in the QCM-D measurement chamber with 10 mM TRIS (pH 7). f and D were recorded continuously during the incubation (using a flow rate of 50 μL/min) of the cells (1 × 107 cells/

mL) suspended in the same medium until reaching stable signals. Then the measurement chambers were rinsed with 10 mM TRIS (pH 7), and the binding of the bacteria to the QCM-sensor surface was monitored by the changes in f and D. Experiments were performed in triplicate. Microscopy experiments coupled with QCM-D were performed using Q-Sense Window module 401 (Biolin Scientific, Sweden) and a microscope Axio Imager A1m (Carl Zeiss S.A.S., France). The images were registered and treated using the software Axiovision from Carl Zeiss S.A.S. As β-CD layer formed soft films (dissipation shift ΔD > 0), the adlayer was modeled as a homogeneous layer of thickness dQCM and density ρ, using a continuum viscoelastic model.7 The shear elastic modulus G′ and the shear loss modulus G″ represent the layer viscoelastic properties. The frequency dependence of the viscoelastic properties was approximated by power laws with exponents α′ and α″ ⎛ f ⎞α ′ ⎛ f ⎞α ″ G′(f ) = G0′⎜⎜ ⎟⎟ , G″(f ) = G0″⎜⎜ ⎟⎟ ⎝ f0 ⎠ ⎝ f0 ⎠

(1)

where G0′ and G0″ are the shear storage and loss moduli at the (arbitrarily chosen) reference frequency f 0 = 15 MHz. QCM-D data (at all available overtones) at selected time points were fitted with the software QTM8,9 (see Supporting Information for related information). ρdQCM, ρG0′, ρG0″, α′, and α″ were adjustable fitting parameters. The semi-infinite bulk solution was assumed to be Newtonian with a viscosity ηl = 0.89 mPa s and a density of ρl = 1.0 g/cm3. Postulating that the film is highly hydrated, its density ρ was also fixed to 1.0 g/ cm3. Details of the fitting procedure have been described previously.10 2.5. Atomic Force Microscopy (AFM). AFM measurements were performed with a Bruker AFM, “AFM_Dimension1” (ICON Head), under ambient conditions. Silicon cantilevers Bruker NCHV were used for tapping mode operation. Data treatment, display, and analysis were performed using Bruker “Nanoscope Analysis 1.7” & Image Metrology A/S “Scanning Probe Image Processing 6.3.3”. 2.6. Functionalization of the Flat Silicon Substrates for AFM Studies. Flat silicon substrates were first sonicated in a brown solution (1.4 g of NaOH in a water/EtOH mixture (150 mL/200 mL)) for 30 min in an ultrasonic bath. Next, the substrates were rinsed with water and dried by centrifugation (30 min, 1000 rpm). A layer of resin (JSR 335 positive photoresist available from JSR Corporation) was manually applied with a brush on half of the flat silicon surface and heated during 1 min at 120 °C. Silanization with APTS was performed from the gas phase by placing the samples inside a dry-seal Teflon pot containing 600 μL of the silane. Then, the samples were kept in an oven for 1 h at 60 °C. The excess of unreacted alkoxysilane and the resist was removed by rinsing and sonication in acetone (5 min) and ethanol (5 min). Substrates were then dried by centrifugation (1000 rpm, 30 min) and annealed for 3 h at 110 °C. In the next step, the amine-functionalized silicon substrates were incubated in a mixture of a β-CD-COOH solution (2, 5, or 10 g/L in 0.2 M MES (pH 4.75) containing EDC and sulfo-NHS (same conditions as for the grafting of β-CD moieties on the QCM crystal sensors). They were stirred using an orbital shaker at room temperature for 4 h. The resulting CDfunctionalized flat silicon substrate was rinsed with MES buffer and dried with N2. 2.7. Isothermal Titration Calorimetry. ITC experiments were performed on a Microcal VP-ITC titration micro-calorimeter (Northampton, USA). All titrations were made in PBS buffer pH 7.4 at 25 °C. The reaction cell (V = 1.45 mL) contained the cyclodextrin derivative solution. A series of 30 injections of 10 μL from the computercontrolled 300 μL microsyringe at an interval of 10 min of sodium adamantane acetate (ADAc) were performed into the CD derivative solution with stirring at 300 rpm at 25 °C. The raw experimental data were reported as the amount of heat produced after each injection of ADAc as a function of time. The amount of heat produced per injection was calculated by integration of the area under individual peaks by the instrument software, after taking into account heat of dilution. The experimental data were fitted to a theoretical titration curve using the instrument software (Bindworks program (CSC) or C


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ACS Applied Materials & Interfaces Table 1. Probes and Primer Sequences strain E. coli

S. epidermidis

B. subtilis

S. marcescens

primer/probe

sequence (5′→3′)

Ec-F Ec-R Ec-probe Se-F Se-R Se-probe Bs-F Bs-R Bs-probe Sm-F Sm-R Sm-probe

GTGTGATATCTACCCGCTTCGC AGAACGGTTTGTGGTTAATCAGGA FAM- TCGGCATCCGGTCAGTGGCAGT-BHQ1 TCTTAAGAATGTTACAAGTGGTGCAA AATCTCATGGAGCGCTTCTATAGC HEX-TTGCACTGCTTTGTCAATACCTTGTCTTAAGCCT-BHQ1 GACCGATCAGCTTGTAGAAGTTG TTGAGATGCCTGCATGGATG Cy5- TGCAGTTTACAGCACAGCTCGAGAAAT-BHQ2 AGTGCACGAACAAACTTACAG GTCGTACTCGAAATCGGTCACA Cy5-TTCTGGCCAAGCCACCAGACCTTTAC-BHQ2

the grafting of β-CD moieties on the QCM crystal sensors to graft βCD-COOH on the amine-functionalized pillar arrays. The substrates were incubated in a mixture of a β-CD-COOH solution (2, 5, or 10 g/ L in 0.2 M MES (pH 4.75)) containing EDC and sulfo-NHS. They were stirred using an orbital shaker at room temperature for 12 h. Then, they were rinsed with buffer for 5 min and dried by centrifugation (30 min, 1000 rpm). Packaging of chips. Once the chemical functionalization was completed, a glass cover was glued on top of the chip by screen-printing for glue deposition. Capillaries were glued in input and output to the microfluidic test. 2.10. Bacterial Cell Capture and Release Tests Using the Pillar Array Chips. First of all, a sample containing microorganisms (0.1, 0.3, or 1 mL) at a known concentration (1 × 103 cells/μL) in Milli-Q water (pH 6) was injected with a classical micropipette in a plastic card having a functionalized β-cyclodextrin pillar chip (flow 16.0 μL/s). A micropipette cone was inserted at the outlet of a channel to recover the captured supernatant. Then, 30 μL of elution solution (10 mM TRIS buffer, pH 7, containing DIMEB ([DIMEB] = 540 g/ L)) was injected by the input channel (flow 2.5 μL/s). The eluate was recovered in the exit cone and put in an Eppendorf. A small amount of glass beads and 1 μL of BSA at 10 mg/mL were added. The mixture was placed in Precellys24 to achieve the mechanical lysis. Then, the lysed mixture was centrifuged a few seconds to separate the beads and the eluate. The supernatant containing the DNA now was recovered and placed in a PCR tube to be analyzed by qPCR after adding PCR Mix in the tube. At least three independent experiments per condition were performed. 2.11. Quantitative Real-Time PCR Amplification. For real-time PCR assays using TaqMan probe (Eurofins MWG Operon), PCR mixtures possessed 0.25 U/μL Taq polymerase (AmpliTaq Gold DNA Polymerase, ABI), AmpliTaq Gold Buffer 1X, 3 mM MgCl2, 0.2 mM d NTP mixture, 0.65 M betaine, DNase and RNase free water, and a set of 0.3 μM forward and reverse primers, and 0.3 μM TaqMan probe. The PCR amplicon was designed to have ∼100 base pairs for all strains. After the sample was loaded into the PCR analyzer (Stratagene Mx3005P), it was heated at 95 °C for 10 min to denature DNA templates followed by 40 cycles of a denaturation step at 95 °C for 15 s, annealing and extension step at 60 °C for 30 s. As the volume of samples was limited after mechanical lysis (∼15−20 μL), duplexes PCR were performed: Sm (Cy5 probe) and E. coli (FAM probe), and Bs (Cy5 probe) and Se (HEX probe). A standard curve for each strain with pure DNA was performed for all analysis runs. After PCR amplification was completed, the amplified DNA from samples was quantified using MxPro software. The probes and primer sets used in this study are listed in Table 1.

ORIGIN software (Microcal)), refining the enthalpy change (in kJ/ mol) ΔH, the association constant (in M−1) Ka, and the stoichiometry of the interaction (number of binding sites per receptor) n. In all cases, calculations were performed using the “one set of binding sites” model. 2.8. Microdevice Fabrication. The chips with a very highly developed surface made of a perfectly ordered two-dimensional array of micropillars directly etched in silicon were made at CEA-LETI (Grenoble, France). Microfabrication of silicon pillar chips is briefly described. All photolithography steps were performed with standard microelectronics machines. An initial silicon oxide (SiO2) insulating layer of 1 μm is grown by thermal wet oxidation on 200 mm silicon wafer. Then successive steps comprise photoresist deposition, photolithography, etching, and further Deep reactive-ion etching (DRIE) leading to the high aspect ratio pillar chip. A final thermal wet oxidation is performed. Each pillar has a dimension of 20 μm in diagonals, 200 μm in height, and an interpillar spacing of 20 μm. The micropillar silicon chip has a volume of 15.2 μL, and the contact surface with the sample was designed to be as large as possible (13 cm2), that is, with a small distance between pillars to increase the developed surface but limiting this spacing to avoid very high pressures in the device, which can lead to clogging with some samples or also to leakages by breaking the glue film between microchip and cover. Preliminary experiments have shown that an interpillar spacing of 20 μm satisfies these criteria. This significant contact surface/volume ratio is highly favorable for efficient capture of bacteria, as it increases the probability that a bacteria is close enough from solid surface for capture. The chip was glued on a plastic card containing channels for liquids injection and recovery after flowing through the pillars array. These channels were directly connected to the inlet and outlets of the chip by geometrical alignment. Holes at the entry and outlet of each fluidic circuit were machined so that a pipet cone can be inserted for convenient liquid injection and recovery. At the entry of the microchip, the inlet stream was split by channels successive divisions just prior to the entry in the pillars array to enable a good distribution of the fluid in the pillars chamber. The output had the identical design, and the flow was grouped before flowing in the exit channel of the plastic card and entering the outlet pipet cone for recovery. 2.9. Functionalization of the Pillar Array Chips. Surface Activation. Flat substrates and silicon pillar array chips were first sonicated in a brown solution (1.4 g of NaOH in a water/EtOH mixture (150 mL/200 mL)) for 30 min in an ultrasonic bath. Next, the substrates were rinsed with water and dried by centrifugation (30 min, 1000 rpm). Silanization with APTS: After surface activation with brown solution, silanization with APTS was performed from the gas phase by placing the samples inside a dry-seal Teflon pot containing 600 μL of the silane. Then, the samples were introduced into an oven at 60 °C overnight. Afterward, the substrates were thoroughly rinsed and sonicated in ethanol for 5 min in an ultrasonic bath to remove all noncovalently bound silane and dried by centrifugation (30 min, 1000 rpm). Finally, the substrates were annealed at 110 °C for 3 h. Covalent attachment of β-CD moieties: the same procedure was used as that for

3. RESULTS AND DISCUSSION 3.1. Investigation of β-CD-Modified Surfaces for Capture and Release of E. coli by QCM-D. To evaluate the bacterial capture and elution capabilities of β-cyclodextrin modified surfaces, we first examined the binding of E. coli as a D


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ACS Applied Materials & Interfaces representative Gram-negative bacterium to β-CDs by quartz crystal microbalance with dissipation monitoring. To this end, β-cyclodextrin monofunctionalized with a carboxylic acid group (β-CD-COOH) was immobilized on the gold-coated QCM crystal surface functionalized with 2-aminoethanethiol by an amine-acid coupling reaction. Figure 2A compares the

Table 2. β-CD Layer Thickness Evaluated from QCM-D Data Treatment Using QTM Modeling and by AFM QCM-D

AFM

[β-CD] g/L

dbest fit (nm)

dmin (nm)

dmax (nm)

Δz (nm)

2 10 20

4.2 6.6 6.2

3.4 4.8 3.6

12.1 16.8 21.5

2.8 ± 0.1 2.9 ± 0.2 4.2 ± 0.3

silicon substrates, prepared from the different β-CD-COOH concentrations (Table 2 and Figure 3). Slightly lower values

Figure 3. (A) AFM image of β-CD layer covalently bound to silicon substrate and (B) height histogram of the sample. Concentration of βCD-COOH used for the grafting: 10 g/L.

Figure 2. QCM-D profile (shifts in resonant frequency (A) and in dissipation (B) vs time) illustrating the grafting of β-CD-COOH on quartz crystal gold surfaces functionalized with 2-aminoethanethiol (seventh overtone data). Conditions: 0 s, buffer flow (0.02 M MES pH 4.75); 389 s, activation of surface and immobilization of β-CD-COOH (50 g/L EDC, 30 g/L sulfo-NHS and 2, 10, and 20 g/L of βCDCOOH, respectively, in 0.02 M MES pH 4.75); 6600 s, rinsing with buffer.

ranging from 2.8 to 4.2 nm in comparison to those derived from QCM-D were obtained, which is in line with values reported in the literature. Values ranging from 0.4 to 5.4 nm were indeed obtained when CDs were immobilized via different chemical linkers.11−13 The higher QCM-D values could be attributed to the precision of fitting. On the basis of these results, the bacterial capture of E. coli on the QCM sensor surfaces was investigated on β-CDmodified surfaces using a β-CD-COOH concentration of 10 g/ L. After equilibration of the QCM sensors with TRIS buffer at pH 7, 500 μL of 1 × 107 bacterial cells/mL (50 μL/min flow rate) were passed, and changes in frequency as well as in dissipation shifts were monitored until it became stable (∼5 min). Finally, TRIS buffer (pH 7) was introduced to wash unbound bacteria. As can be seen from Figure 4, there was immediate and rapid adsorption of bacterial cells onto the functionalized sensor crystal, resulting in a decrease of frequency. Bacteria remained attached to the surface after rinsing, as no significant change in the frequency was observed after introduction of the buffer (time t2 in Figure 4). Regarding dissipation, positive shifts were observed after the rinsing step, which indicates that a mobile bacterial layer is created on the βCD-coated surface. Similar QCM-D profiles were previously observed for E. coli adsorbed to glycosylated QCM sensor crystals.14 The stability of the QCM signals (f and D) after rinsing suggests that the adsorbed bacteria are strongly anchored to the β-CD-coated surface through host−guest interactions between membrane components of the bacterial cell and the CD cavities. Indeed, the trapping of bacteria proceeds readily due to the multiple presentation of cellular components, potentially

frequency shifts recorded after injection of solutions containing β-CD-COOH at different concentrations (2, 10, and 20 g/L) in MES buffer (pH 4.75), the water-soluble carbodiimide, 1-ethyl3-(3-(dimethylamino)propyl)carbodiimide hydrochloride (EDC), and N-hydroxysulfosuccinimide (sulfo-NHS) on the amine-functionalized QCM crystal surface. After ∼2 h reaction and rinsing steps with MES buffer, we observed significant negative frequency shifts, corresponding to the covalent fixation of β-CD on the surface. Note that the shifts in resonant frequency and in dissipation for each coupling condition (βCD-COOH concentration) after the rinsing steps did not change by varying the time of reaction between 2 and 12 h (one night), indicating that the amine−acid coupling reaction was completed within 2 h. The frequency and dissipation shifts were found to be dependent on the concentration of β-CDCOOH in the injected solution due to the increase in viscosity of the solutions of increasing concentrations (Figure 2A,B). These QCM-D data (shift in resonance frequency Δf and in dissipation ΔD measured at different overtones after the rinsing steps) were used to evaluate the thicknesses of the chemisorbed β-CD layers. The thicknesses determined from the best fit of the QCM-D data were found to be similar for the three CD concentrations, 2, 10, and 20 g/L (dbest fit in the range of 4.2− 6.6. nm, Table 2). AFM experiments were also performed to measure the thickness of the CD layers in the dried state on E


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Figure 4. QCM-D profile (shifts in resonant frequency (A) and in dissipation (B) vs time) illustrating the capture of bacterial cells (E. coli) on a βCD-sensor crystal prepared with [β-CD-COOH] = 10 g/L and their elution with a solution of DIMEB at 540 g/L in TRIS buffer (seventh overtone data). (C) Optical images of the QCM-sensor surface showing bacterial capture and release. t0 = TRIS buffer (pH 7); t1 = bacteria at 1 × 107 cells/ mL in TRIS buffer; t2 = buffer solution; t3 = DIMEB at 540 g/L in TRIS buffer; t4 = buffer solution.

complexes with DIMEB at high concentration (230 g/L) and washings with water, bacteria lost completely the fluorescence (Figure S1C). In contrast, no desorption was observed after injection of native β-CD at a concentration near its maximum solubility (18 g/L) in water (Figure S2). Likewise, injection of DIMEB at a concentration of 18 g/L did not result in the release of bacteria from the surface, even after a series of two additional injections at the same concentration (Figure S3). These experiments confirmed our earlier statement that bacteria are bound to the β-CD-coated in a strong but reversible fashion through multivalent interactions. Indeed, a concentration of more than 1 order of magnitude higher than the maximum solubility of natural β-CD is required to truly compete with the multiple presentation of β-CD offered by the quartz crystal surface. It is worth noting that these conditions allow rapid and quantitative bacteria recovery in mild conditions at neutral pH. As the complexation properties of the β-CD cavity are strongly dependent on the regioselectivity as well as on the degree of methylation of the glucose units, in a subsequent experiment, we attempted to elute bacteria using another highly water-soluble methylated β-cyclodextrin, namely, randomly methylated β-CD (RAMEB, degree of methylation = 1.6−2). Because of partial methylation of the OH (3) groups of the βCD rim, the complexation properties of this methylated β-CD may be altered as previously reported for heptakis(2,3,6-tri-Omethyl)-β-cyclodextrin (TRIMEB).17 This was checked by isothermal titration calorimetry (ITC) studies of RAMEB, DIMEB, and β-CD complexes with sodium adamantane acetate

acting as guests for the CD cavity, which induces a high affinity to the surface. To confirm the selective interaction between bacteria and the surface, a solution of competing heptakis(2,6-di-O-methyl)-βcyclodextrin (DIMEB) in TRIS buffer (pH 7) was injected after bacterial adsorption. This methylated CD was selected due to its high solubility in water allowing to prepare more concentrated solutions than with the parent β-CD. It is indeed well-described in the literature that methylation of β-CD results in a dramatic increase in water solubility, because methylation reduces formation of intermolecular hydrogen bonds.15,16 DIMEB has thus an aqueous solubility 30 times higher than that of the parent β-CD (560 vs 18.5 g/L)15 and still retains efficiency for complexation.17 When a solution of DIMEB at a concentration near its maximum solubility in MES buffer (540 g/L) was passed on the QCM sensor with E. coli bound on the surface modified with β-CD-COOH, the bacterial cells were fully released from the surface (Figure 4A,B). Notably, the kinetics of bacterial desorption was very rapid. Optical image of the QCM−sensor surface confirmed effective bacterial release on the β-CD-coated surface (Figure 4C). When bacteria on the surface are exposed to a solution of free DIMEB, competition between the cellular components/surface CD sites interactions and the cellular components/DIMEB interactions occur, resulting in desorption of bacteria. This release of β-CD by DIMEB was also illustrated by additional experiments using fluorescently labeled β-CD. After incubation of E. coli bacteria with the fluorescent β-CD (0.75 g/L), bacteria became fluorescent even after intense washings with water (Figure S1A). Following incubation of the fluorescent β-CD/bacteria F


Research Article


accessible aromatic side chains on the surface of noncarbohydrate-binding proteins (insulin SerB9Asp and S6).30 Interestingly, tryptophan and tyrosine are the only hydrophobic residues that are found with equal frequency in the interior and on the surface of proteins.31 Thus, β-CD and DIMEB are likely to interact with proteins exposed on the surface by inclusion complexation. Several crystal structures of carbohydrate-binding proteins in complex with β-CD also indicate interactions with aromatic amino acids,30 but they occur through nonpolar stacking between aromatic amino acids and part of the CD sugars. Hydrogen bonds between the glucose units and nonaromatic amino acids also play a key role in the CD binding to the protein.32 This is the case of the periplasmic maltodextrin binding protein (MBP), which serves as an initial receptor for the active transport of malto-oligosaccharides in E. coli.33,34 Importantly, this protein acts in conjunction with the outer membrane λ receptor protein to facilitate specifically the transport of maltodextrins across the outer membrane. This means that maltodextrins are bound at the outer membrane of E. coli by the λ receptor, which has been reported to bind αcyclodextrin but with a lower affinity than MBP.34 Thus, bacterial capture by β-CD may be partly due to their selective/ specific interaction with several surface-exposed proteins, including the λ receptor protein. Regarding elution by DIMEB, the regioselective methylation of the OH (2) and OH (6) of this cyclodextrin probably makes it more prone to selectively interact with surface-exposed proteins via hydrophobic interactions and hydrogen bonding in comparison to RAMEB. 3.2. Efficiency for Capture and Release of Different Species Bacteria by β-CD-Modified Surfaces. On the basis of these results, we investigated whether the proposed platform may be potentially used for the capture of other bacterial strains. To evaluate this, adsorption on β-CD-coated QCM-D crystal substrates of two Gram-positive bacteria (B. subtilis and S. epidermidis) as well as another Gram-negative bacterium (S. marcescens), which belongs to the family Enterobacteriaceae as E. coli, was investigated. Figure 5 shows QCM-D measurements following injection of 500 μL of each bacterial cell (1 × 107 cells/mL; 50 μL/min flow rate). For clarity, only the data recorded at one overtone (n = 7) are shown, while the four other overtones (n = 3, 5, 9, 11) exhibited the same behavior. For all bacterial strains, negative frequency shifts are observed, indicating their adsorption on the QCM-D substrate. The most significant frequency change is obtained for the Gram-negative bacterium. At the same time, an increase of dissipation is observed, which is associated with increased viscosity of the adsorbed layer. Although a dissipation change is observed for S. epidermidis, the frequency shift is relatively small, which may indicate little adsorption of this Gram-positive bacterium. The significant dissipation shift observed can be ascribed to the property of S. epidermidis to rapidly adhere to surface to form viscous films. Regarding the adsorption of the Gram-positive bacterium B. subtilis, this exhibits an intermediate profile in comparison to those of S. marcescens and S. epidermidis. Importantly, quantitative desorption of bacterial cells was observed for the three strains upon elution with a solution of DIMEB at a concentration of 540 g/L in TRIS buffer (pH 7; Figure S6). Further comparison of the frequency and dissipation shifts at the different overtones obtained for these bacteria and for E. coli shows that bacterial binding on the β-CD-coated surface is higher for the Gram-negative bacteria (Figure 6). Of these

(ADAc, Figure S4). From the three types of cyclodextrin tested, RAMEB exhibited the lowest affinity toward ADAc. Indeed, the stability constant of the ADAc/RAMEB complex (Ka = 10 500 M−1) was found to be 1.4-fold lower than the ADAc/DIMEB complex (Ka = 14 500 M−1), while the stability constant of the latter complex was reduced by a factor of 6 relative to the ADAc/β-CD complex (Ka = 88 100 M−1). When a solution of RAMEB at the same concentration of DIMEB (540 g/L) in the elution solvent was passed on the QCM sensor with E. coli bound on the surface modified with β-CD-COOH ([β-CDCOOH] = 10 g/L in the injected reaction medium), no desorption of bacterial cells was observed (Figure S5). Unlike the native β-CD, its methylated derivatives are surface-active and soluble in organic solvents.18 Thus, a proper combination of a complexation power, surface activity, and lipophilicity may enable DIMEB to interact with the bacterial cell membrane and the wall effectively. Note, however, that DIMEB was previously reported to be more efficient to interact with the membrane of E. coli, while TRIMEB showed no significant interaction though it is surface-active similar to DIMEB.19 Moreover, in the present study, optical microscopy observations revealed that the bacterial cells are still alive after elution with the concentrated solution of DIMEB (based on plate-count experiments, data not shown), which rules out cell lysis. Collectively, these results suggest that the potency of DIMEB-induced release of bacteria is related to its selective interaction with some components of the outer bacterial membrane combined with its high watersolubility. Different types of interactions between β-CD and its methylated derivatives and cell membranes have already been described in the literature. DIMEB similarly to β-CD can potentially interact with cholesterol, phospholipids, and/or proteins in the membrane.4 β-CD and its methylated derivatives have been reported to remove cholesterol from cell membrane.4 This process has been attributed to the ability of cholesterol molecules to diffuse directly from the plasma membrane into the hydrophobic core of a cyclodextrin molecule packed near the membrane surface.20 Yet, as two stacked β-CD are necessary to completely include cholesterol,21,22 full extraction of cholesterol seems rather unlikely, at least for the β-CD molecules immobilized on the surface. As observed for cholesterol, CDs are also able to remove phospholipids from membranes.23 Such an extraction of phospholipids has been proposed to cause the solubilization of lipid vesicles.24−27 It was shown that methylation of β-CD resulted in a much stronger interaction with lipids compared to TRIMEB and β-CD.26 Although models for the interaction of CDs with lipid membranes have been proposed,28,29 the molecular mechanisms of the process of lipid extraction are not understood. It is unclear how the extravesicular CDs get access to the fatty acyl chains buried within the bilayer core. Indeed, contrary to cholesterol, which exhibits large slipping mobility resulting in its protrusion out of the membrane surface, such a slipping mobility perpendicular to the membrane surface is energetically unfavorable for long-chain phospholipids and should only occur with a very low probability.23 So, although these data may provide some evidence for the efficacy of DIMEB to release bacterial cells from the surface, they provide few clues as to why β-CD immobilized on the surface can effectively capture E. coli. Regarding proteins, besides forming inclusion complexes with noncarbohydrate-binding proteins, βCD can specifically interact with carbohydrate-binding proteins by a molecular recognition mechanism. Formation of inclusion complexes with β-CD has been reported to occur with solventG


Research Article


Figure 7. Energy dissipation (ΔD) vs frequency (Δf) dependence of the binding of Gram-positive and Gram-negative bacteria at the 3rd (A) and 11th (B) overtones.

Figure 5. QCM-D profile (shifts in resonant frequency (A) and in dissipation (B) vs time) illustrating the capture of Gram-positive and Gram-negative bacteria on β-CD-sensor crystals prepared with [β-CDCOOH] = 10 g/L (seventh overtone data). t0 = TRIS buffer (pH 7); t1 = bacteria at 1 × 107 cells mL−1 in TRIS buffer; t2 = buffer solution.

the influence of the bacterial adsorption on the viscoelastic damping of the crystal resonance and thus infer viscoelastic properties of the adsorbed layer. For instance, a low ΔD/Δf value indicates mass addition without significant dissipation increase, which is characteristic of a fairly rigid layer. In contrast, a large ΔD/Δf value signals a soft, dissipative film. Figure 7 compares the ΔD/Δf plots for the adsorption of the four bacteria strains onto the β-CD-coated QCM crystal surfaces at the 11th and 3rd overtones, which probe processes closer to the sensor surface and the furthest away, respectively. In the case of E. coli, S. marcescens, and S. epidermidis, there is a clear linear relationship between the change in dissipation and the change in frequency during bacterial adsorption on the surface. The slope of ΔD/Δf is much steeper for the binding of S. epidermidis than for the other bacteria, confirming our previous speculation that S. epidermidis adsorbs in a very viscous layer. E. coli adsorption consist of two steps: an initial step exhibiting mass addition without significant dissipation decrease, and a second step that induces a very steep ΔD/Δf slope. This reflects rapid initial attachment of bacterial cells on the β-CD-coated surface followed by changes in the contact area between the adhering cells and the surface, which is accompanied by more trapped water. For B. subtilis, a markedly different behavior on surface is observed: the ΔD/Δf plot is initially linear with a tendency to level off at high coverage, indicating some rearrangement in the bacterial film resulting in a stiffer bacterial film. Although this Gram-positive bacterium is surrounded by a rigid peptidoglycan cell wall that protects it from its environment, these results indicate its capability to bind β-CD. Interestingly, monitoring growth of B. subtilis on minimal medium containing either α-, β-, or γ-cyclodextrin as the sole carbon and energy source revealed that B. subtilis is able to grow preferentially on β- or γ-cyclodextrins.36 This has been related to the existence in the B. subtilis genome of a gene encoding a protein potentially involved in cyclodextrin utilization. This protein was found to specifically interact with

Figure 6. Comparison of the shifts in resonant frequency and in dissipation resulting from the binding of Gram-positive and Gramnegative bacteria to the β-CD-coated surface ([β-CD-COOH] = 10 g/ L; seventh overtone data).

latter bacteria, E. coli exhibits more propensity for adsorbing on the surface. This fact allows us to speculate that the cell wall (which envelopes cellular membranes) can provide bacterial cells with some protection against cyclodextrins, but in the Gram-negative bacteria, especially for E. coli, the outer membrane is exposed to cyclodextrins, and an interaction with the λ receptor as mentioned above cannot be excluded. An alternative presentation of the QCM-D data is shown in Figure 7 in which the dissipation is plotted versus the change in frequency. Presenting the data in this form eliminates time as an explicit parameter and makes it possible to directly compare the ratio between ΔD and Δf, that is, the induced energy dissipation per coupled unit mass.35 One is able to see directly H


Research Article


Figure 8. Quantitative evaluation by qRT-PCR of the bacterial capture and relative elution yields for β-CD-coated surfaces prepared with [β-CDCOOH] = 2, 5, and 10 g/L. (A, B) E. coli and (C, D) B. subtilis.

α-, β-, or γ-cyclodextrin with comparable affinities in the micromolar range as described for the product of the malE gene of E. coli, MBP.32 In this regard, in studies of the maltose system in other members of the family Enterobacteriaceae,37 it has been found that MBP is also produced by S. marcescens and that this bacterium transported maltose with high affinity. 3.3. Bacterial Cell Capture and Release Using SurfaceModified Si Pillar Arrays. As a proof of concept, we next studied bacterial cell capture and subsequent release using a micropillar-integrated microfluidic device (see Figure 1). To evaluate this, we selected as model micro-organisms E. coli and B. subtilis, which showed the best binding to the β-CD-coated QCM-D crystal substrates among the Gram-negative and Gram-positive bacteria tested, respectively. The micropillar arrays were modified by an EDC-mediated amidation similarly to the QCM-D crystal substrates using solutions of β-CDCOOH at three different concentrations (2, 5, and 10 g/L) in MES buffer (pH 4.75). Samples, with a volume comprised between 0.1 and 1 mL, of bacterial cells (1 × 103 cells/μL) in ultrapure water were injected through the pillar arrays with a flow rate of 16 μL/s (1 mL/min). This flow was selected as being high enough for a proof of concept of a rapid analysis. After it was rinsed first with 30 μL of TRIS buffer (pH 7) containing DIMEB ([DIMEB] = 540 g/L) and second with 30 μL of TRIS buffer (pH 7), the yields of bacterial capture and elution were measured by qRT-PCR. It appeared that elution can be performed in the small 30 μL volume when surfaces are prepared with 2 and 5 g/L of β-CD-COOH, whereas the elution was incomplete in this volume for 10 g/L of β-CDCOOH, as bacteria are significantly detected in a second rinsing step after DIMEB elution. This result suggests that interactions between bacterial cells and surface are stronger when the surface coverage with CD is increased, leading to a higher bacterial cell capture but also to a more difficult elution step, requiring higher volumes of eluant. Note that cells remain

viable after their capture and release (based on plate count experiments exhibiting a number of viable colonies from final samples after concentration in the microdevice in good agreement with the concentration of bacteria determined in the same samples by qRT-PCR, data not shown) and that no inhibition of the qRT-PCR analysis due to the presence of DIMEB was observed. In spite of imprecisions due to measurement errors (related to cell-to-cell variation, sample preparation, and other effects due to experimental design), the dependence of the capture yield and relative elution yield (recovered bacteria after elution/captured bacteria), estimated by qRT-PCR, on the concentration of β-CD-COOH used to modify the surface showed a similar trend for both types of bacterial cell (Figure 8). The bacterial capture yield is improved by increasing the concentration of β-CD-COOH, which can be related to a higher surface coverage by β-CD. This trend is much more pronounced in the case of E. coli. However, the relative elution yield remains nearly unchanged. The best results were obtained with E. coli, for which the relative elution yield was equal to or higher than 60% for all the concentrations of β-CD-COOH used to modify the micropillar array (Figure 8). This is in line with the QCM-D experiments showing efficient capture of E. coli by the β-CD-modified surface. Interestingly, we did not observe significant variations in the capture when the injected sample volume was increased from 0.1 to 1 mL for the surfaces prepared with 2 and 5 g/L compatible with elution in 30 μL (Figures S7 and S8). Gathering these results, it appears that best performances are a balance between the concentration factor (volume of bacteria suspension initially injected/volume of recovered bacteria suspension after elution) and the absolute yield (recovered bacteria after elution/bacteria initially injected in the microdevice). A 33-fold volume reduction can be achieved from an initial sample volume of 1 mL for surfaces prepared with [βI


Research Article

ACS Applied Materials & Interfaces CD-COOH] = 5 g/L, with an absolute yield of 44% for E. coli and 18% for B. subtilis. Performance of our microdevice may be optimized by modifying the design parameters of the microchip, like the pillars spacing, entries design, and also microfluidic parameters like flow rate, especially during elution as back and forth liquid flow at a high flow rate, may improve elution at high surface coverage by CD. Moreover, an intermediate surface coverage by using 7.5 g/L of β-CD-COOH could be interestingly tested, as it could be the best compromise regarding the balance between the concentration factor and the absolute yield.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Rachel Auzély-Velty: 0000-0003-2038-7604

4. CONCLUSION In conclusion, we report for the first time the potential of βCD-coated surfaces for bacterial capture via selective interactions between the CD cavity and bacterial cell surface components. The outstanding bacterial capture capability of such surfaces enables concentration of bacterial aqueous suspensions by elution in a volume of liquid lower than the sample volume initially injected containing DIMEB as a competitive host molecule. This capture/elution strategy is performed in physiological conditions and can be integrated in a microfluidic device, thereby allowing the identification and quantification of target species by qRT-PCR. Though the binding mechanism of bacterial cells could not be completely elucidated, QCM-D analysis of the binding of several Gramnegative and Gram-positive bacteria showed the potential to probe different bacterial species. Among the different types of bacteria tested, E. coli exhibited more propensity for adsorbing on the β-CD-modified surface, which may be due to a combination of inclusion complex formation with noncarbohydrate-binding proteins and molecular recognition with carbohydrate-binding proteins exposed on the bacterial cell surface. Finally, we presented a first proof-of-concept demonstrating the potential of β-CD-modified microchips for concentration of bacteria. The results obtained with E. coli suggest that this approach could be broadly applicable among Gram-negative bacteria, which share common cell membrane structures. This will be the focus of future studies together with the analysis of more complex real samples. Though our capture/elution strategy may offer less specifity/selectivity than biorecognition molecules such as antibody, aptamers, or lectins, it is attractive to address some problems related to cost and lifetime for bacterial capture.

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bacterial cells bound to the sensor crystal, effect of injected bacterial suspension volume on the E. coli capture and relative elution yields for β-CD-coated surfaces prepared with [β-CD-COOH] = 2 and 5 g/L (PDF)

Notes

The authors declare no competing financial interest. For QTM: D. Johannsmann, Technical University of Clausthal, Germany; http://www.tu-clausthal.de/en/research/johannsmann-group/qcm-modeling; option “small load approximation”.

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ACKNOWLEDGMENTS This work has been supported by the Arcane Labex program, funded by the French National Research Agency (ARCANE Project No. ANR-12-LABX-003). The authors thank the NMR platform of ICMG (FR2607) for its support; Dr. L. CocheGuérente for her assistance in QCM-D analyses performed in the Chimie NanoBio-ICMG platform (Grenoble Alpes Univ., Grenoble, France); C. Fontelaye for silicon chips functionalization and M. Alessio for pillar chips packaging; D. Mariolle for the AFM experiments.

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REFERENCES

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.7b02194. Observation of interaction between E. coli bacteria and fluorescently labeled β-CD by fluorescence microscopy, QCM-D profile during addition of β-CD ([β-CD] = 18 g/L) and TRIS buffer on bacterial cells E. coli bound to the sensor crystal, QCM-D profile during successive additions of DIMEB ([DIMEB] = 18 g/L) and TRIS buffer on bacterial cells E. coli bound to the sensor crystal, calorimetric titration of natural β-CD, DIMEB and RAMEB with ADAc, QCM-D profile during addition of RAMEB ([RAMEB] = 540 g/L) and TRIS buffer on bacterial cells E. coli bound to the sensor crystal, QCM-D profile during addition of DIMEB ([DIMEB] = 540 g/L) and TRIS buffer on Gram-positive and Gram-negative J


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Î²-CD-Functionalized Microdevice for Rapid Capture and Release of

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