# Microfluidics-Based Electrochemical Detection of Antimicrobial-Resistant DNA Sequence in Lysed Escherichia coli Medium

Hiu Mun Man, Choayb Omar, Martina Freisa, David Bouville, Téo Baptiste, Anne-Marie Haghiri-Gosnet, Hervé Jacquier, Isabelle Le Potier, Jean Gamby

► To cite this version:

Hiu Mun Man, Choayb Omar, Martina Freisa, David Bouville, Téo Baptiste, et al. Microfluidics-Based Electrochemical Detection of Antimicrobial-Resistant DNA Sequence in Lysed Escherichia coli Medium. ACS Electrochemistry, 2025, 10.1021/acselectrochem.4c00203. [https://doi.org/10.1021/acselectrochem.4c00203]

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Distributed under a Creative Commons Attribution 4.0 International License
Microfluidics-based electrochemical detection of antimicrobial-resistant DNA sequence

in lysed *Escherichia coli* medium

Hiu Mun Man<sup>a</sup>, Choayb Omar<sup>a</sup>, Martina Freisa<sup>a</sup>, David Bouville<sup>a</sup>, Téo Baptiste<sup>a</sup>, Anne-Marie Haghiri Gosnet<sup>a</sup>

Hervé Jacquier<sup>b,c</sup>, Isabelle Le Potier<sup>a</sup>, and Jean Gamby<sup>a*</sup>

<sup>a</sup>Centre de Nanosciences et de Nanotechnologies, Université Paris-Saclay, CNRS, 91120 Palaiseau, France

<sup>b</sup>Unité de Bactériologie, Département de Prévention, Diagnostic et Traitement des Infections, Hôpital Henri Mondor, APHP, 94000 Créteil, France

<sup>c</sup>EA 7380 DYNAMIC, Université Paris-Est Créteil, 94010 Créteil, France

*Corresponding author: jean.gamby@cnrs.fr
**Abstract**

Facing the emergence of multi-drug-resistant bacteria, rapid and sensitive detection of the antimicrobial resistant genes in such organisms is crucial for appropriate and timely treatment management. In this work, we have fabricated a microfluidic DNA biosensor immobilized with 6-mercaptohexanol and probe DNA sequence, aimed for the detection of antibiotic-resistant genes in *Escherichia coli* (*E. coli*). As proof-of-concept, we have demonstrated the successful electrochemical detection of the target DNA sequence down to 10 fM in the presence of diluted lysed *E. coli* cells, by cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS). We used a new approach to analyze the CV and EIS plots based on the influence of non-linear mass transport phenomenon more representative of the surface modification under flow conditions in microchannel. The custom fitting of the model allows us to estimate the apparent charge transfer resistance and thus evaluate the surface coverage and surface molecular immobilization on the electrode, which holds great potential for a wide range of bio-electrochemical sensing applications in complex sample matrices.

**Keywords:**

*Escherichia coli*, DNA biosensor, 6-mercaptohexanol, microfluidics, antimicrobial resistance
# INTRODUCTION

Mortality due to bacterial infections is a major public health issue, particularly in the context of increasing antibiotic resistance ¹. *Escherichia coli* (*E. coli*) is the most abundant aerobic bacterium present in the intestinal microbiota of vertebrates, but is also one of the main bacterial causes of bacterial infections. This bacterium is notably capable of acquiring numerous resistance genes to many classes of antibiotics, notably through horizontal transfer of mobile genetic elements. The emergence of antibiotic resistance in *E. coli* is particularly worrying, especially resistance to 3rd generation cephalosporins (3GCs), beta-lactam antibiotics frequently used in the probabilistic treatment of bacterial infections. Resistance to 3GCs is generally mediated by an enzyme called extended-spectrum beta-lactamase (ESBL).

The frequency of ESBL-producing *E. coli* has exploded over the last twenty years, due to the epidemic success of *E. coli* strains that have acquired the gene encoding a CTX-M-type ESBL².

Diagnosis of bacterial infections is generally based on culture techniques, which allow antibiotic susceptibility testing after several days' incubation. Thus, in case of infection by resistant bacteria, the introduction of effective antibiotic treatment is sometimes delayed by the time required for traditional microbiological diagnosis. This delay in optimal management is associated with increased mortality for the most severe infections³. Worldwide, antibiotic resistance in *E. coli* is described as one of the main causes of mortality attributable to bacterial infections¹. To speed up microbiological diagnosis, molecular-based methods have emerged in recent years. However, the susceptibility of these methods is often incompatible with very low levels of bacterial inoculum, as found in blood infections.
In this context, electrochemical DNA biosensing arises to be an attractive alternative to overcome the limitations of standard PCR-based methods due to the high sensitivity, portability, short analysis time and low cost⁴⁻⁷. Microfluidic technology has significantly advanced DNA biosensing by reducing sample sizes while enhancing sensitivity and enabling multiplexed measurements⁸. This enables rapid clinical diagnostics, making devices suitable for settings with limited resources.

Recent advancements in electrochemical DNA sensing have shifted from using redox-labeled DNA to methods that preserve the physicochemical properties and bio-affinity of nucleic acids. This improvement is achieved by (1) incorporating a redox reporter directly into the electrolyte solution⁹, (2) modifying the electrode surface¹⁰, and (3) employing intercalation or electrostatic interactions for binding nucleic acids¹¹,¹². Self-assembled monolayers (SAMs) of thiol molecules facilitate the uniform arrangement of nucleic acids on the electrode surface¹³. Thiol linkers can be used to reduce non-specific adsorption of non-complementary nucleic acids⁷.

The decreased electron transfer rates after the functionalization of SAM on the electrode are often evidenced in the literature by the changes in peak currents in cyclic voltammetry (CV) or charge transfer resistance in electrochemical impedance spectroscopy (EIS)¹⁴⁻¹⁷. However, these electrochemical signals are averaged over the entire electrode surface, making it difficult to distinguish whether signal changes result from reduced electroactive surface area or diminished electron transfer kinetics. This is particularly important to consider when biosensing is incorporated into microfluidic devices, where mass transport and electron transfer are the two major processes in confined spaces. Our group recently demonstrated a non-linear phenomenon related to mass transport using a model where the electrochemical
signal is governed by the redox reporter's transport through a partially blocked thiol layer on
the microchannel electrode ¹⁸. In this model, the direct adsorption of thiol-linked
oligonucleotide probes on the electrode surface creates small holes, affecting the mass
transport dynamics. We have shown with the detection of microRNAs (*miR-30b* and *miR-122*)
at platinum and gold surfaces that the observed current decrease during DNA immobilization
protocols does not correspond to a decrease of the charge transfer rate *k*₀, but is ascribed to
the small changes in mass transport within the microchannel-electrode. More recently, we
have optimized the ratio of tris(2-carboxyethyl)phosphine (TCEP) and 6-mercaptohexanol in
the determination of DNA in a microfluidic device ¹⁹.

The combination of microfluidics and electrochemistry for the detection of antibiotic
resistance/susceptibility has been exploited in the literature in recent years using a wide range
of electrode materials (gold<sup>20,21</sup>, pencil graphite<sup>22</sup>, cellulose based substrates<sup>23</sup> and
microfabricated substrates<sup>19</sup>). Song and co-workers have proposed a polyadenine-DNA
tetrahedron probe-based electrochemical sensor for the detection of multiple antibacterial
genes including *MCR-1*, *bla*<sub>NDM</sub>, *bla*<sub>KPC</sub> and *bla*<sub>OXA</sub> <sup>21</sup>. Very recently, Gopalakrishnan et al.
have developed an electrochemical microfluidic device (ε-μD) combining carbon screen-
printed electrodes and microfluidic channels for the sensitive detection of bacteria (*E. coli*
and *Bacillus subtilis*) <sup>24</sup>. The efficacy of various antibiotic drugs including ampicillin and
tetracycline were evaluated in spiked urine samples.

This present work is centered on the detection and analysis of the *CTX-M-8* gene which
belongs to the β-lactamase genes *bla*<sub>CTX-M</sub> subgroup2 in antibiotic resistance strains of *E. coli*.
The detection of this 23-nucleotide long DNA sequence was successfully achieved in the
presence of a complex medium (*E. coli* cell lysates) over a wide range of concentrations. We
utilized the EIS measurement model proposed by Freisa *et al.*<sup>18</sup> to analyze the responses
obtained from the DNA platinum biosensor modified with 6-mercaptohexanol. The
integration of EIS and DNA sensing microfluidic platforms enhances the capability for high-
throughput screening and analysis, making it feasible for use in point-of-care devices. This
combination could lead to the development of portable early diagnostic tools that provide
real-time multiplexed measurements in the near future.
# EXPERIMENTAL

## DNA sequences and chemicals

The selected antibiotic-resistance DNA sequences ($bla_{CTX-M-8}$ denoted in this work GP8 probe and target, as seen in **Table 1**), were purchased from Eurogentec, Belgium. The reverse-phase HPLC-purified DNA sequences were used without further purification upon receipt and diluted in 0.5 M NaCl in the following experiments. 6-mercaptohexanol (MCH), sodium chloride, potassium hydroxide, potassium hexacyanoferrate (III) and potassium hexacyanoferrate (II) trihydrate were obtained from Sigma Aldrich, France. Tris(2-carboxyethyl) phosphine (TCEP), LIVE/DEAD BacLight Bacterial Viability Kits and Bacterial Protein Extraction Reagent (B-PER) were purchased from Thermo Fisher Scientific, France.

<table><caption>**Table 1** Selected nucleotide sequences used in this work.</caption><thead><tr><td></td><td>Sequence</td><td>Modification</td></tr></thead><tbody><tr><td>GP8 probe (5' → 3')</td><td>GCG-GCG-CTG-GAG-AAA-AGC-AGC-GG</td><td>5' Thiol modifier -SH-C6</td></tr><tr><td>GP8 target (3' → 5')</td><td>CGC-CGC-GAC-CTC-TTT-TCG-TCG-CC</td><td>-</td></tr></tbody></table>

## Fabrication of microfluidic biosensor

The two-electrode system microfluidic device was fabricated following the work developed in our group 25-29. A graphical representation of the device is demonstrated in **Figure 1a**. Briefly, the microfabrication processes include SU-8 photolithography of electrode patterns (a working electrode of 30 µm x 300 µm and a counter electrode of 2 mm x 300 µm, respectively, as shown in **Figure 1b**) onto a glass wafer, by spin coating a primer hexamethyldisilazane (HDMS) followed by a photoresist AZ5214 for 30 s at a speed of 4000 rpm and an acceleration of 2000 rpm s⁻¹ to achieve a thickness of 1.4 µm. The coated wafer was baked at 110 °C for 1 min 30 s and the exposure of UV light through a hard photomask. The glass wafer was heated again post exposure and developed in the developer AZ 2026
MIF. Subsequently, a 100 nm layer of platinum was deposited on top of a 10 nm-thick
adhesion layer of titanium by evaporation. The fluidic channels were fabricated by soft
lithography of PDMS on a patterned silicon wafer. The fluidic inlets and outlets were
achieved using stainless steel punches (Rapid-core, punch diameter = 0.5 mm). The PDMS
and glass substrate were assembled by O₂ plasma-activated bonding for 30 s, followed by a
final curing step at 60 °C for a minimum of 4 hours.

<figure><img src="image_2.png" /><figcaption>**Figure 1** a) A graphical representation of the fabricated microfluidic device. b) A micrograph depicting the geometric of the working and counter electrodes. c) The experimental protocol used in this work: (1) Pretreatment of platinum electrodes by KOH cycling, (2) functionalization of thiol and DNA probes, (3) introduction and measurement of lysed E. coli suspension as a background signal in the microfluidic channel and (4) introduction and measurement target DNA-spiked bacterial lysates.</figcaption></figure>

Electrochemical measurements

The fabricated platinum microelectrodes were pre-treated by repeated cycling in 0.5 M potassium hydroxide (-0.2 – -1.2 V, scan rate: 50 mV s⁻¹, 10 cycles) to ensure a clean electrochemical surface.

The DNA probe GP8 was diluted in 5 mM TCEP to prevent the reformation of disulfide
bridges of DNA. MCH was added to the DNA probe/TCEP solution, achieving the final
concentration of DNA probe of 10⁻⁶ M. The concentrations of TCEP and MCH was kept at 5
mM TCEP and 50 mM MCH in this work, which was the optimized ratio previously determined to ensure an effective immobilization of DNA probes¹⁹. The probe/TCEP/MCH solution was injected into the microchannel and immobilized under static conditions for two hours at room temperature, and then the microchannel was washed with 0.5 M NaCl.

The sensing protocol is illustrated in **Figure 1c**. For the electrochemical measurements in the presence of *E. coli* cell lysates, the lysed solution (either undiluted or at a 50% v/v dilution) was injected into the microchannel after the immobilization of the DNA probe/MCH/TCEP and incubated for 30 minutes, followed by rinsing the channel with 0.5 M NaCl. This procedure provided a 'blank' signal for the lysed bacteria. Subsequently, the lysed bacteria suspension was spiked with increasing concentrations of synthetic GP8 target DNA (10⁻¹⁴, 10⁻¹², 10⁻¹⁰ and 10⁻⁷ M in 50% diluted lysates, and 10⁻¹⁴, 10⁻¹² and 10⁻¹⁰ M in the presence of undiluted lysates) with a 30-minute incubation in the microchannel at each concentration.

For the measurements in the absence of *E. coli* cell lysates, the immobilization of probe/TCEP/MCH was followed the hybridization process of the complementary DNA sequence of GP8 (target DNA) at increasing concentrations, which involved a 30-minute incubation in static conditions, followed by rinsing with 0.5 M NaCl.

Electrochemical characterizations of the DNA biosensor were performed after each step described above, in the traditionally used redox couple, 20 mM equimolar [Fe(CN)₆]³⁻/⁴⁻ containing 0.5 M of NaCl, under flow conditions at 0.5 µL s⁻¹ by cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS). CV was measured in the potential window of 0.2 V to –0.2 V with a scan rate of 20 mV s⁻¹. EIS analyses were recorded at VDC = 0 V and VAC = 10 mV in the frequency range of 1 MHz to 0.1 Hz. A potentiostat (BioLogic SP-300, operated by the proprietary software EC-lab) was used for all measurements.
The flow control (a constant flow of 0.5 µL s⁻¹) was achieved by the neMESYS syringe pump system (cetoni GmbH). After the characterization of bare electrode, the DNA probe-immobilized electrode was measured three times by CV and EIS methods consecutively to ensure the stability of the electrodes. The electrical equivalent circuit used in the measurement protocol for electrochemical DNA biosensor to detect antimicrobial DNA sequences in spiked lysed bacterial media is shown in **Scheme 1**. The description of each element is given in the next section (see **Table 2**).

## EIS model fitting

The software created by Watson and Orazem (Florida Research Foundation) is a Python-based toolbox to analyze impedance spectroscopy data, which estimates the capacitance, ohmic resistance and the apparent charge transfer resistance³⁰. The modified Randles equivalent circuit shown in **Scheme 1** was used to model the experimental data¹⁸.

<figure><img src="image_3.png" /><figcaption>**Scheme 1** The electrical equivalent circuit used for fitting the responses of electrochemical DNA biosensor for the detection of antimicrobial DNA sequences in spiked lysed bacterial media.</figcaption></figure>

As we reported previously ¹⁸, the modified circuit includes a global solution resistance $R_e$, a local solution resistance at the surface of the SAM-modified electrode $R_e'$, the charge transfer resistance $R_{ct}$ and the convection diffusion resistance $R_D$ (when $Z_{D f→0}$). Two capacitors were considered in this model, including the double layer capacitance $C_{dl}$ (here the constant phase element CPE, attributed to the circuit elements $Q_{dl}$ and $\alpha$) and the blocking layer capacitance for the thiolated-DNA layer. The double layer capacitance is deduced as follows in equation (1):
$$C_{dl,eq} = \left[ Q_{dl} \left( \frac{1}{R_e + R'_e} + \frac{1}{R_{ct}} \right)^{\alpha-1} \right]^{\frac{1}{\alpha}} \quad (1)$$

The heterogeneous charge transfer rate constant $k_0$ is calculated with the apparent $R_{ct}$ value using the following equation (2):

$$k_0 = \frac{RT}{F^2 S_{WE} R_{ct, \eta \to 0} C^0} \quad (2)$$

where T is the temperature in Kelvin (293.16 K), R refers to the universal gas constant (8.3145 J K⁻¹ mol⁻¹), S<sub>WE</sub> is the electrode surface area (9 × 10⁻⁵ cm²), R<sub>ct, η→0</sub> is the apparent charge transfer resistance when overpotential equal to 0 V, C⁰ is the concentration of the redox indicated in moles per m³.

<table><caption>**Table 2** List of the symbols, units and initial guesses used in the equivalent circuit for modelling.</caption><thead><tr><th>Circuit element</th><th>Unit</th><th>Initial guesses</th></tr></thead><tbody><tr><td>Global electrolyte resistance, R<sub>e</sub></td><td>Ω</td><td>1.8 × 10<sup>3</sup> – 4 × 10<sup>3</sup></td></tr><tr><td>Local electrolyte resistance, R<sub>e</sub>'</td><td>Ω</td><td>1 – 10<sup>5</sup></td></tr><tr><td>Characteristic time for diffusion τ<sub>D</sub></td><td>s</td><td>10<sup>-3</sup>– 10</td></tr><tr><td>Diffusion resistance, R<sub>D</sub></td><td>Ω</td><td>1 – 4 × 10<sup>5</sup></td></tr><tr><td>CPE element</td><td>Q<sub>dl</sub> Ω<sup>-1</sup> s<sup>α</sup></td><td>10<sup>-12</sup> – 10<sup>-4</sup></td></tr><tr><td>CPE exponent, α</td><td>-</td><td>0.6 – 1</td></tr><tr><td>Charge transfer resistance, R<sub>ct</sub></td><td>Ω</td><td>10<sup>3</sup> – 10<sup>9</sup></td></tr><tr><td>Blocking capacitance, C<sub>b</sub></td><td>F</td><td>10<sup>-12</sup>– 10<sup>-4</sup></td></tr></tbody></table>

## Complex sample medium preparation – *Escherichia coli* lysis

A genetically modified, non-pathogenic strain of *E. coli* K-12 was lysed in 300 µL of Bacterial Protein Extraction Reagent, which contains Tris-HCl buffer at pH 7.5 and a proprietary non-ionic detergent. The lysis process was carried out for 15 minutes at room temperature.
To verify the efficiency of bacterial lysis, a LIVE/DEAD Bacterial Viability Stain Kit was used. This kit employs SYTO 9 to stain living bacteria and propidium iodide to stain dead or dying cells, with fluorescence microscopy used to assess the results. The resulting lysed suspension was then filtered through a 0.2 µm filter before proceeding with electrochemical measurements.
# RESULTS

## Cyclic voltammetry

The developed DNA biosensor was characterized by CV in a solution of 20 mM equimolar $[Fe(CN)_6]^{3-}$ containing 0.5 M NaCl with a scan rate of 20 mV s$^{-1}$ and a flow rate of 0.5 µL s$^{-1}$. Figure 2a presents the CV responses of the platinum electrode at various stages of the measurement process. The KOH-pretreated bare Pt electrode exhibited an expected microelectrode response with a sigmoidal curve, reaching a steady-state current at ~4000 nA (from theoretical expression of Levich equation $^{31-33}$) as reported previously for channel-electrode $^{18,25,25}$, which indicates fast and efficient electron transfer, as expected following activation by repeated cycling. The immobilization of the DNA probe/MCH/TCEP significantly reduced the CV signal from -4000 nA to -1600 nA, indicating successful and maximal immobilization of the GP8 probe/MCH, which suggests a non-linear mass transfer on a surface being microstructured with SAM$^{18}$. After incubating with a 50% v/v diluted lysed bacterial suspension, used as a 'blank' measurement, a slight decrease in the CV response was observed. This could indicate that some components of the lysed cells adsorbed onto the electrode surface (non-specific adsorption). The lysed solution was then spiked with increasing concentrations of the GP8 target sequence (complementary to the DNA probe), ranging from $10^{-14}$ to $10^{-7}$ M (denoted throughout this work as Hybridizations 1-4, respectively). A gradual decrease in absolute current was observed, from 1150 nA for the probe down to 750 nA for the highest GP8 target concentration ($10^{-7}$ M).

The normalized current was calculated with respect to the probe current using the following equation (3):

$$i_{\text{normalized}} = |(i_{\text{probe}} - i_{\text{target}})/i_{\text{probe}}| \quad (3)$$

A linear relationship between the normalized current and the logarithm of the concentration was observed in the range of $10^{-14}$ to $10^{-10}$ M, with an R$^2$ value of 0.993, as shown in Figure
**2b.** The red data point in the calibration curve in **Figure 2c** represents the “blank” sample, i.e., the lysed bacteria at a 50% v/v dilution. It is important to note that the current signal obtained for the 50% lysed bacteria sample was found to be equal to a 10<sup>-16</sup> M concentration (graphically extrapolated as in **Figure 2c** that confirms the linear trend, as the R<sup>2</sup> value and the slope remain unchanged with or without the blank at this concentration.

To ensure that the linear response was not a result of the gradual accumulation of lysed *E. coli* on the electrode surface, we conducted the same calibration experiments using undiluted *E. coli* cell lysates, as shown in **Figure 2b**. A sharp decrease in absolute currents was observed after the functionalization with the GP8 probe/MCH, followed by a slight decrease in currents during the incubation with undiluted bacterial cell lysates. Interestingly, the hybridization processes of GP8 targets in the presence of undiluted bacterial cell lysates were observed to be inconsistent and lacked any discernible trend (**Figure 2c**, grey markers). The lack of linearity in undiluted bacterial lysates could be attributed to the lysed fragments (from the peptidoglycan cell wall and the outer membrane containing lipids, proteins, and polysaccharides<sup>34</sup>) rapidly saturating the microchannel. In the case of non-specific adsorption, these lysed fragments act as blocking agents on the free remaining active sites. This explains the current decrease following the incubation of undiluted lysed *E. coli*. The active surface is quickly saturated by the lysed fragments and DNA targets cannot access these free remaining sites for hybridization. Thus, the change in current is independent of the target molar concentration. Conversely, this further confirms that the observed linear response in the case of 50% diluted cell lysate indicates successful DNA hybridization.
<figure><img src="image_4.png" /><figcaption>**Figure 2 a)** Cyclic voltammograms recorded in 20 mM ferri-/ferro-cyanide in 0.5 M NaCl at bare platinum electrode (black curve), after immobilization of probe DNA with TCEP and MCH (red curve), 50 % E. coli cell lysates (blue curve), and increasing concentrations of target DNA from 10 x 10<sup>-14</sup> M to 10 x 10<sup>-7</sup> M, inset shows a zoom to the curves at -0.2 V.

**b)** Cyclic voltammograms recorded at bare platinum electrode (black curve), after immobilization of probe DNA with TCEP and MCH (red curve), undiluted E. coli cell lysates (blue curve), and increasing concentrations of target DNA from 10 x 10<sup>-14</sup> M to 10 x 10<sup>-10</sup> M, inset shows a zoom to the curves at -0.2 V.

**c)** Calibration curves of the normalized currents from DNA hybridization of the target sequence complementary to the GP8 probe sequence in the presence of 50 % bacterial lysates (in black markers) and undiluted bacterial lysates (in grey markers). The red point represents the “blank” recorded in 50% cell lysates without any target DNA. The unfilled red marker represents the “blank” signal recorded in undiluted cell lysates.</figcaption></figure>
Electrochemical impedance spectroscopy

The electrochemical behavior of the fabricated DNA biosensor was investigated using electrochemical impedance spectroscopy (EIS). **Figure 3** presents the experimental EIS responses alongside the fitted data for the KOH-pretreated bare Pt electrode and the probe/MCH-immobilized Pt surface, represented in Nyquist, phase angle, and Bode plots. The custom fitting performed using the measurement model program demonstrated an excellent fit with the experimental results.

The MCH-functionalized DNA electrode was subsequently incubated with the 'blank' (undiluted and 50% v/v diluted bacterial lysates) and varying concentrations (10⁻¹⁴, 10⁻¹², 10⁻¹⁰ and/or 10⁻⁷ M) of GP8 target DNA in the presence of bacterial lysates. Nyquist plots for each step of the measurements, along with the corresponding calibration curves of normalized Rct against the logarithm of GP8 concentration are shown in **Figure 4**. The normalized Rct values were determined with respect to the Rct value of the probe as shown in Equation (4):

$$R_{ct\_normalized} = |(R_{ct\_target} - R_{ct\_probe})/R_{ct\_probe}| \quad (4)$$

The fitted parameters, along with the calculated Cdl and k₀ values for measurements conducted in 50% diluted and undiluted bacterial lysates, are provided in **Table 3**. The calculated k₀ values for free remaining active sites (1-θ) were expressed in relation to the surface coverage ratio (θEIS) which were determined as follows through equations (5) and (6):

$$\theta_{EIS} = 1 - \left( \frac{R_{ct\_bare}}{R_{ct\_step}} \right) \quad (5)$$

$$k_0 = \frac{RT}{F^2 S(1-\theta)R_{ct-step}C^0} \quad (6)$$

Additional fitted parameters and calculated values for all electrodes are available in **Tables S1-S3** in the Supplementary Information (SI).
<figure><img src="image_5.png" /><figcaption>**Figure 3** Electrochemical impedance spectroscopy and equivalent circuit fitting of bare Pt electrode (in black) and probe/MCH-modified Pt electrode (in green) in electrolytes containing 20 mM Fe(CN)₆²⁻ and 20 mM Fe(CN)₆⁴⁻. The EIS responses of experimental data (in circles and squares) and the fitted data (lines), represented in **a)** Nyquist plot, **b)** phase angle plot **c)** Bode plot. The modified Randles circuit used for fitting is included in **a**), reproduced from **Scheme 1**.</figcaption></figure>

We begin by evaluating the fitting of the bare Pt electrode, as shown in **Figure 3**. We expect the model to closely resemble the well-known Randles circuit (shown in the inset of **Figure 3a** in black) for the bare Pt electrode due to the absence of a SAM blocking layer. This expectation is supported by the significantly lower $R'_e$ value at the bare Pt electrode (1 Ω) compared to that at the probe-functionalized electrode (7082 Ω). Additionally, there is a drastic increase in the charge transfer resistance ($R_{ct}$), which increases from 5239 Ω at the bare electrode to $2.07 \times 10^5$ Ω at the probe-immobilized electrode, indicating a successful immobilization of DNA probe on the Pt surface.

For measurements performed in 50% diluted lysates, the global ohmic resistance ($R_e$) of the bare electrode was (2542.5 ± 91.4) Ω. After functionalization, the $R_e$ values for all subsequent steps shifted to approximately (2600 ± 51.1) Ω, as shown in **Table 3**. This shift could be explained by the error values of $R_e$ determined from the fit model.
In **Figure 4a**, the charge transfer resistance ($R_{ct}$) has increased from $2.07 \times 10^5$ Ω cm² at the probe to $3.76 \times 10^5$ Ω at the most concentrated target DNA ($10^{-7}$ M). The normalized $R_{ct}$ is observed to be linearly related to the logarithm of target DNA concentration as illustrated in **Figure 4c**, with an $R^2$ value of 0.990. The surface coverage value $\theta$ (see **Table 3**) increases from 0.975 at the probe to 0.986 at the most concentrated target DNA hybridization, suggesting that the holes (free remaining active sites on surface) are getting gradually filled after each addition of target DNA.

This is verified by repeating the measurements in undiluted bacterial lysate suspension. As shown in **Figures 4b and c**, the apparent normalized charge transfer resistances do not have a linear relationship with the concentration of target DNA, similar to the results obtained in CV. The surface coverage ratios of measurements in undiluted cell lysates reveal that there is not a huge variation in $\theta$ from the probe to the final hybridization step (an increase from 0.980 to 0.987 in **Table 3**), confirming what we postulated with the CV results that the microchannel electrode has been quickly saturated in the presence of undiluted bacterial lysates.

The last column of **Table 3 and Tables S1-S3** revealed the calculated charge transfer rates $k_0$ (see equations (5)-(6)) for each step of the measurement were determined to be the same when their respective surface coverage ratios were taken into account. This confirms our non-linear mass transport model as mentioned in the introduction, where the observed decrease in the measured current intensity is not related to a decrease of charge transfer rate $k_0$ but to altered mass transport.
<figure><img src="image_6.png" /><figcaption>**Figure 4** Nyquist plots measured at the platinum electrode at each step of the experiment in the presence of **a)** 50% diluted lysed E. coli and **b)** undiluted E. coli lysates. The experimental data are shown in markers, and the fitted data are shown in lines. **c)** Calibration curves of normalized charge transfer resistances ($R_{ct}$) from DNA hybridization of the target sequence complementary to the GP8 probe sequence in the presence cell lysis solution, 50 % v/v (in diamonds) and 100 % v/v (in circles). The first red data points represent the “blank” recorded in 50% and 100% cell lysis solution without any target DNA, respectively.</figcaption></figure>
<table><caption>Table 3. Equivalent circuit element fitting parameters and the calculated C<sub>dl</sub>, θ and k<sub>0</sub> for the DNA biosensor in 50% and undiluted E. coli lysates.</caption><thead><tr><th rowspan="2">Experimental</th><th colspan="8">Fitted parameters</th><th colspan="4">Calculated values</th></tr><tr><th>R<sub>e</sub> (Ω)</th><th>R<sub>e</sub>' (Ω)</th><th>τ<sub>D</sub> (s)</th><th>R<sub>D</sub> (Ω)</th><th>Q<sub>dl</sub> (F s<sup>-a</sup>)</th><th>α</th><th>R<sub>ct</sub> (Ω)</th><th>C<sub>b</sub> (F)</th><th>χ<sup>2</sup>/ν *</th><th>C<sub>dl</sub> (F cm<sup>-2</sup>)</th><th>θ</th><th>k<sub>0</sub> (cm s<sup>-1</sup>)</th></tr></thead><tbody><tr><td>Bare Pt</td><td>2542</td><td>1</td><td>0.1</td><td>12581</td><td>1.8 × 10<sup>-7</sup></td><td>0.6</td><td>5332</td><td>9.2 × 10<sup>-9</sup></td><td>7.9 × 10<sup>-5</sup></td><td>1.91 × 10<sup>-5</sup></td><td>0.000</td><td></td></tr><tr><td>Probe</td><td>2621</td><td>7082</td><td>0.2</td><td>16158</td><td>1.8 × 10<sup>-8</sup></td><td>0.8</td><td>2.07 × 10<sup>5</sup></td><td>4.7 × 10<sup>-9</sup></td><td>5.6 × 10<sup>-5</sup></td><td>5.66 × 10<sup>-5</sup></td><td>0.975</td><td></td></tr><tr><td>50% lysed E. coli lysates</td><td>2646</td><td>7508</td><td>2.2</td><td>12537</td><td>1.7 × 10<sup>-8</sup></td><td>0.8</td><td>2.58 × 10<sup>5</sup></td><td>4.5 × 10<sup>-9</sup></td><td>3.3 × 10<sup>-5</sup></td><td>5.22 × 10<sup>-5</sup></td><td>0.980</td><td></td></tr><tr><td>Hybridization 1</td><td>2660</td><td>8302</td><td>2.6</td><td>13507</td><td>1.5 × 10<sup>-8</sup></td><td>0.8</td><td>2.87 × 10<sup>5</sup></td><td>4.5 × 10<sup>-9</sup></td><td>2.4 × 10<sup>-5</sup></td><td>4.91 × 10<sup>-5</sup></td><td>0.982</td><td>0.027</td></tr><tr><td>Hybridization 2</td><td>2696</td><td>9682</td><td>4.1</td><td>17762</td><td>1.4 × 10<sup>-8</sup></td><td>0.8</td><td>3.23 × 10<sup>5</sup></td><td>4.5 × 10<sup>-9</sup></td><td>2.2 × 10<sup>-5</sup></td><td>4.83 × 10<sup>-5</sup></td><td>0.984</td><td></td></tr><tr><td>Hybridization 3</td><td>2694</td><td>10000</td><td>4.5</td><td>19311</td><td>1.3 × 10<sup>-8</sup></td><td>0.8</td><td>3.50 × 10<sup>5</sup></td><td>4.5 × 10<sup>-9</sup></td><td>1.8 × 10<sup>-5</sup></td><td>4.67 × 10<sup>-5</sup></td><td>0.985</td><td></td></tr><tr><td>Hybridization 4</td><td>2673</td><td>10000</td><td>5.4</td><td>21980</td><td>1.3 × 10<sup>-8</sup></td><td>0.8</td><td>3.76 × 10<sup>5</sup></td><td>4.4 × 10<sup>-9</sup></td><td>1.7 × 10<sup>-5</sup></td><td>4.73 × 10<sup>-5</sup></td><td>0.986</td><td></td></tr><tr><td>Bare Pt</td><td>1800</td><td>335</td><td>0.1</td><td>11102</td><td>5.20 × 10<sup>-8</sup></td><td>0.9</td><td>3520</td><td>9.9 × 10<sup>-10</sup></td><td>2.22 × 10<sup>-3</sup></td><td>1.65 × 10<sup>-5</sup></td><td>0.000</td><td></td></tr><tr><td>Probe</td><td>2533</td><td>4942</td><td>1.0</td><td>12356</td><td>2.46 × 10<sup>-8</sup></td><td>0.8</td><td>1.73 × 10<sup>5</sup></td><td>5.0 × 10<sup>-9</sup></td><td>5.24 × 10<sup>-5</sup></td><td>6.86 × 10<sup>-5</sup></td><td>0.980</td><td></td></tr><tr><td>100% lysed E. coli lysates</td><td>2610</td><td>7138</td><td>0.5</td><td>16181</td><td>1.98 × 10<sup>-8</sup></td><td>0.8</td><td>2.15 × 10<sup>5</sup></td><td>5.1 × 10<sup>-9</sup></td><td>2.92 × 10<sup>-5</sup></td><td>6.08 × 10<sup>-5</sup></td><td>0.984</td><td>0.041</td></tr><tr><td>Hybridization 1</td><td>2615</td><td>7616</td><td>1.8</td><td>11519</td><td>1.74 × 10<sup>-8</sup></td><td>0.8</td><td>2.72 × 10<sup>5</sup></td><td>5.0 × 10<sup>-9</sup></td><td>1.50 × 10<sup>-5</sup></td><td>5.88 × 10<sup>-5</sup></td><td>0.987</td><td></td></tr><tr><td>Hybridization 2</td><td>2610</td><td>7865</td><td>3.0</td><td>9661</td><td>1.80 × 10<sup>-8</sup></td><td>0.8</td><td>2.54 × 10<sup>5</sup></td><td>4.9 × 10<sup>-9</sup></td><td>1.66 × 10<sup>-5</sup></td><td>5.93 × 10<sup>-5</sup></td><td>0.986</td><td></td></tr><tr><td>Hybridization 3</td><td>2606</td><td>8288</td><td>2.8</td><td>10149</td><td>1.74 × 10<sup>-8</sup></td><td>0.8</td><td>2.63 × 10<sup>5</sup></td><td>4.9 × 10<sup>-9</sup></td><td>1.46 × 10<sup>-5</sup></td><td>5.82 × 10<sup>-5</sup></td><td>0.987</td><td></td></tr></tbody></table>

<footnote>*The χ<sup>2</sup>/ν statistic value is generated from the regression weighted by the stochastic error structure from the experimental data in the measurement program<sup>30</sup>.</footnote>
The surface concentration ($Γ_ρ$) was also evaluated using the surface coverage ($θ$) determined from EIS. Firstly, the surface occupied by DNA probes ($A_{DNA probe}$) is the surface coverage ($θ$) multiplied by the WE electrode surface area ($A_{WE}$). Assuming the DNA strands are oriented in an upright position³⁵ and that the diameter of the DNA head is around 1 nm, the number of DNA probes immobilized on the electrode surface ($N_{SS-DNA}$) is deduced to be around $2.79 × 10^9$ molecules²⁶. The binding site density ($b_m$) was then estimated to be $3.1 × 10^{13}$ molecules cm⁻² using Equation (7) as follows:

$$b_m = \frac{N_{SS-DNA}}{A_{WE}} \qquad (7),$$

where $A_{WE}$ refers to the total electrode surface area ($9 × 10^{-5}$ cm²). We obtained the surface concentration which is given by Equation (8):

$$Γ_ρ = \frac{b_m}{N_A} \qquad (8),$$

where $N_A$ represents the Avogadro's number.

The estimated surface concentrations of the DNA sensor after hybridization in undiluted and 50% diluted *E. coli* lysates are summarized in **Table 4**.

<table><caption>**Table 4.** Theoretical estimates of surface concentration ($Γ_ρ$) for various target DNA concentrations in the presence of undiluted and 50% diluted lysed *E. coli*.</caption><thead><tr><th colspan="2"></th><th colspan="2">Γ<sub>ρ</sub> (mol cm<sup>-2</sup>)</th></tr><tr><th colspan="2"></th><th>Undiluted <i>E. coli</i><br>lysates (n=2)</th><th>50% diluted <i>E. coli</i><br>lysates (n=3)</th></tr></thead><tbody><tr><td rowspan="5">Target DNA<br>concentration (M)</td><td>Blank (Lysed <i>E. coli</i>)</td><td>52.07 × 10<sup>-12</sup> ± 8 × 10<sup>-14</sup></td><td>51.99 × 10<sup>-12</sup> ± 1 × 10<sup>-13</sup></td></tr><tr><td>10<sup>-14</sup></td><td>52.20 × 10<sup>-12</sup> ± 3 × 10<sup>-14</sup></td><td>52.09 × 10<sup>-12</sup> ± 1 × 10<sup>-13</sup></td></tr><tr><td>10<sup>-12</sup></td><td>52.15 × 10<sup>-12</sup> ± 3 × 10<sup>-14</sup></td><td>52.17 × 10<sup>-12</sup> ± 1 × 10<sup>-13</sup></td></tr><tr><td>10<sup>-10</sup></td><td>52.14 × 10<sup>-12</sup> ± 7 × 10<sup>-15</sup></td><td>52.22 × 10<sup>-12</sup> ± 9 × 10<sup>-14</sup></td></tr><tr><td>10<sup>-7</sup></td><td>-</td><td>52.26 × 10<sup>-12</sup> ± 8 × 10<sup>-14</sup></td></tr></tbody></table>
As observed in **Figure 5**, the increase in surface concentration in undiluted *E. coli* lysates is not dependent on the molar concentration of target DNA. It confirms that the change in surface coverage ratios is linked to the blocking process caused by the lysed fragments, as previously mentioned. Conversely, in the case of 50% diluted lysates, the evolution of the surface concentration is dependent on the molar concentration of target DNA.

<figure><img src="image_7.png" /><figcaption>**Figure 5** Estimated surface concentration (Γp) of the DNA biosensor after hybridization plotted against the logarithm of molar concentration of target DNA.</figcaption></figure>
In addition, we performed the same set of experiments without bacterial lysates using CV and EIS. **Figure 6** shows the calibration curves of **a)** normalized currents and **b)** normalized charge transfer resistance against the logarithm of GP8 concentration, both with (solid black curves) and without (gray dotted curves) *E. coli* lysates. The slopes for CV are nearly identical, with values of 0.022 and 0.020, respectively. For EIS, the slopes are similarly close, with values of 0.078 and 0.065. Interestingly, in both methods, the signals in the presence of cell lysates are consistently higher than those without, underlining that the increased signal may be due to elevated background noise from the complex sample medium.

<figure><img src="image_8.png" /><figcaption>**Figure 6** Calibration curves of **a)** normalized currents and **b)** normalized charge transfer resistance ($R_{ct}$) from DNA hybridization of the target sequence complementary to the GP8 probe sequence, in the presence of 50 % *E. coli* cell lysates (solid black curves) and in the absence of bacterial cell lysates (dotted grey curves).</figcaption></figure>

It is also interesting to note that the EIS method is more sensitive, as evidenced by the consistently bigger slopes of EIS responses in Figure 6 (0.078 and 0.065, with and without cell lysates, respectively), compared to the CV slopes (~0.02). This could be explained by the non-linear mass transport effects under flow conditions during voltammetric measurements. As EIS measurements are less influenced by the diffusion processes, we would anticipate a
higher sensitivity and reliability with regard to the reactions that take place close the electrode surface. The combination of CV and EIS thus provides a more comprehensive view of the electrode.

**Table 5** presents a comparison of the developed sensor with existing methods for antibiotic resistance detection in the literature. The developed sensor has the advantages of fast analysis time for a direct detection of target nucleic acid sequence. The sensor in the absence of bacterial lysate was able detect as low as 10 aM¹⁹. Moreover, the selectivity of our sensor was previously validated against a non-complementary sequence *bla*<sub>CTX-M-2</sub> across the concentrations 10<sup>-18</sup> M – 10<sup>-8</sup> M, and the voltammetric response was shown to be negligible even at 0.1 nM. This further validates the sensitivity, robustness and selectivity of our fabricated sensor, which will be incorporated in our ongoing researching in streamlining the bacterial lysis workflow into the microfluidic system for the detection of drug-resistant strains of bacteria.
<table><caption>**Table 5.** Comparison of the current work with existing methods for the detection of antibiotic resistance.</caption><thead><tr><th>Method</th><th>Target</th><th>Analysis time</th><th>Detection range</th><th>LOD</th><th>Selectivity</th><th>Sample matrix</th><th>Ref.</th></tr></thead><tbody><tr><td>Colorimetric detection of engineered phage-mediated β-gal enzymatic activity</td><td>Ampicillin-resistant <i>E. coli</i> BLT5403</td><td>7 h</td><td>10<sup>1</sup> – 10<sup>7</sup> CFU mL<sup>-1</sup></td><td>10<sup>2</sup> CFU mL<sup>-1</sup></td><td>Kanamycin and ciprofloxacin at 100 µg mL<sup>-1</sup></td><td><i>E. coli</i> cultures</td><td>[36]</td></tr><tr><td>Colorimetric and electrochemical bioassay based on p-Benzoquinone</td><td>Trimethoprim- and erythromycin- resistant <i>E. coli</i></td><td>1 h</td><td>10<sup>3</sup> – 10<sup>9</sup> CFU mL<sup>-1</sup></td><td>10<sup>3</sup> – 10<sup>4</sup> CFU mL<sup>-1</sup></td><td><i>E. faecalis</i>, <i>S. mutans</i>, <i>S. aureus</i>, and <i>S. pullorum</i> at 10<sup>6</sup> CFU mL<sup>-1</sup></td><td>Microbial cultures</td><td>[37]</td></tr><tr><td>Hydration induced stress-based microcantilever DNA biosensor</td><td><i>Mycobacterium tuberculosis</i> and rifampicin-resistant gene <i>rpoB</i></td><td>1.5 h</td><td>20 ng mL<sup>-1</sup> – 200 fg mL<sup>-1</sup></td><td>2 pg mL<sup>-1</sup></td><td>Excess of digested bacterial DNA</td><td>Microbial cultures</td><td>[38]</td></tr><tr><td>SiO<sub>2</sub>@Fe<sub>3</sub>O<sub>4</sub> nanoparticles-based photoelectrochemical DNA biosensor</td><td>Penicillin-resistant genes <i>bla</i><sub>CTX-M-1</sub> and <i>bla</i><sub>TEM</sub></td><td>2 h</td><td>1 – 10000 nM</td><td>0.5 nM</td><td>3-base and whole-base mismatched sequences at 0.5 µM</td><td><i>E. coli</i> plasmids</td><td>[39]</td></tr><tr><td>Polyadenine tetrahedra-based DNA biosensor</td><td>Colistin-resistant gene <i>MCR-1</i> and Carbapenem-resistant genes <i>bla</i><sub>NDM</sub>, <i>bla</i><sub>KPC</sub>, and <i>bla</i><sub>OXA</sub></td><td>2 h</td><td>1 fM – 1 nM</td><td>10 fM</td><td><i>MCR-1</i>, <i>bla</i><sub>NDM</sub>, <i>bla</i><sub>KPC</sub>, and <i>bla</i><sub>OXA</sub>, <i>bla</i><sub>IMP</sub>, <i>bla</i><sub>VIM</sub></td><td><i>MCR-1</i> plasmid</td><td>[21]</td></tr><tr><td>Carbon nanotubes-based pencil graphite electrode for DNA biosensing</td><td><i>Bla</i><sub>OXA-48</sub>, <i>bla</i><sub>VIM</sub></td><td>30 min</td><td>1 – 15 µg/mL</td><td>2.5 pmol/ 50 µL</td><td>Non-complementary sequence of <i>Bla</i><sub>OXA-48</sub></td><td>Symmetric and asymmetric</td><td>[22]</td></tr></tbody></table>
<table><tr><td>Electrochemical microfluidic device based on carbon screen printed electrodes</td><td><i>E. coli</i> ATCC25922<br/><i>Bacillus subtilis</i> ATCC6051</td><td>3 h</td><td>10<sup>5</sup> – 10<sup>8</sup> CFU mL<sup>-1</sup></td><td>525 CFU mL<sup>-1</sup></td><td>Not available</td><td><i>Spiked urine</i></td><td>[24]</td></tr><tr><td>DNA probe-based electrochemical platinum biosensor on a microfluidic platform</td><td><i>bla</i><sub>CTX-M-8</sub></td><td>30 min</td><td>0.1 fM – 10 µM</td><td>10 pM</td><td>Non-complementary sequence <i>bla</i><sub>CTX-M-2</sub> at 10<sup>-18</sup> M – 10<sup>-8</sup> M</td><td><i>E. coli</i> lysates</td><td>[19] and this work</td></tr></table>
CONCLUSIONS

In this study, we developed a microfluidic DNA biosensor incorporating 6-mercaptohexanol and a probe DNA sequence, specifically aimed at detecting antibiotic resistance genes in *Escherichia coli* (*E. coli*). The use of platinum microelectrodes instead of gold in microfluidics represents a very promising alternative for nucleic acid biosensing but it requires the development of robust processes (microfabrication, surface treatment, and functionalization) for the selective and sensitive capture of DNA targets. The developed protocol in this work has the merit of high sensitivity while at the same time limiting non-specific interactions with complex samples. We demonstrated the electrochemical detection of target DNA sequences at concentrations as low as 10 fM, in the presence of diluted lysed *E. coli* cells through cyclic voltammetry and electrochemical impedance spectroscopy. While the detection limit in the presence of cell lysates shows a small reduction (from 0.1 fM to 10 fM) compared to ideal experimental conditions, the sensor's selectivity and sensitivity remain robust, as indicated by the nearly identical calibration slopes. Additionally, this applicative work is the first validation where we confirmed a recent model about non-linear mass transport phenomena on microchannel electrodes under flow condition. Future optimization work regarding on-chip lysis must be done to ensure a reliable quantification of resistant DNA in pathogenic strains of bacteria. Nonetheless, these findings provide valuable insights for the further development of a fully integrated portable microfluidic platform for bacterial cell lysis and quantification using antibiotic-resistant strains of bacteria.
# ASSOCIATED CONTENT

## Supporting Information

The Supporting Information is available free of charge on the ACS Publications website.

Supplementary tables summarizing all the data for several experimental conditions (50% *E. coli* lysates, 100% (undiluted) *E. coli* lysates, and without *E. coli* lysates).

## AUTHOR INFORMATION

### Corresponding Author

**Jean Gamby** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

Email: jean.gamby@cnrs.fr

### Authors

**Hiu Mun Man** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

**Choayb Omar** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.
**Martina Freisa** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

**David Bouville** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

**Téo Baptiste** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

**Anne-Marie Haghiri-Gosnet** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

**Hervé Jacquier** – Unité de Bactériologie, Département de Prévention, Diagnostic et Traitement des Infections, Hôpital Henri Mondor, APHP, Créteil, France, and EA 7380 DYNAMIC, Université Paris-Est Créteil, Créteil, France

**Isabelle Le Potier** – Team Smart Biosystems, MicroNanoBioFluidics department, UMR 9001 - Centre de Nanosciences et de Nanotechnologies (C2N), CNRS, Université Paris-Saclay, Université Paris Cité, 10 boulevard Thomas Gobert, 91120 Palaiseau, France.

## ACKNOWLEDGMENTS
The authors thank the DIMELEC project ANR-19-CE09-0016 for funding and RENATECH clean room facilities at C2N, Palaiseau, France. C.O. and M.F. thank the doctoral school "Electrical, Optical, Bio-Physics and Engineering" (ED575) and Paris-Saclay University for PhD grants. The authors are also grateful for Dr. Laurent Thouin (ENS Paris, Département de Chimie) for fruitful discussions about nonlinear mass transfer model understanding from hydrodynamics voltammetry.

## REFERENCES

(1) Antimicrobial Resistance Collaborators. Global Burden of Bacterial Antimicrobial Resistance in 2019: A Systematic Analysis. *Lancet* **2022**, *399* (10325), 629–655.
https://doi.org/10.1016/S0140-6736(21)02724-0.

(2) Castanheira, M.; Simner, P. J.; Bradford, P. A. Extended-Spectrum β-Lactamases: An Update on Their Characteristics, Epidemiology and Detection. *JAC Antimicrob Resist* **2021**, *3* (3), dlab092.
https://doi.org/10.1093/jacamr/dlab092.

(3) Kumar, A.; Roberts, D.; Wood, K. E.; Light, B.; Parrillo, J. E.; Sharma, S.; Suppes, R.; Feinstein, D.; Zanotti, S.; Taiberg, L.; Gurka, D.; Kumar, A.; Cheang, M. Duration of Hypotension before Initiation of Effective Antimicrobial Therapy Is the Critical Determinant of Survival in Human Septic Shock*. *Critical Care Medicine* **2006**, *34* (6), 1589.
https://doi.org/10.1097/01.CCM.0000217961.75225.E9.

(4) Pellitero, M. A.; Shaver, A.; Arroyo-Currás, N. Critical Review—Approaches for the Electrochemical Interrogation of DNA-Based Sensors: A Critical Review. *J. Electrochem. Soc.* **2019**, *167* (3), 037529. https://doi.org/10.1149/2.0292003JES.

(5) Cagnin, S.; Caraballo, M.; Guiducci, C.; Martini, P.; Ross, M.; SantaAna, M.; Danley, D.; West, T.; Lanfranchi, G. Overview of Electrochemical DNA Biosensors: New Approaches to Detect the Expression of Life. *Sensors (Basel)* **2009**, *9* (4), 3122–3148. https://doi.org/10.3390/s90403122.

(6) Ozer, T.; Geiss, B. J.; Henry, C. S. Review—Chemical and Biological Sensors for Viral Detection. *J. Electrochem. Soc.* **2020**, *167* (3), 037523. https://doi.org/10.1149/2.0232003JES.

(7) Hai, X.; Li, Y.; Zhu, C.; Song, W.; Cao, J.; Bi, S. DNA-Based Label-Free Electrochemical Biosensors: From Principles to Applications. *TrAC Trends in Analytical Chemistry* **2020**, *133*, 116098. https://doi.org/10.1016/j.trac.2020.116098.

(8) Siavashy, S.; Soltani, M.; Rahimi, S.; Hosseinali, M.; Guilandokht, Z.; Raahemifar, K. Recent Advancements in Microfluidic-Based Biosensors for Detection of Genes and Proteins: Applications and Techniques. *Biosensors and Bioelectronics: X* **2024**, *19*, 100489.
https://doi.org/10.1016/j.biosx.2024.100489.

(9) Shen, H.; Yang, J.; Chen, Z.; Chen, X.; Wang, L.; Hu, J.; Feihu, J.; Xie, G.; Feng, W. A Novel Label-Free and Reusable Electrochemical Cytosensor for Highly Sensitive Detection and Specific Collection of CTCs. *Biosensors and Bioelectronics* **2016**, *81*.
https://doi.org/10.1016/j.bios.2016.03.048.

(10) Deng, K.; Liu, X.; Li, C.; Huang, H. Sensitive Electrochemical Sensing Platform for microRNAs Detection Based on Shortened Multi-Walled Carbon Nanotubes with High-Loaded Thionin. *Biosensors and Bioelectronics* **2018**, *117*, 168–174. https://doi.org/10.1016/j.bios.2018.05.055.

(11) Xiao, Y.; Lubin, A. A.; Baker, B. R.; Plaxco, K. W.; Heeger, A. J. Single-Step Electronic Detection of Femtomolar DNA by Target-Induced Strand Displacement in an Electrode-Bound Duplex. *Proc. Natl. Acad. Sci. U.S.A.* **2006**, *103* (45), 16677–16680.
https://doi.org/10.1073/pnas.0607693103.
(12) Mazaafrianto, D. N.; Ishida, A.; Maeki, M.; Tani, H.; Tokeshi, M. Label-Free Electrochemical Sensor for Ochratoxin A Using a Microfabricated Electrode with Immobilized Aptamer. *ACS Omega* **2018**, *3* (12), 16823–16830. https://doi.org/10.1021/acsomega.8b01996.

(13) Gooding, J. J.; Darwish, N. The Rise of Self-Assembled Monolayers for Fabricating Electrochemical Biosensors--an Interfacial Perspective. *Chem Rec* **2012**, *12* (1), 92–105. https://doi.org/10.1002/tcr.201100013.

(14) Patolsky, F.; Lichtenstein, A.; Willner, I. Electronic Transduction of DNA Sensing Processes on Surfaces: Amplification of DNA Detection and Analysis of Single-Base Mismatches by Tagged Liposomes. *J. Am. Chem. Soc.* **2001**, *123* (22), 5194–5205. https://doi.org/10.1021/ja0036256.

(15) Cardoso, A. R.; Moreira, F. T. C.; Fernandes, R.; Sales, M. G. F. Novel and Simple Electrochemical Biosensor Monitoring Attomolar Levels of miRNA-155 in Breast Cancer. *Biosens Bioelectron* **2016**, *80*, 621–630. https://doi.org/10.1016/j.bios.2016.02.035.

(16) Long, Y.-T.; Li, C.-Z.; Kraatz, H.-B.; Lee, J. S. AC Impedance Spectroscopy of Native DNA and M-DNA. *Biophysical Journal* **2003**, *84* (5), 3218–3225. https://doi.org/10.1016/S0006-3495(03)70046-0.

(17) Radi, A.-E.; Acero Sánchez, J. L.; Baldrich, E.; O'Sullivan, C. K. Reagentless, Reusable, Ultrasensitive Electrochemical Molecular Beacon Aptasensor. *J. Am. Chem. Soc.* **2006**, *128* (1), 117–124. https://doi.org/10.1021/ja053121d.

(18) Freisa, M.; Poujouly, C.; Le Potier, I.; Dinh, T. H. N.; Couraud, L.; Bouville, D.; Sella, C.; Thouin, L.; Gamby, J. Highlighting the Impact of Blocking Monolayers on DNA Electrochemical Sensors. Theoretical and Experimental Investigations under Flow Conditions. *Electrochimica Acta* **2024**, *505*, 145006. https://doi.org/10.1016/j.electacta.2024.145006.

(19) Omar, C.; Freisa, M.; Man, H. M.; Kechekche, D.; Dinh, T. H. N.; Haghiri-Gosnet, A.-M.; Le Potier, I.; Gamby, J. Optimizing Tris(2-Carboxyethyl) Phosphine and Mercaptohexanol Concentrations for Thiolated Oligonucleotide Immobilization on Platinum Electrodes in Microfluidic Platforms. *Langmuir* **2024**, *40* (50), 26371–26788. https://doi.org/10.1021/acs.langmuir.4c03566.

(20) Wan, C.; Qu, A.; Li, M.; Tang, R.; Fu, L.; Liu, X.; Wang, P.; Wu, C. Electrochemical Sensor for Directional Recognition and Measurement of Antibiotic Resistance Genes in Water. *Anal. Chem.* **2022**, *94* (2), 732–739. https://doi.org/10.1021/acs.analchem.1c03100.

(21) Song, Y.; Feng, J.; Wang, X.; Wen, Y.; Xu, L.; Huo, Y.; Wang, L.; Tao, Q.; Yang, Z.; Liu, G.; Chen, M.; Li, L.; Yan, J. A Multi-Channel Electrochemical Biosensor Based on Polyadenine Tetrahedra for the Detection of Multiple Drug Resistance Genes. *Analyst* **2024**, *149* (12), 3425–3432. https://doi.org/10.1039/D4AN00488D.

(22) Subak, H.; Yilmaz, F. F.; Ozkan-Ariksoysal, D. Ready-to-Use Diagnostic Kit Based on Electrochemical Nanobiosensor for Antibiotic Resistance Gene Determination. *Microchemical Journal* **2024**, *204*, 110976. https://doi.org/10.1016/j.microc.2024.110976.

(23) Jiang, H.; Sun, Z.; Guo, Q.; Weng, X. Microfluidic Thread-Based Electrochemical Aptasensor for Rapid Detection of Vibrio Parahaemolyticus. *Biosensors and Bioelectronics* **2021**, *182*, 113191. https://doi.org/10.1016/j.bios.2021.113191.

(24) Gopalakrishnan, S.; Mall, D.; Pushpavanam, S.; Karmakar, R. Rapid Antimicrobial Susceptibility Testing Using Carbon Screen Printed Electrodes in a Microfluidic Device. *Sci Rep* **2025**, *15* (1), 5133. https://doi.org/10.1038/s41598-024-84286-3.

(25) Horny, M.-C.; Freisa, M.; Poujouly, C.; Gonzalez-Losada, P.; Le Gall, J.; Dinh, T.-H.-N.; Bouville, D.; Le Potier, I.; Pallandre, A.; Gamby, J. Electrochemical Nucleic Acid Biosensor on a Microfluidic Chip to Understand the Coupling between Electrochemistry and Microfluidics. *J. Chem. Educ.* **2024**, *101* (2), 605–611. https://doi.org/10.1021/acs.jchemed.3c01054.

(26) Horny, M.-C.; Lazerges, M.; Siaugue, J.-M.; Pallandre, A.; Rose, D.; Bedioui, F.; Deslouis, C.; Haghiri-Gosnet, A.-M.; Gamby, J. Electrochemical DNA Biosensors Based on Long-Range Electron Transfer: Investigating the Efficiency of a Fluidic Channel Microelectrode Compared to an Ultramicroelectrode in a Two-Electrode Setup. *Lab Chip* **2016**, *16* (22), 4373–4381. https://doi.org/10.1039/C6LC00869K.

(27) Poujouly, C.; Le Gall, J.; Freisa, M.; Kechkeche, D.; Bouville, D.; Khemir, J.; Gonzalez-Losada, P.; Gamby, J. Microfluidic Chip for the Electrochemical Detection of MicroRNAs: Methylene
Blue Increasing the Specificity of the Biosensor. *Front. Chem.* **2022**, *10*.
https://doi.org/10.3389/fchem.2022.868909.

(28) Poujouly, C.; Gonzalez-Losada, P.; Boukraa, R.; Freisa, M.; Le Gall, J.; Bouville, D.; Deslouis, C.; Gamby, J. Diffusion-Convection Impedance for a Micro-Band Electrode under Microfluidic Conditions. *Electrochemistry Communications* **2022**, *137*, 107262.
https://doi.org/10.1016/j.elecom.2022.107262.

(29) Horny, M.-C.; Billon, F.; Deslouis, C.; lazerges, M.; Dupuis, V.; Siaugue, J.-M.; Pailleret, A.; Gamby, J. Amorphous Carbon Nitride Microband Integrated in a Microfluidic Device for DNA Biosensors Applications. *Journal of Electroanalytical Chemistry* **2021**, *895*, 115395.
https://doi.org/10.1016/j.jelechem.2021.115395.

(30) Watson, W.; Orazem, M. E. EIS: Measurement Model Program. **2020**.
https://doi.org/10.17605/OSF.IO/NFMJ8.

(31) Levich, V. G. *Physicochemical Hydrodynamics*; Prentice-Hall international series in the physical and chemical engineering sciences; Prentice-Hall: Englewood Cliffs, N.J., 1962.

(32) Compton, R. G.; Fisher, A. C.; Wellington, R. G.; Dobson, P. J.; Leigh, P. A. Hydrodynamic Voltammetry with Microelectrodes: Channel Microband Electrodes; Theory and Experiment. *J. Phys. Chem.* **1993**, *97* (40), 10410–10415. https://doi.org/10.1021/j100142a024.

(33) Amatore, C.; Da Mota, N.; Sella, C.; Thouin, L. Theory and Experiments of Transport at Channel Microband Electrodes under Laminar Flows. 1. Steady-State Regimes at a Single Electrode. *Anal. Chem.* **2007**, *79* (22), 8502–8510. https://doi.org/10.1021/ac070971y.

(34) Shehadul Islam, M.; Aryasomayajula, A.; Selvaganapathy, P. R. A Review on Macroscale and Microscale Cell Lysis Methods. *Micromachines* **2017**, *8* (3), 83.
https://doi.org/10.3390/mi8030083.

(35) Kelley, S. O.; Barton, J. K.; Jackson, N. M.; McPherson, L. D.; Potter, A. B.; Spain, E. M.; Allen, M. J.; Hill, M. G. Orienting DNA Helices on Gold Using Applied Electric Fields. *Langmuir* **1998**, *14* (24), 6781–6784. https://doi.org/10.1021/la980874n.

(36) Chen, J.; Alcaine, S. D.; Jackson, A. A.; Rotello, V. M.; Nugen, S. R. Development of Engineered Bacteriophages for Escherichia Coli Detection and High-Throughput Antibiotic Resistance Determination. *ACS Sens.* **2017**, *2* (4), 484–489.
https://doi.org/10.1021/acssensors.7b00021.

(37) Sun, J.; Warden, A. R.; Huang, J.; Wang, W.; Ding, X. Colorimetric and Electrochemical Detection of Escherichia Coli and Antibiotic Resistance Based on a P-Benzoquinone-Mediated Bioassay. *Anal. Chem.* **2019**, *91* (12), 7524–7530. https://doi.org/10.1021/acs.analchem.8b04997.

(38) Domínguez, C. M.; Kosaka, P. M.; Sotillo, A.; Mingorance, J.; Tamayo, J.; Calleja, M. Label-Free DNA-Based Detection of Mycobacterium Tuberculosis and Rifampicin Resistance through Hydration Induced Stress in Microcantilevers. *Anal. Chem.* **2015**, *87* (3), 1494–1498.
https://doi.org/10.1021/ac504523f.

(39) Li, X.; Lu, J.; Feng, L.; Zhang, L.; Gong, J. Smart pH-Regulated Switchable Nanoprobes for Photoelectrochemical Multiplex Detection of Antibiotic Resistance Genes. *Anal. Chem.* **2020**, *92* (16), 11476–11483. https://doi.org/10.1021/acs.analchem.0c02839.
TOC – Graphical abstract

Antimicrobial resistance gene sequence detection

<figure><img src="image_9.png" /></figure>