# Optimizing self-assembled monolayer construction for aptasensors and preventing false signals. The case of erythromycin detection

Teodora Lupoi, Sarah Baccara, Yann R. Leroux, Bogdan Feier, Mathieu Pasturel, Jean-Luc Cercus, Cecilia Cristea, Florence Geneste

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## ► To cite this version:

Teodora Lupoi, Sarah Baccara, Yann R. Leroux, Bogdan Feier, Mathieu Pasturel, et al.. Optimizing self-assembled monolayer construction for aptasensors and preventing false signals. The case of erythromycin detection. Electrochimica Acta, 2025, 525, pp.146097. 10.1016/j.electacta.2025.146097. hal-05080397

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Optimizing self-assembled monolayer construction for aptasensors and
preventing false signals. The case of erythromycin detection.

Teodora Lupoi¹,², Sarah Baccara³, Yann R. Leroux¹, Bogdan Feier², Mathieu Pasturel¹, Jean-Luc Cercus¹, Cecilia Cristea²*, Florence Geneste¹*

¹Univ Rennes, CNRS, ISCR, UMR 6226, F-35000 Rennes, France

²“Iuliu Hatieganu” University of Medicine and Pharmacy, Faculty of Pharmacy, Department of Analytical Chemistry, 400349, Cluj-Napoca, Romania

³University of Carthage, Faculty of Science of Bizerte, Laboratory of Application of Chemistry to Natural Resources and Substances and the Environment, 7021 Bizerte, Tunisia

**Abstract**

Although an aptamer selected for erythromycin (ERY) was synthesized and characterized, no electrochemical sensor has been reported yet. This study is the first development of an aptamer-based electrochemical sensor for ERY detection. Two immobilization platforms of the thiol-modified aptamer were evaluated: a gold-coated carbon screen-printed electrode (C-SPE) and a pure [111] gold electrode. Label-free ERY detection was performed by differential pulsed voltammetry (DPV) with $K_4[Fe(CN)_6]/K_3[Fe(CN)_6]$ as a redox probe. Instability in surface chemistry initially led to a false signal on the C-SPE in the presence of ERY. To address this, we examined the stability of the mixed self-assembled monolayer (SAM) formed by the aptamer and a blocking agent. Extended thiol immobilization times enhanced the mixed SAM stability but did not improve the sensor analytical performance. Applying the optimized SAM construction to the pure gold electrode resulted in a stable aptasensor with a stable electrochemical signal during blank incubation. A calibration curve was obtained on the pure gold platform in an ERY concentration range of $1 \times 10^{-6}$ M to $2 \times 10^{-4}$ M with a limit of detection of $3.2 \times 10^{-7}$ M. In addition to the advancements made in the electrochemical detection of ERY using an aptamer-based sensor, this work provides insights into the stability of MCH/aptamer layers depending on their construction method.

**Keywords:** aptamer-based electrochemical sensor, self-assembled monolayers, prevention of false signals, stability enhancement, erythromycin detection.

**1. Introduction.**

Electrochemical aptamer-based (E-AB) sensors are an attractive alternative to antibody-based detection techniques because they are chemically synthesized, easily modified, adaptable to a wide range of targets including ions, small molecules, nucleic acids, proteins, and cells, and are relatively inexpensive and stable [1-3]. This last point is one of the most important advantages of this type of biosensor, as it allows for long-term storage periods, use in non-physiological environments, and even regeneration to achieve sustainable biosensors. The general detection principle of E-AB sensors is that target molecules induce conformational changes in the aptamer through various interactions between the oligonucleotides and the
target, such as electrostatic, Van der Waals, hydrophobic, hydrogen bonding interactions. These conformational changes lead to an increase or decrease in the electrochemical signal of a redox probe, either in solution or attached to the aptamer. The sensitivity of the E-AB sensors depends directly on the difference in signal value in the absence and presence of the target, which means that signal stability is crucial for achieving high accuracy and repeatability.

Although numerous articles deal with the preparation of new E-AB sensors, few have addressed the issue of signal drift. The instability of the signal has been attributed to protein fouling that occurs when analyzing biological samples such as blood [4] or to false signals generated when methylene blue is used as a redox reporter [5]. Moreover, signal drift has also been observed in model analytical media after repeated measurements or even after simple washing with buffer solution [6, 7]. It has been shown that the desorption of the SAM monolayer composed of the aptamer and a blocking agent from the gold electrode is responsible for this undesired signal change [4, 8]. This presents a challenge for the development of E-AB sensors, as recently highlighted [9].

The most used blocking agent is 6-mercapto-1-hexanol (MCH). Its role is to reduce non-specific adsorption and to allow the immobilized aptamer (APT) to adopt an appropriate conformation by diluting it on the gold surface [10, 11]. Recent efforts have been made to explore new co-adsorbates that improve electrochemical signal and stability. Although thiols with C11 chains are known to produce more stable SAM, they also increase the resistance to charge transfer and the target accessibility to the aptamer [12, 13]. Intermediate chain lengths, such as C6, offer a good compromise between stability and sensitivity to E-AB sensors [14]. The nature of the end group of the thiol alkyl chain also plays an important role in biofouling resistance. Phosphatidylcholine moieties [15] and zwitterionic structures [12], for example, significantly improve baseline stability due to their antifouling properties. It has also been shown that hydrophobic thiols lead to higher monolayer packing density than hydrophilic ones, thus increasing SAM stability [16-18]. Interestingly, recent studies have reported that binding the aptamer to gold via a gold-alkyne bond results in a more stable E-AB sensor compared with gold-thiol binding [19]. The authors have attributed this enhanced stability to the lower sensitivity of the gold-alkyne bond to oxidation at ambient temperature. Other strategies to improve SAM stability, such as di-trithiol anchoring groups and crosslinking of the layer, have also been reported [16].

Another factor affecting SAM stability is the electroanalytical method used. Studies have shown that reducing the potential window during analysis improves signal stability [4]. Accounting for changes in peak-to-peak separation due to electron-transfer variations in cyclic voltammetry (CV) has also reduced sensor performance variability compared with pulsed techniques [20].

The formation of the SAM is another crucial factor influencing signal stability. It depends on the crystallinity of the gold (Au) surface [21] and its pre-treatment [22]. Several parameters that influence SAM formation of oligonucleotides have been investigated, including aptamer concentration, the aptamer-to-blocking agent ratio [12], incubation time to enhance surface coverage [11] and buffer ionic strength to prevent electrostatic repulsion between adsorbed oligonucleotides [23]. Different methods have been tested for forming the oligonucleotide/MCH mixed layer. The most common approach is to add MCH after
oligonucleotide immobilization to displace non-specifically adsorbed macromolecules. However, it has been reported that using the reverse order may be more efficient in preventing physisorption [24]. Co-deposition has also been explored, showing a greater signal change in the presence of the target without reducing sensor stability, even when exposed to a blood sample at 37°C, compared with the conventional step-by-step approach [25]. Additionally, pulse-assisted thiol exchange has been used to improve adsorption kinetics and achieve high surface coverage [26-28]. However, while many articles have focused on the effect of the modification protocol on surface coverage, few have investigated the stability of the blank as a function of the modification process. Impedimetric measurements highlighted the importance of conditioning the electrode in the buffered measurement electrolyte for 12 h to achieve a reproducible signal [8]. The authors have demonstrated that the impedance drift was primarily due to the gradual reorganization of the MCH SAM.

Erythromycin (ERY) is a widely used macrolide antibiotic, selected as a target for this study due to its environmental relevance. The presence of ERY in water bodies increases the risk of antimicrobial resistance and ecotoxicity. Monitoring ERY levels can thus serve as an important indicator of pollution from emerging contaminants, emphasizing the need for its detection [29]. Notably, an erythromycin-selective aptamer has recently been synthesized using the SELEX method [30]. During our development of an erythromycin E-AB sensor with this new aptamer on screen-printed carbon electrodes modified with gold nanoparticles, we observed that the drift in the blank signal could be mistaken for changes associated with the target molecule, even after optimizing the aptamer immobilization conditions. We identified a link between the blank signal stabilization and the formation of the mixed aptamer/MCH SAM by varying parameters such as the order of thiol derivatives introduction, reaction times, mixing open circuit voltage (OCV) and applied potential methods. Our findings showed that the blank signal was minimized when using a gold surface instead of carbon-gold mixed surfaces. Stabilizing and minimizing the blank signal ultimately allowed us to observe a significant signal change in the presence of ERY. This study will focus on the stability of the MCH/APT system and demonstrates that the MCH/APT/Au electrode has the potential to detect ERY.

## 2. Materials and methods.

### 2.1 Reagents and materials.

KCl, NaCl, MgCl₂, HCl, H₂SO₄, 2-Amino-2-(hydroxymethyl)-1,3-propanediol (TRIS), 6-Mercapto-1-hexanol (MCH), ERY, nuclease-free water, K₄[Fe(CN)₆], K₃[Fe(CN)₆], HAuCl₄, and H₂O₂ were purchased from Sigma-Aldrich. All the reagents had analytical purity and were used without further purification.

The sequence and affinity constant of the aptamer (APT) specific for ERY were taken from literature [30]: Kd = 20 ± 9 nM and the sequence was 5'-AGG-AAT-TCA-CGT-CTC-ACT-GGA-TTC-ACG-CAC-GCC-AAG-GAC-TGC-ACT-TAA-GGT-TAG-ATA-GCC-CCA-TGC-AGT-GAG-TCA-GGA-TAT-CG-C6-SH-3'. The APT was synthesized by Eurogentec Belgium and was received as a lyophilized powder which was solubilized in TRIS buffer pH 7.4 (10 mM Tris, 100 mM NaCl, 100 mM KCl, 5 mM MgCl₂) prepared in nuclease-free water. The resulting APT stock solution had a 20 µM concentration and was divided into 20 µL aliquots stored at -20 °C. The other solutions were prepared in ultrapure water (18.2 MΩ, Millipore Simplicity).
## 2.2 Preparation of the modified electrodes and analyte detection.

Gold nanoparticles (AuNPs) were deposited on C-SPE by CV using a 2 mM HAuCl₄ solution prepared in 0.5 M H₂SO₄ by applying 40 cycles from 1 V to -0.2 V vs Ag pseudoreference at a scan rate of 0.1 V s⁻¹ [31]. After the AuNPs deposition, the surface was washed with ultrapure water. When using gold wafers, the substrate was thoroughly rinsed with acetone and ultrapure water and treated for 5 min with freshly prepared Piranha solution (H₂SO₄:H₂O₂-3:1, v:v). The surface was then carefully washed with ultrapure water to eliminate any Piranha traces.

For APT immobilization, the 20 µM APT solution stored at -20 °C was diluted to the desired concentration with TRIS buffer pH 7.4 and submitted to a thermal treatment for 5 min at 95°C followed by 1 min at -20 °C to ensure the proper folding of the APT [32].

Two immobilization strategies of the thiol-modified APT were employed, an electrochemically-assisted method and a standard OCV one. Jambrec et al. proved that the immobilization kinetics of single-stranded DNA molecules can be enhanced when switching rapidly between a positive and a negative potential [27, 33]. We adapted the protocol with respect to the pzc of the gold surface (0.2 V) with the following values: step 1 = 0.5 V, step 2 = -0.2 V, step duration = 45 ms, total duration = 5 minutes. The standard OCV strategy involved leaving the clean gold substrates in the APT solution at 4°C for 12 hours. The remaining gold sites were blocked with a 1 µM MCH solution prepared in TRIS buffer pH 7.4, applying the pulsed method for 5 minutes and then washing the surface with TRIS buffer. Other strategies of forming the mixed APT – MCH SAM were tested, with the MCH immobilized before the APT and the two immobilized simultaneously. The APT and MCH concentrations were also optimized.

After the two thiols immobilization, the electrode was rinsed for one minute with ultrapure water and the stability of the blank signal was investigated by CV analysis in [Fe(CN)₆]⁴⁻/³⁻ solution (1:1 5 mM) prepared in TRIS after successive incubations in the same buffer. The incubation with ERY in TRIS buffer pH 7.4 was performed without stirring at room temperature for the optimal time of 20 min. The electrode was rinsed 3 times with 1 mL TRIS buffer pH 7.4 to remove any non-specifically adsorbed ERY and re-analyzed in the [Fe(CN)₆]⁴⁻/³⁻ solution. The signal linked to the presence of ERY was calculated as a percentage of increase or decrease in current intensity corresponding to the redox probe after incubation using the equation (1):

$$S_{ERY} = \frac{|I_{BLANK} - I_{ERY}|}{I_{BLANK}} \times 100 \quad (1)$$

## 2.3 Analysis.

The electrochemical measurements were carried out with an Origalys OGF500 potentiostat coupled to an impedance module. The first support used for the sensor construction was a screen-printed electrode (C-SPE) containing carbon working and counter electrodes and a silver pseudo-reference electrode supplied by Metrohm DropSens. The second support was a silicon wafer covered with a 100 nm gold layer purchased from Soka technology. The gold surface was integrated into an electrochemical cell with an Ag/AgCl/saturated KCl reference and Pt wire counter-electrode.
Each modification step was characterized by CV, differential pulse voltammetry (DPV), and electrochemical impedance spectroscopy (EIS) in a mixture of K₄[Fe(CN)₆]/ K₃[Fe(CN)₆] (1:1, 5 mM) prepared in TRIS buffer pH 7.4. For the CV measurements, the potential was scanned from 0 V to 0.6 V vs Ag pseudo-reference at a scan rate of 0.1 V s⁻¹. In DPV, the parameters were optimized on a C-SPE in a K₄[Fe(CN)₆]/ K₃[Fe(CN)₆] solution prepared in TRIS (Table S1). The initial potential was -0.2 V and the end potential 0.5 V, the scan rate 0.01 V s⁻¹, with a pulse height of 0.1 V and pulse width of 25 ms. Because of the instability of the Ag pseudo-reference, the peak potential in CV and DPV was corrected vs a saturated calomel electrode (SCE). The EIS measurements were conducted on a frequency range of 10000 to 0.1 Hz with a wave amplitude of 0.01 V and open circuit potential conditions.

The surface morphology of the electrode was analyzed by JSM-7100F Field Emission Scanning Electron microscopy equipped by EDS (Energy Dispersive X-Ray Spectroscopy) Detector SDD type (Silicon Drift Detector) X-Max 80 mm² Oxford Instruments AZtecEnergy. The accelerating voltage was 10 kV with a beam current of 1 nA and a spectra collection time of 100 s.

The affinity of the aptamer for ERY was determined by measuring the equilibrium dissociation constant (Kd) using surface plasmon resonance (SPR). All SPR experiments were conducted on a three-channel Biosensing BI-2500 instrument (Biosensing Instrument Inc.)

X-ray diffraction patterns have been measured in the 30-120° angular domain (step size ≈ 0.029°, integrated counting time = 89.5 s/step) on a Bruker D8 Advance diffractometer working with monochromatized Cu Kα1 radiation (λ= 1.5406 Å) and equipped with a LynxEye detector. Preliminary fitting of the patterns by the Le Bail method has been performed using the FullProf software [34].

**3. Results and discussions.**

3.1 C-SPE electrode functionalization and characterization.

The construction of the aptasensor on the C-SPE support involves three modification steps: AuNPs deposition, aptamer immobilization and blocking agent immobilization, as illustrated in Scheme 1.

<figure><img src="image_3.png" /><figcaption>Scheme 1. Fabrication of the aptasensor and erythromycin detection principle.</figcaption></figure>
### 3.1.1. Gold deposition.

The first functionalization step aimed to cover the surface of the C-SPE with AuNPs, which act as anchoring points for the aptamer. Moreover, AuNPs significantly increase the conductivity of the electrode and the active surface area thanks to their high surface-to-volume ratio. AuNPs were electrodeposited onto carbon surfaces by electrochemical reduction from an HAuCl₄ solution, as previously reported in the literature as a fast and efficient preparation method [35]. Figure 1A shows the CV used for AuNPs deposition from a 2 mM HAuCl₄ solution prepared in 0.5 M H₂SO₄ while scanning the potential from 1 V to -0.2 V, with a scan rate of 0.1 V s⁻¹. On the forward scan, a peak corresponding to the reduction of Au³⁺ to Au⁰ was observed at 0.12 V. This step corresponds to the nucleation phase, during which gold is deposited on the carbon electrode, forming nuclei that serve as seeds for particles growth. Starting with the second scan, the reduction peak was shifted to 0.38 V due to the preferential deposition of Au on the already formed AuNPs because of the lower energy required to deposit gold onto gold. This phase corresponds to particle growth. The resulting platform is called AuNPs/C-SPE. The number of cycles of gold deposition varied from 30 to 50 cycles to investigate the influence on the active surface area. The morphology of the surfaces was examined by SEM and images of the bare and modified surfaces after 30, 40 and 50 cycles of gold deposition are gathered in Figure 1B and Figure S1. For 30 cycles, the average diameter of the particles was 200 nm with 13% surface coverage. When using 40 cycles, particles with similar size were obtained (182 nm), but the surface coverage was increased to 19% (Figure S2 A and B). In both cases, the particles were uniformly distributed, and their diameters followed a Gaussian distribution. When using 50 cycles, two distinct regions were observed, one with particles of 245 nm diameter and another with 73 nm diameter, indicating high variability in particle size and an uncontrolled deposition process (Figure S2 C).

The active surface area was also estimated using the Randles-Sevcik equation (2) for one electron exchange using a 5 mM K₄[Fe(CN)₆] solution in TRIS buffer:

$$I_p = 2.69 \times 10^5 A C \sqrt{D \nu} \quad (2)$$

where $I_p$ is the anodic peak current (A), A is the area of the electrode (cm²), C is the concentration of the redox probe (mol/cm³), D is the diffusion coefficient (cm² s⁻¹), ν is the scan rate (V s⁻¹) and the constant with a value of 2.69 × 10⁵ has units of C mol⁻¹V⁻¹/².

The calculated surface area of the bare C-SPE was 3.6 ± 0.1 mm². The surface area after gold deposition was 5.7 ± 0.2 mm² for 30 cycles, 6.4 ± 0.3 mm² for 40 cycles, and 6.0 ± 0.1 mm² for 50 cycles. The most important increase of 74% in surface area was obtained for 40 deposition cycles and this number was kept for the future aptasensor fabrication protocol. A larger surface area is desirable for aptamer immobilization, as more gold facilitates the attachment of a greater number of aptamers on the surface.
<figure><img src="image_4.png" /><figcaption>Figure 1. A) CV of the gold deposition from a 2 mM HAuCl₄ solution in 0.5 M H₂SO₄, scan rate 0.1 V s⁻¹ (40 cycles), B) SEM images of the C-SPE before and after 40 cycles of gold deposition at different magnifications.</figcaption></figure>

The modified surface after 40 cycles was characterized electrochemically by registering the signal before and after gold deposition using DPV and EIS in 5 mM K₄[Fe(CN)₆]/K₃[Fe(CN)₆], as shown in Figure 2. An increase in peak current intensity was observed after the deposition of AuNPs, due to the enhancement of the active surface area of the electrode. The modification step was supported by EIS analysis, where a reduction in charge transfer resistance (Rct) from 334 to 103 Ω was noted after the deposition step due to a higher conductivity of gold.

<figure><img src="image_5.png" /><figcaption>Figure 2. Electrochemical characterization by A) DPV and B) EIS in 5 mM K₄[Fe(CN)₆]/K₃[Fe(CN)₆] in TRIS buffer of the bare electrode after washing with ethanol</figcaption></figure>
(black), after AuNPs deposition (orange), 1 µM APT immobilization by MPA (blue), 1 µM MCH immobilization by MPA (green).

The crystallinity of the AuNPs deposited on the C-SPE was investigated. For the AuNPs, a pyrolyzed photoresist film (PPF) was used as a substrate to replace the C-SPE due to its simplicity. Figure S3 shows the X-ray diffraction patterns of gold deposited on PPF (green) together with the simulated one using the crystallographic data of gold (blue). In the experimental pattern, the silicon substrate Bragg peaks are dominating, with a [100] orientation for the substrate used for PPF coating.

The XRD pattern of gold nanoparticles deposited on PPF is more characteristic of a random isotropic powder, despite the slightly more intense (111) and (222) peaks hinting at a bit of preferential orientation. The small number and intensity of available peaks do not enable an accurate determination of the crystallite size, but preliminary fits using the Le Bail method indicate diffracting domains around 30 nm.

### 3.1.2. APT immobilization.

After the deposition of AuNPs, the aptamer functionalized with a thiol group was immobilized on the gold surface, obtaining the APT/AuNPs/C-SPE platform. The immobilization step is of utmost importance as it can influence the subsequent affinity for the target, the stability of the sensor, and its reproducibility. Considering this, two immobilization strategies were tested. The first method is the most used in the literature and involves the spontaneous formation of DNA SAM at OCV overnight. The second method is assisted by fast alternating pulses and was discussed and applied by Jambrec *et al* [27]. Applying positive potentials relative to the potential of zero charge (pzc) of the gold electrode will attract anions toward the surface and repel cations into the bulk solution. The cations that shield the negatively charged aptamer will be repelled from the surface and the aptamer will be attracted to the positive surface until it lies flat. Then the applied negative potential will repel the anions and force the aptamer to have an upright orientation. Fast switching between a positive and a negative potential with respect to the pzc value will create a stirring effect and accelerate the immobilization kinetics, leading to more reproducible platforms and better surface coverage [27]. Therefore, the multipulse amperometry (MPA) method was applied relative to the pzc value of 0.2 V of the C-SPE/AuNPs with a positive pulse of 0.5 V vs Ag and negative pulse of -0.2 V vs Ag, a pulse duration of 45 ms, and a total duration of 5 minutes in a 1 µM APT solution in TRIS buffer.

As seen in Figure 2, the immobilization of the APT can be confirmed by electrochemistry. The APT is a biological macromolecule with insulating properties and its immobilization leads to a decrease in current intensity, and an increase in Rct. Moreover, the APT is negatively charged causing repulsion with the negatively charged [Fe(CN)]³⁻/₄⁻ redox probe. The immobilization method and the concentration of the aptamer were optimized. Since a higher aptamer density is preferred, greater passivation is sought. The passivation percentage in DPV measurements after APT immobilization was used as a measure of aptamer loading, with higher passivation indicating a greater amount of APT immobilized on the surface. For the immobilization strategy, both the OCV and MPA methods produced similar results when using a 1 µM APT solution in TRIS, as shown in Figure 3A. The OCV method achieved passivation of 47%, while the MPA method resulted in a slightly higher passivation of 55%, relative to the peak current on AuNPs/C-SPE. The MPA method was preferred in these first
experiments conducted with AuNPs/C-SPE electrodes due to its significantly shorter duration (5 minutes) compared to the 12 hours required for overnight immobilization using the OCV method.

Another important parameter to consider is the APT concentration in the immobilization solution. Ideally, a high density of immobilized aptamers is desired without excessive use of aptamer. Passivation percentages were calculated following APT immobilization with solutions of 0.1, 0.5, 1, and 2 µM, as summarized in Figure 3B. The passivation increased with maximum coverage achieved at 1 µM, the concentration selected for future immobilization.

<figure><img src="image_6.png" /><figcaption>Figure 3. The passivation percentage calculated from the peak current in DPV following the APT and MCH immobilization compared with the AuNPs/C-SPE surface, for the optimization of A) the method of immobilization of 1 µM APT by OCV for 12h or MPA, B) the APT concentration (MPA immobilization), C) the ratio between APT (1 µM) and MCH (MPA immobilization).</figcaption></figure>

### 3.1.3. Blocking agent immobilization.

The final step in aptasensor construction involves blocking the unoccupied gold sites with a co-adsorbed thiol. This backfilling aims to remove any non-specifically adsorbed aptamer, improve the organization of the mixed SAM, and to produce an anti-fouling effect. MCH was chosen as the co-thiol because its length matches that of the C6 spacer on the aptamer, which is introduced between the oligonucleotide chain and the thiol group [36]. Additionally, the pulse-assisted method MPA has been shown to enhance the immobilization of uncharged thiol molecules [27] and was therefore utilized for the immobilization of MCH. MCH was immobilized from the TRIS buffer solution using the same pulse-assisted method used for the APT immobilization. SAMs are dynamic structures and replacement of one thiol by another is a frequently observed phenomenon [37, 38]. To avoid the replacement of the APT by MCH, the concentration of MCH was optimized. Different ratios of the APT to MCH at 10:1, 1:1, 1:5, and 1:50 ratios were tested, as shown in Figure 3C. For a 10:1 ratio, no additional passivation is observed beyond that provided by the APT, which means the MCH did not successfully block the remaining free gold sites. A significant increase in passivation is observed at a 1:1 ratio and remains stable with increasing MCH concentrations. A 1:1 ratio (1 µM APT:1 µM MCH) was selected as optimal for effective blocking while ensuring the aptamer remained attached to the surface. It is worth noting that we did not observe complete
passivation after APT/MCH immobilization, probably due to some defects in the SAM layer
and the possible permeation of ions into the short-chain thiol MCH layer [39].

## 3.2 False positive: the aptasensor instability in blank with the modified C-SPE.

After optimizing the MCH/APT/AuNPs/C-SPE platform, the sensor was incubated with TRIS buffer for 20 minutes at room temperature to record the blank signal. Afterward, the electrode was left in contact with ERY solution in TRIS buffer. The electrode was finally rinsed with TRIS buffer to remove any non-specifically adsorbed ERY molecules and the signal was recorded again. Because there is variability between the surface areas of C-SPEs, causing variations in sensor response, the signal was calculated as a percentage of increase or decrease in peak current compared to the blank. The response of one sensor after successive incubations with solutions of increasing ERY concentration was first studied and is shown in Figure 4A. The current intensity corresponding to the oxidation of $K_4[Fe(CN)_6]$ increases, which was initially attributed to the increased permeability of the SAM due to the interaction of the aptamer with ERY. The resulting calibration curve that plots the percentage of increase of the current intensity vs ERY concentration shows a logarithmic increase (Figure 4B). This is a common effect in aptasensors which is generally attributed to the saturation effect of the aptamers by the analyte. The same logarithmic increase is observed on other modified AuNPs/C-SPE after incubation with increasing concentrations of ERY (Figure S4A). However, even when using the percentage in peak increase as a signal, different MCH/APT/AuNPs/C-SPE sensors have a different response to ERY, indicating poor reproducibility among the aptasensors.

The calibration curve was also recorded on 6 independent electrodes after incubations with different ERY concentrations. In this case, no signal increase is observed with increasing ERY concentration (Figure S4 B). All electrodes show an approximately 11% increase in current intensity following incubation with ERY solutions at concentrations ranging from 5 x $10^{-8}$ M to 5 x $10^{-5}$ M for 20 minutes.

For reference purposes, multiple incubations were then performed in a blank TRIS buffer solution on the same electrode. As shown in Figure 4C, the percentage of peak increase correlates with the incubation time (red line). When simulating a calibration curve with the same concentrations of ERY used as the x-axis in Figure 4B, an identical logarithmic response is observed as shown by the dashed blue line. This reference experiment indicates that the ERY signal observed in Figure 4A is a false positive. Most aptasensors studies do not examine the impact of multiple blank incubations on the electrochemical signal, assuming no response occurs without the target. Our findings highlight the critical importance of investigating the signal in blanks to avoid false positive responses.
<figure><img src="image_7.png" /><figcaption>Figure 4. A) DPV characterization in 5 mM K₄[Fe(CN)₆]/K₃[Fe(CN)₆] of one C-SPE aptasensor after successive incubations with increasing ERY concentrations. Calibration curve plotted on an aptasensor that underwent successive incubations with B) ERY and C) TRIS. Percentage of peak vs. incubation time (red line)</figcaption></figure>

Other studies highlighted the effects of surface chemistry on the stability and performance of aptasensors [8,9,12,13]. As our findings indicate an evolution of the electrochemical signal over time, the stability of the developed platform was tested starting with the bare C-SPE and after each functionalization step.

### 3.3 Origin of the modified C-SPE instability.

The stability of the C-SPE electrodes in TRIS buffer (blank) was evaluated following a preconditioning step, which included rinsing the electrodes with ethanol to remove any residues from the fabrication ink. Figure 5A shows the evolution of the peak current corresponding to the oxidation of 5 mM K₄[Fe(CN)₆] in DPV. The peak current is stable after 5 incubations of 20 minutes each in the TRIS buffer. Additionally, the electrodes demonstrate good reproducibility, with the peak current measured across five electrodes being 151 ± 14 µA. The initial supporting platform remains stable in the blank solution, indicating that it did not contribute to the signal changes observed in the MCH/APT/AuNPs/C-SPE electrode. The stability was also tested after AuNPs deposition, using the same redox probe solution. Figure 5B displays the peak current recorded on the AuNPs/C-SPE platform. The peak current increases notably following the gold deposition, reaching a value of 501 ± 21 µA. Subsequent analyses show minimum variation in this peak current, indicating that until this modification step, the electrode was stable.

The next step was to study the stability of the SAM formed solely by the aptamer or MCH, aiming to identify which component contributes to the mixed SAM instability. After the immobilization of 1 µM APT for 5 minutes by MPA, the peak current decreased to 193 ± 22 µA. After successive TRIS incubations, the peak current increases continuously and reaches a maximum after incubation overnight in TRIS, corresponding to the value obtained on the AuNPs/C-SPE surface (Figure 5C). The APT SAM is unstable in solution, with the increased current intensity suggesting that the aptamer detaches from the surface, enhancing electron transfer between the redox probe and the electrode surface. The same experiment was conducted after 1 µM MCH immobilization using the pulse-assisted method. In this case, the current stabilizes early on and remains stable even after the overnight incubation in TRIS. These results show that the instability in the final aptasensor can be attributed to SAM
formation, specifically due to the APT. Because of their large size, negative charge, and susceptibility to humidity, aptamers tend to change conformation and experience extended rearrangement phases before reaching equilibrium on the electrode surface [40]. While back-filling in MCH helps the proper aptamer orientation [41], it did not enhance the stability of the mixed SAM. This suggests that mixed SAM instability may stem from SAM formation.

<figure><img src="image_8.png" /><figcaption>Figure 5. The evolution of the peak current in DPV corresponding to 5 mM K₄ [Fe(CN)₆] /K₃[Fe(CN)₆] in TRIS buffer after successive blank incubations of A) C-SPE, B) AuNPs/C-SPE, C) APT/AuNPs/C-SPE, and D) MCH/AuNPs/C-SPE, error bars corresponding to 5 electrodes.</figcaption></figure>

## 3.4 Stabilization strategies.

### 3.4.1 SAM formation.

The first approach to surface stabilization was the adjustment of the SAM formation conditions. SAM formation is a dynamic process, influenced by factors such as thiol concentration, temperature, incubation time, solvent, and other thiols present in the solution. As seen in Figure 6A, APT immobilization using MPA results in unstable SAMs, with subsequent MCH immobilization failing to stabilize them. To achieve organized SAMs, APT immobilization time was extended as previously proposed [8, 42]. Figure 6B illustrates the stability of SAM formed after overnight aptamer immobilization. The current remains stable after successive blank incubations in the TRIS buffer. As previously shown in Figure 3A, the passivation obtained after overnight APT immobilization by OCV is similar to that with the MPA method. However, stable APT SAM is achieved with the OCV method after 16 hours, unlike the unstable SAM with MPA (Figure 6A). Although the MPA method offers good coverage and quick setup, the OCV method proved preferable for ERY aptamer SAM formation due to its stability. MCH immobilization was subsequently performed for two hours at OCV. Following MCH immobilization, the peak current decreases and stabilizes
after the second blank incubation. An alternative formation strategy involved APT
immobilization by MPA followed by overnight MCH immobilization (Figure S5A). In this
case, the mixed SAM remains unstable after successive TRIS incubations. As APT
rearrangement is the limiting step, extending APT immobilization time is more relevant than
extending MCH immobilization time. Notably, co-immobilization of APT and MCH in a 1:1
ratio overnight yielded a stable signal after two TRIS buffer incubations (Figure S5B).
However, the final passivation is less important compared to the sequential immobilization,
indicating a lower thiol density on the gold surface. These findings emphasize the importance
of extended APT immobilization time to obtain a reproducible blank signal, likely due to
enhanced SAM organization.

Previous studies have shown that coating the surface with MCH, followed by aptamer
immobilization either through thiol exchange or via pinholes in the MCH SAM, leads to a
more uniform aptamer distribution [24]. This approach effectively minimizes the
agglomeration of the APT. In Figure S5C, the order of thiol immobilization was reversed,
with MCH immobilized first, followed by APT, both by MPA. The signal with the MCH
SAM is stable as shown in Figure 5D. APT immobilization results in a stable peak current,
suggesting that initial immobilization of the blocking agent can yield stable SAMs, even if the
MPA method was used to immobilize the APT.

For all four SAM formation strategies, no ERY response is detected after a 20-minute incubation with a 50 µM solution (Figure 6B and S5). Platforms with good blank stability are achieved, but analyte detection performance is not attained. To verify that the immobilized aptamer remained functional and capable of binding its target, we conducted SPR binding assays using increasing concentrations of ERY. The Langmuir binding isotherm is shown in Figure S6. The resulting sensorgrams demonstrate a concentration-dependent response, confirming that the aptamer retains its affinity for the target. The dissociation constant is 115 ± 23 nM, indicating good affinity between the aptamer and ERY, and the maximum response signal (Rmax) is estimated at 85.47 mdeg. This validation is crucial in distinguishing between potential surface chemistry issues and actual target recognition limitations.

Because of the high carbon content, the oxidation peak of the redox probe remains significant
even after APT and MCH immobilization. Therefore, the strong carbon platform signal may
mask changes from ERY complexation and aptamer conformation alteration. Moreover, Au
morphology strongly affects binary SAM distribution, APT density, and packing [28]. For
these reasons, additional tests were conducted on a crystalline gold surface with [111]
orientation to improve aptasensor analytical performance. Moreover, the crystalline gold
surface should promote more compact SAMs, enhancing stability and sensitivity to aptamer
conformational changes.
<figure><img src="image_9.png" /><figcaption>Figure 6. The values of the peak currents obtained in DPV in 5 mM K₄[Fe(CN)₆]/K₃[Fe(CN)₆] after functionalization and blank incubations of SAMs formed on AuNPs/C-SPE platform by the immobilization of A) APT and MCH by MPA, B) APT overnight and MCH 2 h.</figcaption></figure>

### 3.4.2 Gold substrate.

A comparison between the blank stability of the AuNPs/C-SPE platform with a gold surface (AuE) was conducted. MCH and APT were first immobilized independently overnight by OCP and by MPA from 1 µM solutions under the same conditions as above. The analysis in the redox probe solution shows passivation after MCH immobilization for both methods. This passivation stabilizes at values of 87 % and 62 % compared to the bare AuE for OCV overnight and MPA, respectively, after the first wash in TRIS buffer, probably due to the desorption of molecules from the surface (Figure 7A). Although ideal conditions were met for a compact MCH SAM, the layer does not cover the entire surface of the electrode, leaving space for the redox probe to exchange electrons with the surface. These results are consistent with those reported by Li *et al*. who showed that MCH is a poor blocking agent, due to its low capacity to stabilize through intermolecular interactions [43]. It is worth noting that even though the passivation by MPA is less significant than by OCV, the MCH SAM is stable after several incubations in TRIS buffer. The incubation with ERY 50 µM on the MCH/AuE platforms does not generate any change in signal, proving that ERY does not adsorb on the surface and cannot be detected without the presence of APT (Figure 7D).

In the case of APT, both immobilization methods led to similar passivation with 72 % for the electrodes (Figure 3A). However, a difference between the two methods regarding blank stability was observed. The SAM formed by the MPA method is less stable after several blank incubations, leading to a continuous increase in peak current, while the SAM prepared by the OCV method is stable during 5 incubations in the blank. Both observations correlate with the previous results obtained on the C-SPE platform even though the signal variation was lower (Figure 5). Although the pulsed method is faster and leads to similar electrode passivation, it results in less stable SAMs compared to those formed using the OCV method. The incubation of APT/AuE generated at OCV overnight with 50 µM ERY results in a slight decrease in current intensity of 1.42 %, indicating that the SAM did not attain an organization suitable for the analyte detection and the presence of APT alone is not enough for ERY detection (Figure 7D).
The mixed SAM was formed by sequentially immobilizing APT and then MCH by MPA and OCP (Figure 7C). Interestingly, the signal observed for both surfaces behaved very similarly, except for greater instability after APT immobilization by MPA. Unlike the C-SPE platform, the blank of the mixed APT-MCH SAM is stable even when APT is immobilized by MPA. This difference in behavior can be explained by the [111] crystallinity of the gold surface, which is known to improve the stability of SAMs [28, 38]. As shown in Figure S3 (red), the gold layer deposited on silicon exhibits a high crystallinity and an almost perfect [111] orientation, with the only visible narrow peaks being (111) and (222) in the 30 – 120° 2θ range.

Interestingly, incubation with a 50 µM ERY solution on these well-stabilized SAM electrodes led to passivation of 9.53 % on the platform generated by MPA and 15.60 % on the platform generated by OCV (Figure 7D). Considering the high stability of the blank signal and the obvious change upon ERY incubation, the drop in peak current corresponds to the APT conformational change after interaction with ERY.

<figure><img src="image_10.png" /><figcaption>Figure 7. The values of the peak currents obtained in DPV in 5 mM K₄[Fe(CN)₆]/K₃[Fe(CN)₆] after blank incubations of SAMs formed by MPA and OCV on AuE platform by the immobilization of A) MCH 1 µM, B) APT 1 µM, C) APT 1 µM and MCH 1 µM, D) The signal of 50 µM ERY solution after 20 minutes of incubation on the MCH SAM, APT SAM, and APT/MCH SAM.</figcaption></figure>

The decrease in current intensity is the most commonly reported in the literature in the case of label-free aptasensors [44] and indicates successful ERY detection.

To validate the aptasensor response following incubation with ERY multiple concentrations were tested. Figure 8 shows a linear increase in passivation percentage as a function of ERY concentration. Since the tested concentrations do not saturate the aptasensor, the response remains within the linear range, eliminating the need for logarithmic correction. The signal
corresponding to ERY, which is the passivation percentage, increased linearly within the concentration range of 1 × 10⁻⁶ to 2 × 10⁻⁴ M. The limit of detection (LOD) (3 × the standard deviation of five blank determinations) was determined from equation 3, where St is the signal given by the analyte at LOD, Sb is the average of five field blanks and σ the variability in the field blank. An LOD of 3.2 × 10⁻⁷ M was calculated. Although this detection limit does not meet the stringent requirements for environmental detection of ERY (1.3 × 10⁻⁹ M) [29], it is notable that, through optimized SAM formation and platform selection, we achieved ERY detection via aptasensing. While surpassing the initial scope of our study, a lower detection limit could potentially be reached by increasing the electrode surface area while maintaining the same SAM construction protocol.

$$St - Sb \geq 3\sigma (3)$$

<figure><img src="image_11.png" /><figcaption>Figure 8. The response of the MCH/APT/AuE platform after successive incubations with increasing ERY concentrations obtained on 6 independent electrodes.</figcaption></figure>

## 4. Conclusion.

In this study, an electrochemical aptasensor specific for ERY detection was developed by the self-assembly of the aptamer on gold-based electrodes. To the best of our knowledge, this study marks the first attempt to use an aptamer for the electrochemical detection of erythromycin and demonstrates the ability of the MCH/APT/Au electrode to detect ERY.

The first construction approach used a C-SPE coated with AuNPs and the immobilization of the APT and the blocking agent, MCH, was performed by a pulse-assisted method. While the SAM construction was quick and efficient, the resulting MCH/APT/AuNPs/C-SPE was unstable upon successive blank incubations. This led to unstable signal readings and a false positive result after the incubation with ERY. The mixed SAM instability was due to surface rearrangement generated by the aptamer on the electrode surface. The stability of the mixed SAM was increased using different SAM formation strategies, with prolonged aptamer and MCH immobilization times yielding the most satisfactory result. A stable response in the blank was observed after the immobilization of the APT overnight and the MCH for two
hours at OCV, emphasizing the importance of prolonged times in forming organized SAMs, especially for macromolecules such as aptamers. However, the signal of ERY remained insignificant on this platform.

The second approach used a [111] gold electrode as supporting material on which the APT and MCH were immobilized by OCV. The mixed SAM obtained on the gold electrode manifested good stability after blank incubations. After the incubation with ERY, the peak current in DPV corresponding to the redox probe decreased, indicating a change in aptamer conformation after interaction with the target. A calibration curve was obtained with a linear range from $1 \times 10^{-6}$ M to $2 \times 10^{-4}$ M and an LOD of $3.2 \times 10^{-7}$ M. This work underscores the critical importance of both the intermediate and final stability of the sensor. Although the influence of blanks on aptasensor response is rarely addressed in the literature, it is essential to ensure reliable target detection and to avoid false positive signals. This work provides insight into the stability of MCH/APT systems.

**Funding:** This work received financial support under the EUR LUMOMAT project and the Investments for the Future program ANR-18-EURE-0012 Nanosens.

**Acknowledgments:** Teodora Lupoi thanks the program "Bourses du Gouvernement français France Excellence" for her scholarship. We thank Origalys ElectroChem for lending us the impedance module used for surface characterization measurements.

## References

[1] J. Pan, W. Xu, W. Li, S. Chen, Y. Dai, S. Yu, Q. Zhou, F. Xia, Electrochemical aptamer-based sensors with tunable detection range, Anal. Chem., 95 (2023) 420-432.

[2] C.R. Rousseau, H. Kumakli, R.J. White, Perspective—Assessing Electrochemical, Aptamer-Based Sensors for Dynamic Monitoring of Cellular Signaling, ECS Sensors Plus, 2 (2023) 042401.

[3] L.F. Yang, M. Ling, N. Kacherovsky, S.H. Pun, Aptamers 101: aptamer discovery and in vitro applications in biosensors and separations, Chem. Sci., 14 (2023) 4961-4978.

[4] K.K. Leung, A.M. Downs, G. Ortega, M. Kurnik, K.W. Plaxco, Elucidating the mechanisms underlying the signal drift of electrochemical aptamer-based sensors in whole blood, ACS Sens., 6 (2021) 3340-3347.

[5] N. Fontaine, P. Dauphin-Ducharme, Confounding Effects on the Response of Electrochemical Aptamer-Based Biosensors, Curr. Opin. Electrochem., (2023) 101361.

[6] A. Bogomolova, E. Komarova, K. Reber, T. Gerasimov, O. Yavuz, S. Bhatt, M. Aldissi, Challenges of electrochemical impedance spectroscopy in protein biosensing, Anal. Chem., 81 (2009) 3944-3949.

[7] W.L. Whitehouse, L.H. Lo, A.B. Kinghorn, S.C. Shiu, J.A. Tanner, Structure-Switching Electrochemical Aptasensor for Rapid, Reagentless, and Single-Step Nanomolar Detection of C-Reactive Protein, ACS Appl. Bio Mater., 7 (2024) 3721-3730.

[8] X. Xu, A. Makaraviciute, S. Kumar, C. Wen, M. Sjödin, E. Abdurakhmanov, U.H. Danielson, L. Nyholm, Z. Zhang, Structural changes of mercaptohexanol self-assembled monolayers on gold and their influence on impedimetric aptamer sensors, Anal. Chem., 91 (2019) 14697-14704.

[9] J.J. Gooding, The Importance of the Stability of Surface Chemistry in Sensors, ACS Sens., 9 (2024) 2203-2204.
[10] R. Levicky, T.M. Herne, M.J. Tarlov, S.K. Satija, Using self-assembly to control the structure of DNA monolayers on gold: a neutron reflectivity study, J. Am. Chem. Soc., 120 (1998) 9787-9792.

[11] T.M. Herne, M.J. Tarlov, Characterization of DNA probes immobilized on gold surfaces, J. Am. Chem. Soc., 119 (1997) 8916-8920.

[12] M.C. Brothers, D. Moore, M. St. Lawrence, J. Harris, R.M. Joseph, E. Ratcliff, O.N. Ruiz, N. Glavin, S.S. Kim, Impact of self-assembled monolayer design and electrochemical factors on impedance-based biosensing, Sensors, 20 (2020) 2246.

[13] F. Ricci, N. Zari, F. Caprio, S. Recine, A. Amine, D. Moscone, G. Palleschi, K.W. Plaxco, Surface chemistry effects on the performance of an electrochemical DNA sensor, Bioelectrochemistry, 76 (2009) 208-213.

[14] S. Li, Y. Wang, Z. Zhang, Y. Wang, H. Li, F. Xia, Exploring end-group effect of alkanethiol self-assembled monolayers on electrochemical aptamer-based sensors in biological fluids, Anal. Chem., 93 (2021) 5849-5855.

[15] H. Li, P. Dauphin- Ducharme, N. Arroyo- Currás, C.H. Tran, P.A. Vieira, S. Li, C. Shin, J. Somerson, T.E. Kippin, K.W. Plaxco, A biomimetic phosphatidylcholine- terminated monolayer greatly improves the in vivo performance of electrochemical aptamer- based sensors, Angew. Chem. Int. Ed., 56 (2017) 7492-7495.

[16] A. Shaver, N. Arroyo-Currás, The challenge of long-term stability for nucleic acid-based electrochemical sensors, Curr. Opin. Electrochem., 32 (2022) 100902.

[17] A. Shaver, S.D. Curtis, N. Arroyo-Curras, Alkanethiol monolayer end groups affect the long-term operational stability and signaling of electrochemical, aptamer-based sensors in biological fluids, ACS Appl. Mater. Interfaces, 12 (2020) 11214-11223.

[18] Z. Zhang, Y. Wang, Z. Mei, Y. Wang, H. Li, S. Li, F. Xia, Incorporating hydrophobic moieties into self-assembled monolayers to enable electrochemical aptamer-based sensors deployed directly in a complex matrix, ACS Sens., 7 (2022) 2615-2624.

[19] L.A. Olivan, K. Hand, R.J. White, Utilization of Spontaneous Alkyne-Gold Self-Assembly Chemistry as an Alternative Method for Fabricating Electrochemical Aptamer-Based Sensors, Langmuir, (2024).

[20] M.A. Pellitero, S.D. Curtis, N. Arroyo-Currás, Interrogation of electrochemical aptamer-based sensors via peak-to-peak separation in cyclic voltammetry improves the temporal stability and batch-to-batch variability in biological fluids, ACS Sens., 6 (2021) 1199-1207.

[21] T. Ma, I. Martens, D. Bizzotto, Thermal Stability of Thiolated DNA SAMs in Buffer: Revealing the Influence of Surface Crystallography and DNA Coverage via In Situ Combinatorial Surface Analysis, Langmuir, 36 (2020) 14495-14506.

[22] L.S.J. Ho, J.L. Limson, R. Fogel, Certain methods of electrode pretreatment create misleading responses in impedimetric aptamer biosensors, ACS Omega, 4 (2019) 5839-5847.

[23] A.W. Peterson, R.J. Heaton, R.M. Georgiadis, The effect of surface probe density on DNA hybridization, Nucleic Acids Res., 29 (2001) 5163-5168.

[24] J.N. Murphy, A.K. Cheng, H.-Z. Yu, D. Bizzotto, On the nature of DNA self-assembled monolayers on Au: measuring surface heterogeneity with electrochemical in situ fluorescence microscopy, J. Am. Chem. Soc., 131 (2009) 4042-4050.

[25] Y. Wu, J. Shi, T.E. Kippin, K.W. Plaxco, Codeposition Enhances the Performance of Electrochemical Aptamer-Based Sensors, Langmuir, 40 (2024) 8703-8710.

[26] F. Ma, R.B. Lennox, Potential-assisted deposition of alkanethiols on Au: Controlled preparation of single-and mixed-component SAMs, Langmuir, 16 (2000) 6188-6190.

[27] D. Jambrec, M. Gebala, F. La Mantia, W. Schuhmann, Potential- Assisted DNA immobilization as a prerequisite for fast and controlled formation of DNA monolayers, Angew. Chem. Int. Ed., 54 (2015) 15064-15068.
[28] K.K. Leung, A.D. Gaxiola, H.-Z. Yu, D. Bizzotto, Tailoring the DNA SAM surface density on different surface crystallographic features using potential assisted thiol exchange, Electrochim. Acta, 261 (2018) 188-197.

[29] B.H. Schafhauser, L.A. Kristofco, C.M.R. de Oliveira, B.W. Brooks, Global review and analysis of erythromycin in the environment: occurrence, bioaccumulation and antibiotic resistance hazards, Environ. Pollut., 238 (2018) 440-451.

[30] Y. Du, D. Liu, M. Wang, F. Guo, J.S. Lin, Preparation of DNA aptamer and development of lateral flow aptasensor combining recombinase polymerase amplification for detection of erythromycin, Biosens. Bioelectron., 181 (2021) 113157.

[31] M. Tertiş, B. Ciui, M. Suciu, R. Săndulescu, C. Cristea, Label-free electrochemical aptasensor based on gold and polypyrrole nanoparticles for interleukin 6 detection, Electrochim. Acta, 258 (2017) 1208-1218.

[32] D.W. Ussery, DNA denaturation, Encyclopedia of genetics, (2001) 550-553.

[33] C. Singhal, S. Gupta, J. Dhingra, S. Pandey, S. Chatterjee, R. Bargakshatriya, D.K. Avasthi, S. Pramanik, S. Chaudhuri, Label-free electrochemical aptasensor for detection of Acinetobacter baumannii: Unveiling the kinetic behavior of reduced graphene oxide v/s graphene oxide, Electrochim. Acta, 492 (2024) 144240.

[34] J. Rodríguez-Carvajal, Recent advances in magnetic structure determination by neutron powder diffraction, Phys. Rev. B Condens. Matter, 192 (1993) 55-69.

[35] T. Hezard, K. Fajerwerg, D. Evrard, V. Collière, P. Behra, P. Gros, Gold nanoparticles electrodeposited on glassy carbon using cyclic voltammetry: Application to Hg (II) trace analysis, J. Electroanal. Chem., 664 (2012) 46-52.

[36] O. Thipmanee, S. Samanman, S. Sankoh, A. Numnuam, W. Limbut, P. Kanatharana, T. Vilaivan, P. Thavarungkul, Label-free capacitive DNA sensor using immobilized pyrrolidinyl PNA probe: Effect of the length and terminating head group of the blocking thiols, Biosens. Bioelectron., 38 (2012) 430-435.

[37] Y. Song, Y. Liu, M. Yang, B. Zhang, Z. Li, Diluting thiol-derivatized oligonucleotide monolayers on Au (1 1 1) by mercaptohexanol replacement reaction, Appl. Surf. Sci., 252 (2006) 5693-5699.

[38] J.C. Love, L.A. Estroff, J.K. Kriebel, R.G. Nuzzo, G.M. Whitesides, Self-assembled monolayers of thiolates on metals as a form of nanotechnology, Chem. Rev., 105 (2005) 1103-1170.

[39] Z. Li, T. Niu, Z. Zhang, S. Bi, Potential control characteristics of short-chain thiols of thioctic acid and mercaptohexanol self-assembled on gold, Electrochim. Acta, 55 (2010) 6907-6916.

[40] K.B. Frøhling, T.S. Alstrøm, M. Bache, M.S. Schmidt, M.N. Schmidt, J. Larsen, M.H. Jakobsen, A. Boisen, Surface-enhanced Raman spectroscopic study of DNA and 6-mercapto-1-hexanol interactions using large area mapping, Vib. Spectrosc, 86 (2016) 331-336.

[41] M. Gębala, W. Schuhmann, Controlled orientation of DNA in a binary SAM as a key for the successful determination of DNA hybridization by means of electrochemical impedance spectroscopy, ChemPhysChem, 11 (2010) 2887-2895.

[42] D.Y. Petrovykh, H. Kimura-Suda, L.J. Whitman, M.J. Tarlov, Quantitative analysis and characterization of DNA immobilized on gold, J. Am. Chem. Soc., 125 (2003) 5219-5226.

[43] Z. Li, T. Niu, Z. Zhang, G. Feng, S. Bi, Studies on the effect of solvents on self-assembly of thioctic acid and Mercaptohexanol on gold, Thin Solid Films, 519 (2011) 4225-4233.

[44] F.V. Oberhaus, D. Frense, D. Beckmann, Immobilization techniques for aptamers on gold electrodes for the electrochemical detection of proteins: a review, Biosensors, 10 (2020) 45.