Skip to main content
NCBI home page
Scientific Reports logoLink to Scientific Reports
. 2025 Mar 14;15:8859. doi: 10.1038/s41598-025-93025-1

A novel strategy for controllable electrofabrication of molecularly imprinted polymer biosensors utilizing embedded Prussian blue nanoparticles

Bahareh Babamiri 1,#, Mohammadreza Farrokhnia 1,#, Mehdi Mohammadi 2,✉, Amir Sanati Nezhad 1,3,✉
PMCID: PMC11909151  PMID: 40087363

Abstract

The reproducibility of ultrasensitive biosensors is vital for clinical research, scalable manufacturing, commercialization, and reliable clinical decision-making, as batch-to-batch variations introduce significant uncertainty. However, most biosensors lack robust quality control (QC) measures. This study introduces an innovative QC strategy to produce highly reproducible molecularly imprinted polymer (MIP) biosensors by leveraging real-time data from the electrofabrication process. Prussian Blue nanoparticles (PB NPs) embedded within the MIP structure enable precise monitoring of surface properties, conductivity, MIP film thickness, and template extraction efficiency. The QC strategy utilizes variations in the current intensity of PB NPs during fabrication to implement real-time, non-destructive QC protocols at critical fabrication stages, minimizing measurement variability and ensuring consistency. This approach was validated by fabricating MIP biosensors for detecting agmatine metabolite and glial fibrillary acidic protein (GFAP) in phosphate-buffered saline (PBS). The QC strategy reduced relative standard deviation (RSD) by 79% for agmatine (RSD = 2.05% QC, RSD = 9.68% control) and 87% for GFAP (RSD = 1.44% QC, RSD = 11.67% control). Moreover, quality-controlled biosensors achieved success rates of 45% for agmatine and 36% for GFAP detection, significantly outperforming bare screen-printed electrodes. This work marks a significant advancement in biosensor development by integrating robust QC protocols directly into the fabrication process. By embedding PB NPs and monitoring electrochemical signals in real-time, this strategy delivers an unprecedented level of reproducibility, scalability, and reliability for MIP biosensors, addressing critical challenges in point-of-care diagnostics and commercial applications.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-93025-1.

Keywords: Electrochemical biosensor, Quality control, Electrodeposition, Molecularly imprinted polymers, Prussian blue nanoparticles

Subject terms: Bioanalytical chemistry, Sensors

Introduction

The rapid and precise detection of biomolecules in bodily fluids—such as metabolites, nucleic acids, proteins, and pathogens-is fundamental for early disease diagnosis and health monitoring1−4. To address this need, the development of biosensors that are highly sensitive, selective, reliable, cost-effective, and reproducible is critical for facilitating timely medical evaluations and effective disease management, particularly in point-of-care settings where trained personnel may not be available. Electrochemical biosensors, with their exceptional sensitivity, specificity, portability, and ease of use, have emerged as indispensable tools in the design of advanced chemical and biological sensors5–11. Reproducibility is a cornerstone for ensuring reliable biosensor performance and experimental validity, particularly when adhering to regulatory standards12–14. ISO 13485 (Medical Devices - Quality Management Systems) provides a comprehensive framework for quality management during the development of medical devices, while Good Manufacturing Practices (GMP) ensure consistent production processes and product quality. Furthermore, compliance with FDA regulations and CE marking requirements guarantees that biosensors meet rigorous safety, efficacy, and reliability criteria, making them suitable for both clinical applications and commercialization.

Electrochemical biosensors can be miniaturized, batch-fabricated, and integrated into portable, implantable, and wearable devices for point-of-care diagnosis and prognosis15–17. Their use in miniaturized sensing and diagnostic platforms has led to the development of various screen-printed electrodes18,19. However, batch-to-batch variations in ink properties and substrate characteristics cause uncertainties in electrode behavior, affecting conductivity, resistance, capacitance, and electroactive surface area20–22. Additionally, shelf stability and functionality over time are crucial for POC applications14,23. Proper storage procedures are also essential to maintain stable electrochemical performance. Variations in fabrication processes and storage can impact the reliability of biosensors, necessitating comprehensive monitoring of each manufacturing and storage stage to ensure consistent performance.

Implementing quality control (QC) and quality management system (QMS) strategies during biosensor manufacturing and storage can create uniform, reproducible electrode surfaces and reliable biosensors. A QMS establishes a comprehensive framework of processes, procedures, and documentation for all fabrication stages, ensuring reproducibility and reliability. Their QC involves systematic monitoring and evaluation to identify and correct deviations, using techniques like electrode characterization, raw material testing, and process validation. Covering the entire biosensor lifecycle, a QMS defines protocols for design control, risk management, production planning, and validation to minimize variations and achieve consistent, high-quality biosensors. This system ensures that each fabrication step is documented, controlled, and traceable, leading to reliable, reproducible results24–28.

Most biosensing technologies employ biological recognition elements, such as enzymes, antibodies, nucleic acids, and aptamers, as capture probes, offering high efficacy and selective affinity to target biomolecules. However, these biorecognition elements face issues such as long-term stability, high costs, single-use nature, and degradation in biological media29,30. The development of MIPs as efficient biomimetic receptors has significantly addressed longstanding challenges in sensor technology. MIPs offer exceptional chemical and thermal stability, a long shelf life, ease of fabrication, cost-effectiveness, and high selectivity for target molecules by mimicking natural molecular recognition processes. Additionally, their reusability and resistance to enzymatic degradation make MIPs ideal for long-term, sustainable, and economical applications31–35. However, advancing electrochemical MIP biosensors presents several challenges. These include the careful selection of suitable monomers and cross-linkers, precise control over the thickness of the polymeric layer, effective template removal without compromising recognition sites, and ensuring the reproducibility of MIP films during large-scale production36–39. Overcoming these obstacles is critical for fully leveraging the potential of MIP-based biosensors in diverse applications.

The electro-fabrication procedure, a critical step in biosensor development, demands stringent QC measures to ensure reproducibility and reliability. Electrodeposition of embedded redox probes such as Prussian blue (PB) in electrochemical MIP biosensors enables monitoring of the electrical signal at each fabrication step, providing an effective QC measure, and offering a simple and highly controllable method to achieve reproducible MIP biosensors. Prussian blue nanoparticles are recognized as excellent electron mediators due to their ability to undergo reversible redox transitions between ferric and ferrous states. This unique property facilitates efficient electron transfer at the electrode surface. By integrating PB NPs within the molecularly imprinted polymer matrix, the study capitalizes on their current intensity as a real-time indicator for monitoring the electropolymerization and template extraction processes40,41.

Additionally, the electropolymerization of conductive monomers facilitates the direct, in situ fabrication of MIPs on electrode surfaces. This technique offers a rapid and straightforward method to produce electrically conductive polymeric films with controllable thickness, uniform morphology, reproducible outcomes, and real-time polymer growth monitoring, enhancing the precision of MIP-based biosensor development42–46. Such strategic tuning is crucial for modulating charge transfer through the electrode, thereby improving the overall performance of the biosensor.

Variations in ink properties, substrate characteristics, and fabrication methods of biosensors can lead to disparities in electrochemical performance, impacting the accuracy and precision of MIP biosensors47,48. Thus, developing an efficient QC strategy to address these challenges and enhance the reliability and reproducibility of biosensors is crucial. Screening electrodes during the fabrication process is essential to identify and eliminate defects or irregularities that may compromise the overall performance of the biosensors.

This work presents an innovative QC strategy for fabricating MIP biosensors, aimed at achieving repeatable and reliable performance in detecting metabolites and proteins. This strategy includes non-destructive QC steps based on the electro-fabrication processes. Key steps involve the electrodeposition of a redox probe, electropolymerization of the MIP film using pyrrole as a functional monomer, and extraction of the template molecule (Fig. 1). Two extraction approaches were utilized: electro-cleaning (Approach A) and solvent extraction (Approach B). Each step in the MIP sensor fabrication process was monitored using cyclic voltammetry (CV), square wave voltammetry (SWV), and electrochemical impedance spectroscopy (EIS). This approach is crucial for developing reproducible MIP biosensors, aiming to meet the needs of ISO 13485 certification for large-scale manufacturing and commercialization. Adhering to this systematic QC strategy also ensures time and cost savings, facilitating advancements in biosensing technologies for biomedical applications.

Fig. 1.

Fig. 1

Open in a new tab

The preparation process of metabolite and protein molecularly imprinted polymer (MIP) biosensors, incorporating various quality control measures for their reproducible production [PB: Prussian blue, PBS: phosphate-buffered saline, EIS: electrochemical impedance spectroscopy, SWV: square wave voltammetry].

Results and discussion

Characterization of embedded Prussian blue nanoparticles

The developed QC strategy monitors variations in the current intensity of Prussian blue (PB) nanoparticles (NPs) during the electrofabrication of MIP biosensors. PB was selected as the internal redox probe for its excellent electrochemical properties, including a reversible redox reaction and rapid electron transfer. The stability of PB NPs is critical for ensuring the reproducibility of MIP biosensors, directly influencing the effectiveness of the QC strategy. Figure 2a shows stable oxidation and reduction peaks of PB NPs over 60 continuous CV scans, underscoring their potential for monitoring and selecting reproducible electrodes.

Fig. 2.

Fig. 2

Open in a new tab

Characterization of Prussian blue nanoparticles (PB NPs) embedded into electrodes. (a) Consecutive cyclic voltammetry (CV) scans of the modified electrode with energy-dispersive spectroscopy PB NPs in 0.1 M potassium chloride (KCl), (b) Field emission scanning electron microscopy (FE-SEM) images of the electrodeposited PB NPs, (c) Gauss fitting of particle size distribution of PB NPs, (d) Energy-dispersive spectroscopy (EDS) mapping of the PB NPs, the element distribution diagram of Fe, C, N.

The surface properties of PB NPs were analyzed using field emission scanning electron microscopy (FE-SEM) and energy-dispersive spectroscopy (EDS). The analysis confirms that PB NPs with cubic structures are uniformly electrodeposited onto the electrode surface, exhibiting a consistent size distribution ranging from 80 to 200 nm (Fig. 2b, c). Figure 2c illustrates the size distribution of PB NPs as a histogram (blue bars) alongside a fitted bell curve (red line) representing the normalized frequency of particle sizes. This data was derived from the FE-SEM images and analyzed using ImageJ software. The average nanoparticle size was calculated to be 108.88 nm, with a standard deviation of 28.69 nm. The strong alignment between the bell curve and the histogram demonstrates that the particle size distribution closely follows a normal distribution. These findings highlight a predominantly uniform size distribution of PB NPs on the electrode surface, ensuring consistent deposition and contributing to the reliability of the biosensor’s performance.

EDS elemental mapping (Fig. 2d) further confirms the even distribution of Fe, C, and N within the PB NP structure. These results validate the successful electrodeposition of PB NPs as an embedded redox probe on the electrode surface.

QC strategy for creating reproducible MIP biosensors

The inherent variability in manufacturing processes and the challenges associated with preparing reliable and reproducible MIP biosensors for biomolecule recognition highlight the critical need for a straightforward and controllable MIP fabrication process complemented by stringent QC measures. To address these issues, several strategies have been proposed, including the use of advanced characterization techniques, optimization of electrode printing parameters, and the development of standardized protocols for electrode evaluation49–51. By concentrating on key parameters that influence electrode performance, we have established a systematic and efficient QC approach that effectively addresses these challenges.

In this strategy, we eliminated redundant destructive steps by focusing solely on the results produced automatically during the electro-fabrication process. The manufacturing protocol for MIP biosensors incorporates this new strategy (Fig. 3). The flowchart details the production process of MIP-based biosensors, emphasizing four non-destructive QC steps: QC1 (Visual test and storage conditions of bare electrodes), QC2 (Electrodeposition step), QC3 (Electropolymerization step), and QC4 (Extraction step). Each step’s threshold is technically determined based on the MIP biosensor design, materials used in the platform structure, and the results of each electrode group.

Fig. 3.

Fig. 3

Open in a new tab

Flowchart detailing the creation of reproducible molecularly imprinted polymer (MIP)-based biosensors, highlighting four non-destructive quality control steps (QC1 to QC4).

In the initial step, bare electrodes received from the producer underwent visual inspection and assessment of storage conditions (QC1) from the time of manufacturing to receipt for biosensor production. Electrodes that passed QC1 moved to the PB electrodeposition stage. During this stage, the anodic current peak obtained from the CV method during PB NP electrodeposition was used to evaluate electrochemical performance (QC2). Each electrode was assessed based on a predetermined current intensity peak threshold, allowing for less than 5% deviation. Electrodes that did not meet this criterion were discarded. Qualified electrodes advanced to the electropolymerization stage (QC3), where variations in the current intensity of PB NPs during electropolymerization served as a criterion for further selection. Following the extraction of template molecules, QC4 was implemented to finalize the selection of reproducible MIP biosensors based on their electrochemical performance.

Electrochemical profiling during PB electrodeposition, MIP layer formation, and template extraction

The development of reliable and reproducible electrochemical MIP-based biosensors demands uniformity in electrode electrochemical characteristics and surface morphology. Ensuring precise control over the fabrication of the MIP film and the efficient extraction of template molecules is critical for producing consistent and high-performance polymer films. Incorporating PB NPs as embedded redox probes introduces a powerful strategy to enhance reproducibility by selecting electrodes based on their surface properties and electrochemical behavior. In this methodology, a defined peak current intensity threshold for electrodeposited PB NPs on the electrode surface was established for each stage of the MIP-based biosensor fabrication. During the electro-fabrication process, electrodes failing to meet this threshold were systematically eliminated, ensuring uniformity across batches. Furthermore, analyzing variations in the current intensity of PB NPs during the electropolymerization of pyrrole and subsequent template extraction provided an effective real-time QC mechanism. This approach not only ensures consistent electrode performance but also addresses key challenges in the fabrication process, significantly enhancing the reliability and reproducibility of MIP-based biosensors.

To demonstrate the applicability of the developed QC strategy for achieving reproducible biosensors, we used two different batches of electrodes. Freshly manufactured electrodes were used for the agmatine biosensor fabrication, while electrodes produced three months earlier and stored in non-optimal conditions were used for the GFAP biosensor. We utilized two different approaches for the extraction of template molecules: Approach A (Electro-cleaning) for the agmatine MIP biosensor and Approach B (Solvent extraction) for the GFAP MIP biosensor.

The change in the current intensity of PB NPs during the fabrication process of the agmatine MIP biosensor (Electro-cleaning approach) is shown in Fig. 4a-c. The growth process of PB NPs on the electrode’s surface was apparent where the redox peak gradually increased, and the peak potential remained relatively constant with the increase in the scan number (Fig. 4a). The electropolymerization method was applied to create a uniform MIP film of Polypyrrole (Ppy) on the modified electrode’s surface. Figure 4b illustrates CV curves during the electropolymerization process of pyrrole in the presence of agmatine and 3-aminophenyl boronic acid on the surface of the electrodeposited electrode, with PB NPs used as the embedded redox probe. The curves show the oxidation peaks of PB NPs at about + 0.2 V. Throughout the fabrication of MIP film, the current intensity of PB NPs decreased with the increasing number of cycles, almost disappearing by the 10th cycle, confirming the formation of a reliable imprinted polymer film on the surface of the electrode. The complementary cavities to agmatine were obtained upon the extraction of template molecules through the over-oxidation of Ppy (Electro-cleaning approach). The opened imprinting cavities act as channels for the electron transfer of PB NPs. Increasing the current intensity of PB NPs with the increasing number of cycles indicates the successful extraction of almost template agmatine from the MIP structure (Fig. 4c). A similar pattern is observed for the change in the current intensity of PB NPs during electrofabrication (electrodeposition and electropolymerization) of the GFAP MIP biosensor using the solvent extraction approach (Fig. 4d, e). Therefore, an efficient approach for QC evaluation of electrodes was established through precise monitoring of the changes in the current intensity of the PB NPs, serving as an internal embedded redox probe during the fabrication procedure. This systematic approach assesses electrode performance throughout all its fabrication steps, significantly enhancing the reliability and reproducibility of the biosensors.

Fig. 4.

Fig. 4

Open in a new tab

Electrochemical profiling during PB electrodeposition, MIP layer formation, and template extraction. (a) CV representing the electrodeposition of the PB NPs for the agmatine biosensor, (b) CV representing the MIP formation on the surface of the modified electrode, (c) CV during the extraction of agmatine from the formed Ppy matrix in 500 mM hydrochloric acid (HCl) with a scan rate of 50 mV s−1, (d) CV representing the electrodeposition of the PB NPs for glial fibrillary acidic protein (GFAP) biosensor, (e) CV representing the MIP formation on the surface of the modified electrode for GFAP biosensor.

Surface characterization of MIP biosensors

The surface morphology of the electropolymerized MIP films was analyzed using SEM. SEM images (Figures S1a, b) revealed that the agmatine and GFAP MIP biosensors feature porous and uniform MIP films with a high-density distribution of imprinted sites. In contrast, the NIP electrodes exhibited significantly lower porosity with unevenly distributed cavities (Figures S1c, d). Furthermore, AFM was employed to examine the surface topography, homogeneity, and surface roughness of the MIP films. AFM images (Figures S1e, f) demonstrated that the agmatine and GFAP MIP biosensors possess a consistent and uniform surface topology. The observed homogeneity, uniform surface roughness, and high-density distribution of porous structures collectively confirm the successful electro-fabrication of the MIP biosensors, ensuring their suitability for reliable analytical applications.

Evaluating the QC strategy for producing MIP biosensors

To assess the effectiveness of our fabrication protocol, we initiated the fabrication of MIP biosensors using two different batches of electrodes: 36 freshly manufactured screen-printed electrodes for the agmatine MIP biosensor, and 36 electrodes produced three months earlier for the GFAP MIP biosensor. Once all electrodes passed QC1 (visual inspection and storage condition quality control), we conducted the main QC step following the electrodeposition of the redox probe (QC2). This step was based on predetermined current intensity (anodic peak current (ipa) corresponds to the PB NPs electrodeposition step at potential + 0.2 V relates to the last cycle (#60)) thresholds: 95 mA for agmatine biosensors and 52 mA for GFAP biosensors. The non-optimal storage conditions of bare electrodes resulted in reduced electrochemical performance, leading to lower current intensity in GFAP biosensors compared to agmatine biosensors.

A total of 28 electrodes from the agmatine biosensors and 19 from the GFAP biosensors successfully passed the QC2 electrodeposition step. The electrochemical results for this step are depicted in Figs. 5a and 6a. The minimum anodic peak current (ipa) corresponds to the first cycle of the PB NP electrodeposition, while the maximum ipa is recorded at the final cycle (#60). Throughout the electrodeposition process, the peak current intensity gradually increases, reaching its maximum at the end of the process, which serves as the key QC parameter. While the initial peak current demonstrates relative consistency across different bare electrodes, significant deviations emerge after the 60th cycle of electrodeposition. These variations are likely caused by differences in surface morphology or the electroactive surface area of the electrodes. This finding highlights that relying solely on the first electrochemical peak for evaluation is insufficient for ensuring electrode reproducibility, a method traditionally used for commercial screen-printed electrodes52. By implementing a QC strategy that monitors multiple cycles, a more robust and accurate assessment of the electrochemical behavior of electrodes can be achieved. This comprehensive approach enhances the reliability and consistency of the fabricated biosensors, addressing critical challenges in electrode reproducibility and performance.

Fig. 5.

Fig. 5

Open in a new tab

Anodic peak current results from cyclic voltammetry for (a) electrodeposition of the redox probe (QC2), (b) electropolymerization of the MIP solution (QC3), (c) electro-cleaning of agmatine (QC4), (d) square wave voltammetry voltammograms of 24 electrodes after electro-cleaning of the template molecule, measured in 0.01 M phosphate buffer saline (pH 7.4), and (e) linear calibration curve of three randomly selected MIP biosensors for agmatine detection (ΔI represents the SW peak height).

Fig. 6.

Fig. 6

Open in a new tab

The anodic peak current of cyclic voltammetry results for (a) electrodeposition of redox probe (QC2) and (b) electropolymerization of MIP solution (QC3), (c) EIS of 16 different electrodes after solvent extraction of template molecule measured in 0.01 M PBS (pH 7.4), (d) The corresponding linear calibration curve of three randomized selected MIP biosensors for detection of GFAP.

Figures 5b and 6b show the anodic peaks from the 1st and 10th cycles of the electropolymerization process for preparing agmatine and GFAP MIP biosensors, respectively. According to these results, 24 electrodes for the agmatine biosensor batch and 16 electrodes for the GFAP biosensor batch passed QC3, with predetermined thresholds of 30 mA and 60 mA, respectively. As the thickness of the MIP films increased, the conductivity of the electrode surface decreased after 10 cycles of electropolymerization. Therefore, the anodic peak current in the 10th scan (ipa in the 10th scan) was used as the criterion for advancing the electrodes to the subsequent step. Also, Fig. 5c illustrates the anodic peaks from the 1st and 15th cycles of the electro-cleaning step from agmatine biosensors’ cavities. At this step, we determined a selection threshold of 59 mA for agmatine MIP biosensors. Based on this threshold, 16 electrodes of agmatine biosensors passed the extraction step quality control (QC4).

The final step of the QC strategy is illustrated in Figs. 5d and 6c. After extracting the template molecules as described previously, SWV measurements were performed by dispensing a drop of PBS on the surface of the agmatine MIP biosensors for monitoring the current intensity of electrodeposited PB NPs after extraction of agmatine, while EIS measurements were conducted similarly on the GFAP MIP biosensors. Post-SWV analysis in PBS solution revealed that 16 electrodes maintained consistent current intensity after extraction of agmatine (sub-figure of Fig. 5d). Consequently, approximately 44% of electrodes passed the final quality control step QC4. It is noted that improving the quality of bare electrodes has the potential to increase the pass rate to over 72%. Excluding outlier data within the established range of 1200 Ω to 3700 Ω, 13 electrodes successfully passed QC4. This surface resistance (Rct) range was identified for the GFAP MIP biosensor by evaluating the results across all biosensors. The sub-figure in Fig. 6c indicates that approximately 36% of the electrodes met the criteria of this QC strategy, ensuring the production of reliable and reproducible GFAP MIP biosensors.

Analytical performance and reproducibility of the QC-passed MIP biosensors

A QC strategy was employed to select electrodes with consistent current intensities of PB NPs and stable performance during electrodeposition, electropolymerization, and extraction steps. From the batch passing QC4, three MIP biosensors were randomly chosen. These biosensors were used for the quantitative detection of agmatine and GFAP biomarkers to evaluate batch reproducibility. The analytical performance of these MIP biosensors was examined for agmatine detection in the concentration range of 1 nM to 100 µM and GFAP detection in the range of 0.02 pg mL− 1 to 200 ng mL− 1 in 0.01 M PBS (Figs. 5e and 6d). The calibration curves’ slopes and correlation coefficients of these biosensors were in good agreement, indicating the QC strategy’s effectiveness in producing reliable and repeatable MIP biosensors.

The determined limits of detection (LOD) were 0.1 nM for agmatine and 0.01 fg mL− 1 for GFAP, calculated based on a signal-to-noise ratio of 3, using the standard deviation of the MIP biosensor response and the calibration slope. Compared to previously reported agmatine and GFAP detection methods, the developed MIP biosensors exhibited markedly enhanced sensitivity, an expanded linear detection range, and the lowest detection limits to date (Table S1). These advancements highlight the exceptional performance of the MIP biosensors in achieving ultra-sensitive and precise target detection.

Storage conditions

Ensuring a stable and consistent electrochemical response of bare electrodes over extended periods is crucial for the widespread application of screen-printed electrodes. Due to the high porosity, large specific surface area, and reactive nature of the materials used in fabricating bare screen-printed electrodes, environmental storage conditions can significantly influence their electrochemical characteristics. Environmental factors such as oxygen, moisture, and contaminants interact with the electrode surface, leading to surface fouling and the formation of insulating layers impairing electrochemical performance53,54. Consequently, the electrochemical response of electrodes can decline when electrodes from the same batch are stored for extended periods. The changes in the electrochemical behavior of bare electrodes are highly dependent on storage conditions. Therefore, optimizing storage conditions and implementing QC methods during the storage of bare electrodes are essential for achieving reproducible biosensors in mass production.

To address this challenge, we investigated storage conditions for the bare electrodes and optimized the storage factors that affect the efficiency and electrochemical behavior of bare electrodes during long-term storage. EIS, as a non-destructive method, is highly sensitive to changes in surface properties, including species adsorption, surface roughness changes, and insulating layer formation. Therefore, monitoring the charge transfer resistance (Rct) is an efficient approach to evaluate the stability and degradation of graphene intermixed poly(3,4-ethylenedioxythiophene) polystyrene sulfonate (PEDOT: PSS) carbon (GiPEC) screen-printed electrodes over time. Electrochemical impedance spectroscopy (EIS), a non-destructive and highly sensitive method, was employed to monitor changes in surface properties, including species adsorption, surface roughness alterations, and insulating layer formation. The charge transfer resistance (Rct) was recorded as a key parameter for evaluating the stability and degradation of GiPEC electrodes over time. We systematically investigated the effect of storage conditions, including temperature and vacuum, on the performance of bare electrodes by recording EIS responses using [Fe (CN)₆]⁴⁻/³⁻ as a probe over two months. After receiving the GiPEC SPEs, they were stored under four different conditions: unvacuumed at room temperature (RT), unvacuumed at + 4 °C, vacuumed at RT, and vacuumed at + 4 °C. Storage stability was then assessed by recording EIS responses from Day 1 and repeating the measurements monthly over two months.

As illustrated in Fig. 7, after 30 days, the Rct of electrodes stored at 4 °C increased by 11.36%, while for the electrodes stored at RT, Rct surged by approximately 189.09%, indicating a significant decrease in the performance of bare electrodes. On the other hand, for the electrodes stored in a vacuum at 4 °C, Rct increased by 18.73%, whereas under vacuum at RT, it rose by 56.71%. After 60 days, the Rct of electrodes stored at 4 °C and those stored in a vacuum at 4 °C showed negligible changes, while electrodes stored under other conditions exhibited a significant increase in Rct.

Fig. 7.

Fig. 7

Open in a new tab

The effect of storage conditions on the electrochemical characteristics of bare electrodes was evaluated by recording electrochemical impedance spectroscopy (EIS) responses under four different conditions: unvacuumed at room temperature, unvacuumed at + 4 °C, vacuumed at room temperature, and vacuumed at + 4 °C.

As a result, the electrochemical performance of GiPEC screen-printed electrodes experienced a notable decline when stored at RT for two months (Fig. 7). Conversely, the electrodes stored under vacuum at + 4 °C exhibited consistent electrochemical signals over time. This suggests that storing GiPEC screen-printed electrodes at + 4 °C under a vacuum effectively inhibits the decrement of their electrochemical characteristics. A similar signal deduction was also obtained by the CV method. Therefore, storage under vacuum and at + 4 °C represents an optimal environment for preserving the integrity and performance of our bare electrodes, also supported by the relevant literature55–57 This optimized storage condition enhances the reproducibility and long-term stability of biosensors fabricated with these electrodes, paving the way for scalable manufacturing and wider implementation in POC diagnostic applications.

Validating the QC strategy

To validate the introduced QC strategy, bare electrodes were divided into two groups: those with the QC strategy implemented (treatment group) and those without it (control group). These groups were used to fabricate MIP biosensors for detecting agmatine and GFAP. The effectiveness of the QC strategy was evaluated by comparing the analytical performance of the MIP biosensors in PBS. Five MIP biosensors from each batch (treatment and control) were selected visually by a lab researcher for the detection of agmatine and GFAP. All biosensors were prepared using the same experimental procedure. Measurement variation was assessed using the relative standard deviation (RSD), reflecting the reproducibility of the MIP biosensors. Agmatine MIP biosensors in the treatment group showed a 79% reduction in RSD compared to the control group (RSD = 2.05% treatment, RSD = 9.68% control). Similarly, GFAP MIP biosensors in the treatment group exhibited a 87% reduction in RSD compared to the control group (RSD = 1.44% treatment, RSD = 11.67% control). The results demonstrate a significant decrease in measurement variation for MIP biosensors fabricated with the QC strategy, underscoring their reliability for diverse applications, including detecting metabolites and proteins. While the inclusion of QC steps may initially increase fabrication costs due to additional material characterization, instrumentation, and labor, the long-term advantages far outweigh these initial expenses. These benefits include significantly reduced waste, minimized batch-to-batch variability, and improved biosensor reliability (Fig. 8). As a result, production success rates are elevated, and costs associated with defective products are substantially reduced. This approach ultimately proves to be highly cost-effective for scalable production and successful commercialization (Table S2).

Fig. 8.

Fig. 8

Open in a new tab

Effective quality control strategy for controllable electrofabrication of biosensors.

Conclusion

This study introduced an effective QC strategy for screening electrodes to achieve highly reproducible MIP-based biosensors by leveraging real-time data generated during electrofabrication. The integration of PB NPs as embedded redox probes in the MIP biosensors provides a robust approach for monitoring and selecting electrodes based on critical parameters, including surface properties, conductivity, MIP film thickness, and template extraction efficiency, ensuring consistency across sensor batches. This allows for timely adjustments and optimal electrode selection based on key parameters such as conductivity and extraction efficiency, ultimately enhancing the reproducibility and performance of the sensors. By minimizing measurement variation, this strategy ensures greater consistency and reliability in biosensor fabrication. The proposed method employs non-destructive QC protocols at key fabrication stages, facilitating improved decision-making during electrode screening. This quality-controlled electrofabrication process produced biosensors capable of detecting metabolites and proteins with exceptional sensitivity and reproducibility. Results demonstrated that the QC strategy significantly improved the reproducibility of biosensors fabricated without QC measures. This marks a significant reduction in variability compared to bare screen-printed electrodes. By enabling the fabrication of reliable and reproducible biosensors using various electrofabrication processes, this strategy supports scalable production and broad adoption in point-of-care diagnostic applications.

Materials and methods

Materials

Grail fibrillary acidic protein (GFAP), agmatine, pyrrole, 3-aminophenyl boronic acid (APBA), potassium ferricyanide (K3Fe (CN)6), ferric chloride (FeCl3), sodium dodecyl sulfate (SDS), potassium chloride (KCl), hydrochloric acid (HCl), acetic acid, and phosphate buffer saline (PBS, 0.01 M, pH = 7.4) were acquired from Sigma-Aldrich. All the chemicals with the highest purity were commercially available and employed directly without additional purification in all experiments. Also, deionized (DI) water was utilized in all experiments.

Instrumentation

The surface morphologies, roughness parameters, and surface topography of sensing sites were evaluated by field emission scanning electron microscopy (FESEM, Sigma- ZEISS, Germany) and atomic force microscopy (AFM, Bruker Nanoscope, Germany). All electrochemical tests in this study were conducted on Autolab electrochemical analyzer model PGSTAT 204 (Metrohm, USA) operated with NOVA software.

Preparation of MIP biosensors

The fabrication process of MIP biosensors involves three steps: electrodeposition of PB nanoparticles, electropolymerization of the MIP film, and extraction of template molecules using two different approaches: electro-cleaning for agmatine biosensors and solvent extraction for GFAP MIP biosensors.

In the first step, PB NPs were electrodeposited as the embedded redox probe onto the surface of our conductive GiPEC screen-printed electrode58. The electrode consists of poly (3,4-ethylene dioxythiophene) polystyrene sulfonate (PEDOT: PSS) polymer embedded within the base carbon matrix. This process was carried out using CV within the potential range of -0.2 to 0.6 V versus Ag/AgCl at a scan rate of 50 mV s− 1 for 60 cycles. The solution used for electrodeposition contained 5 mM Inline graphic, 5 mM Inline graphic, in the presence of 100 mM KCl and 100 mM HCl.

To fabricate the agmatine MIP biosensor, the target imprinted polymer was electrochemically synthesized on a PB/GiPEC electrode via electropolymerization. This process took place in a PBS solution (pH 6.5) containing 5 mM agmatine, 37.5 mM pyrrole, and 12.5 mM APBA. Electropolymerization was performed by CV scanning from 0 to 1 V for 10 cycles at 50 mV s⁻¹. Following polymerization, the imprinted agmatine in the polymer film was extracted using an over-oxidation method (Approach A: Electro-cleaning) by CV scans from − 0.2 to 0.8 V at 50 mV s⁻¹ in 500 mM HCl for 15 cycles. After the template molecules were extracted, the MIP electrode was rinsed three times with DI water to remove any remaining agmatine residues from the Ppy film.

To prepare the GFAP MIP biosensor, the PB/GiPEC electrode underwent electropolymerization using CV from 0 to 1 V at a scan rate of 50 mV. s⁻¹ for 10 cycles. The process was conducted in a PBS solution (pH 6.5) containing 25 µg. mL− 1 GFAP, 60 µg. mL− 1 pyrrole, and 30 µg. mL⁻¹ APBA. Following electropolymerization, the GFAP was extracted by immersing the electrode in a washing solution of 10% (w/v) SDS and 5% (v/v) acetic acid in the DI water. This extraction method (Approach B: Solvent extraction) involved shaking the electrode at 100 rpm for one hour to create complementary cavities for GFAP. The MIP electrode was then rinsed three times with DI water to remove any remaining GFAP residues and gently dried with a nitrogen flow.

Storage protocol

The storage protocol involved creating different groups (n = 18 electrodes per group). On Day 1, after manufacturing of GiPEC screen-printed electrodes, one control group was left at room temperature (RT, 25 ± 1 °C) for 30 min prior to recording the EIS response. One-fourth of the electrodes were vacuum-sealed and then divided, with half stored in a fridge at + 4 °C and the other half at RT. Another one-fourth were stored without vacuum sealing, subdivided similarly into fridge and RT conditions. Thus, the groups consisted of one control group at RT, one group stored in the fridge at + 4 °C without vacuum, one group stored at RT without vacuum, and two groups stored under vacuum conditions (one at + 4 °C and the other at RT). Storage stability was assessed by recording EIS responses on Day 1 and repeating this each month over two months.

Electrochemical measurements

The electrochemical behavior of the MIP biosensors during the electro-fabrication steps was investigated by CV. For electrodeposition, the potential range was − 0.2 to 0.6 V at a scan rate of 50 mV s− 1. For electropolymerization, the potential range was 0 to 1 V at a scan rate of 50 mV s− 1. The SWV measurements for the final QC assessment of the agmatine MIP biosensor were performed by dispensing 150 µL of 0.01 M PBS solution at pH 7.4 onto the MIP biosensor surface. SWV measurements were taken in an anodic direction range of -0.8 to 0.8 V, with a frequency of 10 Hz, an amplitude of 100 mV, and an E step (step potential of staircase) of 10 mV over a total duration of 25 s. For the final QC assessment of the GFAP MIP biosensor, a 150 µl PBS (0.01 M, pH = 7.4) droplet was dispensed on the prepared electrode to measure the impedance of each electrode via the EIS method. The EIS test was conducted in the frequency range of 20 kHz to 1 Hz with an amplitude of 10 mV and an open circuit potential (Eocp) of 0.17 V.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Supplementary Material 1 (10.6MB, xlsx)
Supplementary Material 2 (1.2MB, docx)

Acknowledgements

The authors acknowledge funding supports from Fluidome Inc.’s sponsor, Mathematics of Information Technology and Complex Systems (Mitacs) Accelerate, the Alberta Innovates, the Canadian Institutes of Health Research (CIHR), and the Canada Research Chair.

Author contributions

Concept and design: B.B., M.F., and M.M; Experimentation and data acquisition: B.B. and M.F; Data analysis and interpretation: B.B., M.F., M.M., and A.S.N.; Sample analysis: B.B. and M.F.; Drafting and revising the manuscript: B.B., M.F., M.M, and A.S.N.. B.B. and M.F. are Co-first authors, Contributed equally.

Data availability

All data generated or analysed during this study are included in this published article and its supplementary information files (Scientific Report-Raw Data). Materials and instrumentation applied for the fabrication of molecularly imprinted polymer biosensors, storage protocol, electrochemical measurements, and atomic force microscopy and scanning electron microscopy analyses of MIP biosensors.

Declarations

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

These authors contributed equally: Bahareh Babamiri and Mohammadreza Farrokhnia.

Contributor Information

Mehdi Mohammadi, Email: mehdi.mohammadiashan@ucalgary.ca.

Amir Sanati Nezhad, Email: amir.sanatinezhad@ucalgary.ca.

References

  • 1.Wang, J. Electrochemical biosensors: towards point-of-care cancer diagnostics. Biosens. Bioelectron.21, 1887–1892 (2006). [DOI] [PubMed] [Google Scholar]
  • 2.Baranwal, J., Barse, B., Gatto, G., Broncova, G. & Kumar, A. Electrochemical sensors and their applications: a review. Chemosensors10, 363 (2022). [Google Scholar]
  • 3.Lee, D. H., Lee, W. Y. & Kim, J. Introducing nanoscale electrochemistry in small-molecule detection for tackling existing limitations of affinity-based label-free biosensing applications. J. Am. Chem. Soc.145, 17767–17778 (2023). [DOI] [PubMed] [Google Scholar]
  • 4.Haghayegh, F., Salahandish, R., Hassani, M. & Sanati-Nezhad, A. Highly stable buffer-based zinc oxide/reduced graphene oxide nanosurface chemistry for rapid Immunosensing of SARS-CoV-2 antigens. ACS Appl. Mater. Interfaces. 14, 10844–10855 (2022). [DOI] [PubMed] [Google Scholar]
  • 5.Maduraiveeran, G., Sasidharan, M. & Ganesan, V. Electrochemical sensor and biosensor platforms based on advanced nanomaterials for biological and biomedical applications. Biosens. Bioelectron.103, 113–129 (2018). [DOI] [PubMed] [Google Scholar]
  • 6.Anirudhan, T. S., Deepa, J. R. & Binussreejayan Electrochemical sensing of cholesterol by molecularly imprinted polymer of silylated graphene oxide and chemically modified nanocellulose polymer. Mater. Sci. Eng. C. 92, 942–956 (2018). [DOI] [PubMed] [Google Scholar]
  • 7.Gurudatt, N. G., Akhtar, M. H., Park, D., Kim, K. B. & Shim, Y. B. Catalytic SrMoO4 nanoparticles and conducting polymer composite sensor for monitoring of K+-induced dopamine release from neuronal cells. J. Mater. Chem. B. 10, 728–736 (2022). [DOI] [PubMed] [Google Scholar]
  • 8.Akhtar, M. H., Hussain, K. K., Gurudatt, N. G., Chandra, P. & Shim, Y. B. Ultrasensitive dual probe immunosensor for the monitoring of nicotine induced brain derived neurotrophic factor released from cancer cells. Biosens. Bioelectron.116, 108–115 (2018). [DOI] [PubMed] [Google Scholar]
  • 9.Akhtar, M. H., Mir, T. A., Gurudatt, N. G., Chung, S. & Shim, Y. B. Sensitive NADH detection in a tumorigenic cell line using a nano-biosensor based on the organic complex formation. Biosens. Bioelectron.85, 488–495 (2016). [DOI] [PubMed] [Google Scholar]
  • 10.Akhtar, M. H., Hussain, K. K., Gurudatt, N. G. & Shim, Y. B. Detection of Ca2+-induced acetylcholine released from leukemic T-cells using an amperometric microfluidic sensor. Biosens. Bioelectron.98, 364–370 (2017). [DOI] [PubMed] [Google Scholar]
  • 11.Abbas, W. et al. Facilely green synthesis of 3D nano-pyramids cu/carbon hybrid sensor electrode materials for simultaneous monitoring of phenolic compounds. Sens. Actuators B Chem.282, 617–625 (2019). [Google Scholar]
  • 12.Anik, U. Electrochemical medical biosensors for POC applications. Med. Biosens. Point Care (POC) Appl. 275–292 (2017).
  • 13.da Silva, E. T. S. G. et al. Electrochemical biosensors in point-of-care devices: recent advances and future trends. ChemElectroChem4, 778–794 (2017). [Google Scholar]
  • 14.Wu, J., Liu, H., Chen, W., Ma, B. & Ju, H. Device integration of electrochemical biosensors. Nat. Rev. Bioeng.1, 346–360 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Shajari, S. et al. MicroSweat: A wearable microfluidic patch for noninvasive and reliable sweat collection enables human stress monitoring. Adv. Sci.10, 2204171 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Joe, C., Lee, B. H., Kim, S. H., Ko, Y. & Gu, M. B. Aptamer duo-based portable electrochemical biosensors for early diagnosis of periodontal disease. Biosens. Bioelectron.199, 113884 (2022). [DOI] [PubMed] [Google Scholar]
  • 17.Patel, T., Huang, J. & Krukiewicz, K. Multifunctional organic monolayer-based coatings for implantable biosensors and bioelectronic devices: review and perspectives. Biosens. Bioelectron.14, 100349 (2023). [Google Scholar]
  • 18.Grau, G. et al. Gravure-printed electronics: recent progress in tooling development, Understanding of printing physics, and realization of printed devices. Flex. Print. Electron.1, 023002 (2016). [Google Scholar]
  • 19.Gao, F. et al. Wearable and flexible electrochemical sensors for sweat analysis: a review. Microsyst. Nanoeng. 9, 1–21 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Harris, A. R. et al. Efficient strategy for quality control of screen-printed carbon ink disposable sensor electrodes based on simultaneous evaluation of resistance, capacitance and Faradaic current by fourier transform AC voltammetry. J. Solid State Chem.12, 1301–1315 (2008). [Google Scholar]
  • 21.Qian, H. et al. Improving high throughput manufacture of laser-inscribed graphene electrodes via hierarchical clustering. Sci. Rep.14, 7980 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Parrilla, M., Cánovas, R., Jeerapan, I., Andrade, F. J. & Wang, J. A textile-based stretchable multi-ion potentiometric sensor. Adv. Healthc. Mater.5, 996–1001 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Kelíšková, P., Matvieiev, O., Janíková, L. & Šelešovská, R. Recent advances in the use of screen-printed electrodes in drug analysis: A review. Curr. Opin. Electrochem.42, 101408 (2023). [Google Scholar]
  • 24.Ravizza, A. et al. Comprehensive review on current and future regulatory requirements on wearable sensors in preclinical and clinical testing. Front. Bioeng. Biotechnol.7, 1–9 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Jain, A. & Ganesh, N. Quality standards for medical devices. Int. J. Drug Regul. Affairs Www Ijdra Com. (2014).
  • 26.Martins, E. G. & De Lima, P. Gouvea Da Costa, S. E. Developing a quality management system implementation process for a medical device manufacturer. J. Manuf. Technol. Manag.26, 955–979 (2015). [Google Scholar]
  • 27.Cong, H. & Zhang, N. Perspectives in translating microfluidic devices from laboratory prototyping into scale-up production. Biomicrofluidics16, 021301 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Ulucan-Karnak, F., Kuru, C. İ. & Akgöl, S. Sensor commercialization and global market. In Advanced Sensor Technology: Biomedical, Environmental, and Construction Applications 879–915 (2022).
  • 29.Rogers, K. R. Principles of Affinity-Based biosensors. Mol. Biotechnol.14 (2000). [DOI] [PubMed]
  • 30.Dykstra, G., Reynolds, B., Smith, R., Zhou, K. & Liu, Y. Electropolymerized molecularly imprinted polymer synthesis guided by an integrated data-driven framework for cortisol detection. ACS Appl. Mater. Interfaces. 14, 25972–25983 (2022). [DOI] [PubMed] [Google Scholar]
  • 31.Raziq, A. et al. Development of a portable MIP-based electrochemical sensor for detection of SARS-CoV-2 antigen. Biosens. Bioelectron.178, 113029 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Yeasmin, S., Ullah, A., Wu, B., Zhang, X. & Cheng, L. J. Enzyme-mimics for sensitive and selective steroid metabolite detection. ACS Appl. Mater. Interfaces. 15, 13971–13982 (2022). [DOI] [PubMed] [Google Scholar]
  • 33.Siavash Moakhar, R. et al. A versatile biomimic nanotemplating fluidic assay for multiplex quantitative monitoring of viral respiratory infections and immune responses in saliva and blood. Adv. Sci.9, 2204246 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Parlak, O., Keene, S. T., Marais, A., Curto, V. F. & Salleo, A. Molecularly selective nanoporous membrane-based wearable organic electrochemical device for noninvasive cortisol sensing. Sci. Adv.4, 2904 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Pilvenyte, G. et al. Molecularly imprinted polymers for the determination of cancer biomarkers. Int. J. Mol. Sci.24, 4105 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Rebelo, T. S. C. R. et al. A disposable saliva electrochemical MIP-based biosensor for detection of the stress biomarker α-Amylase in point-of-care applications. Electrochem2, 427–438 (2021). [Google Scholar]
  • 37.Hassine, A., Ben, Raouafi, N. & Moreira, F. T. C. Novel electrochemical molecularly imprinted polymer-based biosensor for Tau protein detection. Chemosensors9, 238 (2021). [Google Scholar]
  • 38.Zidarič, T., Majer, D., Maver, T., Finšgar, M. & Maver, U. The development of an electropolymerized, molecularly imprinted polymer (MIP) sensor for insulin determination using single-drop analysis. Analyst148, 1102–1115 (2023). [DOI] [PubMed] [Google Scholar]
  • 39.Rajpal, S. & Mishra, P. Next generation biosensors employing molecularly imprinted polymers as sensing elements for in vitro diagnostics. Biosens. Bioelectron.11, 100201 (2022). [Google Scholar]
  • 40.Li, Y. et al. Highly sensitive protein molecularly imprinted electro-chemical sensor based on gold microdendrites electrode and Prussian blue mediated amplification. Biosens. Bioelectron.42, 612–617 (2013). [DOI] [PubMed] [Google Scholar]
  • 41.Tang, W. et al. Touch-based stressless cortisol sensing. Adv. Mater.33, (2021). [DOI] [PubMed]
  • 42.Sadriu, I. et al. Molecularly imprinted polymer modified glassy carbon electrodes for the electrochemical analysis of isoproturon in water. Talanta207, 120222 (2020). [DOI] [PubMed] [Google Scholar]
  • 43.Jafari, S., Dehghani, M., Nasirizadeh, N. & Azimzadeh, M. An Azithromycin electrochemical sensor based on an aniline MIP film electropolymerized on a gold nano urchins/graphene oxide modified glassy carbon electrode. J. E. C. 829, 27–34 (2018). [Google Scholar]
  • 44.Suriyanarayanan, S., Mandal, S., Ramanujam, K. & Nicholls, I. A. Electrochemically synthesized molecularly imprinted polythiophene nanostructures as recognition elements for an aspirin-chemosensor. Sens. Actuators B Chem.253, 428–436 (2017). [Google Scholar]
  • 45.Pilvenyte, G. et al. Molecularly imprinted polymer-based electrochemical sensors for the diagnosis of infectious diseases. Biosensors13, 620 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Mahony, J. O., Nolan, K., Smyth, M. R. & Mizaikoff, B. Molecularly imprinted polymers - Potential and challenges in analytical chemistry. Anal. Chim. Acta. 534, 31–39 (2005). [Google Scholar]
  • 47.Elbadawi, M., Ong, J. J., Pollard, T. D., Gaisford, S. & Basit, A. W. Additive manufacturable materials for electrochemical biosensor electrodes. Adv. Funct. Mater.31, 2006407 (2021). [Google Scholar]
  • 48.Paimard, G., Ghasali, E. & Baeza, M. Screen-printed electrodes: fabrication, modification, and biosensing applications. Chemosensors11, 113 (2023). [Google Scholar]
  • 49.Salahandish, R., Haghayegh, F., Khetani, S., Hassani, M. & Nezhad, A. S. Immuno-affinity potent strip with pre-embedded intermixed PEDOT:PSS conductive polymers and graphene nanosheets for bio-ready electrochemical biosensing of central nervous system injury biomarkers. ACS Appl. Mater. Interfaces. 14, 28651–28662 (2022). [DOI] [PubMed] [Google Scholar]
  • 50.Abdulbari, H. A. & Basheer, E. A. M. Electrochemical biosensors: electrode development, materials, design, and fabrication. Chem. BioEng. Reviews. 4, 92–105 (2017). [Google Scholar]
  • 51.Fernández, H. et al. Multivariate optimization of electrochemical biosensors for the determination of compounds related to food safety—a review. Biosensors13, 694 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Li, Y. et al. Recent advances in molecularly imprinted polymer-based electrochemical sensors. Biosens. Bioelectron.249, 116018 (2024). [DOI] [PubMed] [Google Scholar]
  • 53.Ngoc, H. T. L. P. T. Ho Xuan Anh Vu, Nguyen Dang Giang Chau & Ho Van Minh Hai. Fabrication and evaluation of some electrochemical properties of screen-printed electrodes for use in electrochemical analysis. HUJOS: Nat. Sci.133, 99–108 (2024). [Google Scholar]
  • 54.Behrent, A., Borggraefe, V. & Baeumner, A. J. Laser-induced graphene trending in biosensors: Understanding electrode shelf-life of this highly porous material. Anal. Bioanal Chem.416, 2097–2106 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Hurst, J. M., Kim, M. A., Peng, Z., Li, L. & Liu, H. Assessing and mitigating surface contamination of carbon electrode materials. Chem. Mater.31, 7133–7142 (2019). [Google Scholar]
  • 56.Cameron, J. & Skabara, P. J. The damaging effects of the acidity in PEDOT:PSS on semiconductor device performance and solutions based on non-acidic alternatives. Mater. Horiz. 7, 1759–1772 (2020). [Google Scholar]
  • 57.Kwon, J. et al. Conductive ink with circular life cycle for printed electronics. Adv. Mater.34, 2202177 (2022). [DOI] [PubMed] [Google Scholar]
  • 58.Pinilla, S., Coelho, J., Li, K., Liu, J. & Nicolosi, V. Two-dimensional material inks. Nat. Rev. Mater.7, 717–735 (2022). [Google Scholar]

Associated Data

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

Supplementary Materials

Supplementary Material 1 (10.6MB, xlsx)
Supplementary Material 2 (1.2MB, docx)

Data Availability Statement

All data generated or analysed during this study are included in this published article and its supplementary information files (Scientific Report-Raw Data). Materials and instrumentation applied for the fabrication of molecularly imprinted polymer biosensors, storage protocol, electrochemical measurements, and atomic force microscopy and scanning electron microscopy analyses of MIP biosensors.

Articles from Scientific Reports are provided here courtesy of Nature Publishing Group

RESOURCES