Bioelectronic cell enrichment

Abstract

Herein is described a bioelectronic cell enrichment platform and methods of use thereof. Said bioelectronic cell enrichment platform comprising; a microelectrode array (MEA) comprising a glass substrate coated with a conducting surface layer and a smart polymer, said smart 5 polymer being coated with a capture substrate, said capture substrate being coated with a selective capture reagent, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises gold nanospheres (AuNP).

Description

[0001] BIOELECTRONIC CELL ENRICHMENT FIELD OF THE INVENTION The present invention relates to bioelectronic cell enrichment platforms and use of such platforms. In particular, the bioelectronic cell enrichment platforms of the invention can be used to detect and enrich specific cell types in vitro which may be indicative of a disease, risk of developing a disease or disease status. BACKGROUND TO THE INVENTION Cell samples are commonly analysed to identify different cell types. Different cell types may be indicative of disease, risk of disease or progression of disease. However, in any one cell sample, the majority of cells, perhaps greater than 90%, are likely to be healthy or wild-type cells, even if diseased cells or other cells of interest are present (Masqué-Soler, N. et. al., EBioMedicine 76, (2022)). This high proportion of healthy cells may impact the sensitivity of any subsequent molecular diagnostic assays carried out on the cell sample. Therefore, there is a need for measures to selectively enrich the representation of specific target cell types or groups in a cell sample prior to further analysis. Conventional methods of cell selection and collection are often complex, requiring either size- dependent sorting or invasive enzymatic reactions. Furthermore, releasing collected cells may be complex and harmful to the cells, leading to disrupted cell integrity and cell death. Bioelectronic cell enrichment platforms are an alternative way to selectively isolate and enrich target cells from other cells or contaminants in cell sample. Biological components, such as antibodies or lectins, which selectively bind to the target cells are identified. When the target cells bind, they alter the local electrical environment. This change in electrical properties is detected by electronic sensors in a bioelectronic device, which applies a localized force or voltage to capture the cells, allowing other cells to pass through. The captured target cells can then be retained for further analysis. Capture and subsequent release of target cells in a bioelectronic cell enrichment platform may take place on a conductive substrate, enabling electronic monitoring and quantification (see for example Saez et. al., (2023) Methods Mol Biol.2679:305-314). Improved method using smart polymers, are known. For example, Saez et. al., (2023, as per above) describes as functional polymer film, which combines the thermoresponsive poly(N- isopropylacrylamide) and the conducting poly(3,4-ethylenedioxythiopene) / poly(styrene sulfonate) (PEDOT:PSS), and its use for the capture and release of circulating tumour cells. However, problems with this system include that the capture and particularly release are not selective. Furthermore, the target cells released after capture with existing systems may not be viable as the release mechanisms may be harsh. The current inventors herein provide a improved bioelectronic cell enrichment platform using more selective capture reagents such as lectins. Cell release is gentle which allows downstream testing and / or quantification of captured cells. Previous platforms do not achieve the goal of enrichment together with release for molecular characterisation. SUMMARY OF THE INVENTION Herein we describe a bioelectronic cell enrichment platform comprising; a microelectrode array (MEA) comprising a glass substrate coated with a conducting surface layer and a smart polymer, said smart polymer being coated with a capture substrate, said capture substrate being coated with a selective capture reagent, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises metal nanospheres, suitably gold nanospheres (AuNP). Suitably, the selective capture reagent selectively binds to cell surface markers. For example, the selective capture reagent may be an antibody which binds to cell-surface proteins, or a glycan which binds to cell-surface lectins. Suitably, the selective capture reagent is a lectin, suitably wherein said selective capture reagent is a functionalised lectin, suitably Erythrina Cristagalli lectin. Suitably, the capture substrate comprises AuNP having a diameter of 20-100nm, suitably 30- 70nm, suitably 40-60nm, most suitably 50nm. Suitably, the smart polymer releases at least the captured target cells when heated, suitably when heated to 20-40°C, suitably when heated to 30-40°C, suitably when heated to 23-37°C, most suitably when heated to 32-37°C. In preferred embodiments the target cells are not disrupted or damaged by capture or release. In preferred embodiments, captured target cells are labelled, suitably with a fluorescent label. In some embodiments the smart polymer releases the capture substrate, selective capture reagent and captured target cells when heated, suitably when heated to 20-40°C, suitably when heated to 30-40°C, suitably when heated to 23-37°C, most suitably when heated to 32- 37°C. In preferred embodiments the target cells are not disrupted or damaged by capture or release. Suitably, the conducting surface layer comprises any conducting surface. Suitably, said conducting surface layer comprises one or more of gold, indium, tin oxide, graphene, carbon nanotubes or graphite. In preferred embodiments said conducting surface layer comprises gold. Suitably, the selective capture reagent selectively binds to cells associated with a disease or indicative of disease. Said disease may suitably be cancer, a bacterial disease, a viral disease, an early stage disease, a late stage disease, a genetic disease, an autoimmune disease, H.Pylori or Streptococcus pneumoniae. In preferred embodiments the selective capture reagent selectively binds to cancer cells or cancer associated cells. In one embodiment the selective capture reagent binds to cells associated with oesophageal cancer cells such as oesophageal squamous cell carcinoma cells and oesophageal adenocarcinoma cells. Suitably, the selective capture reagent selectively binds to cells associated with Barret’s oesophagus. In preferred embodiments the selective capture reagent binds to gastric and intestinal cells types, goblet cells and / or columnar epithelium cells. Suitably, the microarray electrode comprises an optical window. Said optical window allows monitoring of cell capture and / or functionalisation of the selective capture reagent, suitably when labelling can be visualised, for example of fluorescent labels have been used. Suitably the optical window is found in the core of the microelectrode array. The present invention further includes a method of selectively obtaining target cells from a cell sample, said method comprising the steps of (a) incubating a cell sample with the bioelectronic cell enrichment platform of the invention as described herein; (b) washing away unbound cells; (c) heating to release captured target cells. Suitably, the heating step (c) is to 20-40°C, suitably 30-40°C, suitably to 23-37°C, most suitably 32-37°C. Suitably, the target cells are disease cells, cells associated with or indicative of a disease or cancer cells. The target cells may be labelled, such as with a fluorescent label. Suitably, there is an additional step of analysing the target cells. Analysing may include microscopy, selective labelling, counting, fixing. Suitably, the method of selectively obtaining target cells from a cell sample, includes the additional steps of:- (i) applying an alternating current voltage to the microelectrode array; (ii) measuring electrochemical impedance spectroscopy (EIS) signals before and after cell capture; (iii) correlating impedance changes to cell capture efficiency. Suitably steps (i)-(iii) occur after step (a) of the method described above. The present invention further includes use of the bioelectronic cell enrichment platform of the invention to identify cells associated with disease or risk of a disease. Suitably, the disease or risk of disease is Barrett’s oesophagus or cancer. Suitably, the disease or risk of disease is a bacterial disease, a viral disease, a genetic disease, an autoimmune disease. Suitably, the bioelectronic cell enrichment platform of the invention may be integrated with a microfluidic system. The invention further includes a bioelectronic cell enrichment platform comprising; a microelectrode array comprising a glass substrate coated with gold and at least two layers of insulating polymer, wherein multiple etching windows are created in the insulating polymer; wherein a smart polymer is layered on the gold inside one or more etching windows, wherein a selective capture reagent is covalently coupled to a capture substrate layer which is electrostatically bound to the smart polymer surface, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises AuNP. SOME ADVANTAGES The present invention has the advantage over non-bioelectronic enrichment platforms of not requiring processing of the cellular sample in formalin fixed paraffin embedded (FFPE) blocks and slides as used in other methods. FFPE embedding processes can introduce chemical modifications, cross-linkages and breaks in nucleic acids, as well as being labour intensive. Capture of target cells may use any selective capture reagent, including for example glucans or lectins. The present invention is not limited to the use of antibodies as the selective capture reagent, though they may be used if preferred. The present invention may be used to enrich cells samples to identify disease or the stage of a disease in a mixed cell sample, avoiding the need for the patient to undergo more invasive procedures to obtain a cell sample with fewer contaminants. The target cells captured by the present invention are released easily by a gentle heating process. As the heating step takes place at or near body temperature, the target cells are undamaged and may therefore be more easily used for further analysis. BRIEF DESCRIPTION OF THE DRAWINGS Figure 1 is a schematic outlining the Cytosponge assay of Paterson A. L., (Diagn Cytopathol 48, 253–264 (2020)) as used in the examples. a) Cytosponge collect cells from the gastroesophageal junction and the entire length of the oesophagus. b) The sampled cells are retrieved from the sponge through a series of washing steps, followed by centrifugation to create a cell pellet. A cell clot is formed by introducing plasma-thrombin. c) The cell clot is fixed with formalin and processed into a paraffin block using standard laboratory protocols. d) Pathology slide-based evaluation of morphology with Haematoxylin and Eosin (H&E) stained sections and the immunohistochemistry (IHC) marker TFF3 (trefoil factor 3, a known protein biomarker). Figure 2 is a schematic diagram showing mart polymer-based cell capture and thermal release mechanism (adapted from Garcia-Hernando et. al., (Biosens Bioelectron 191, 113405 (2021)), a) Schematic depicting the mechanism of cell capture via fibronectin-based adhesion and their release by thermal actuation. b) Microelectrode array consisting of electrodes of different size and geometries c) Chemical structure of the smart polymer- (1)- the conducting polymer PEDOT:PSS with the crosslinker GOPS (3-glycidoxypropyltrimethoxysilane), (2) the thermo-responsive polymer comprising of the NIPAAm monomer, mBAAM (N, N’- Methylenebisacrylamide) crosslinker and DMPA (Dimethylolpropionic Acid) photo-initiator. Figure 3 is a schematic showing the bioelectronic enrichment platform of the invention and use thereof. a) A single Microelectrode array (MEA) chip showing a 6 electrode design. b) Single electrode integrated with a transparent optical window at its core to facilitate dual-mode optical-electrochemical sensing; the chemical structures of the smart polymer (1) PEDOT:PSS (further defined below) and (2) pNIPAAM (further defined below) ; and the polymer spin-coated electrode c) Schematic depicting the strategy employed for localised spin-coating of the hybrid polymer and localised functionalisation of lectins. d) Schematic of the electrochemical configuration used for electrochemical impedance spectroscopy (EIS) measurements. Randles circuit was used to fit and quantify the EIS measurements. RE= Reference electrode. CE = Counter Electrode. WE=Working electrode. Figure 4 is an enlarged diagram of Randles circuit as used in the previous figure. Randles circuit is an equivalent electrical circuit that consists of an active electrolyte resistance in series with the parallel combination of the double-layer capacitance Cdland an impedance (Zw) of a faradaic reaction. It is commonly used in electrochemical impedance spectroscopy (EIS) for interpretation of impedance spectra, often with a constant phase element (CPE) replacing the double layer capacity. The Randles equivalent circuit is one of the simplest possible models describing processes at the electrochemical interface. Figure 5 is a Schematic of the cell capture and thermal release mechanism. Figure 6 shows the optical characterisation of lectin-based cell capture and release exclusively on the PEDOT:PSS-pNIPAAM hybrid polymer. a) Lectins functionalised onto PEDOT:PSS coated glass slide b) Lectins functionalised onto PEDOT:PSS-pNIPAAM coated glass slide. Scale bars - 20μm. Figure 7 shows indium tin oxide (ITO)-based dual mode optical-electrochemical monitoring of lectin-based cell capture and release on the hybrid polymer (see Example 9). a) Schematic of cell capture on lectin functionalised polymer coated ITO platform. b) Schematic of the configuration used for dual optical-electrochemical monitoring, inset – brightfield microscopy of cells captured onto lectin-functionalised electrode (scale bars - 20μm), and subsequently released by thermal exposure. c), d), e) Faradaic EIS measurements obtained during the experiment represented as Bode, Nyquist and Randle’s circuit-based quantification of Rct (charge transfer resistance) respectively (see also figure 4). The solid and dashed lines in the Bode plot represent the magnitude and phase respectively. f) CV plot performed between - 0.3V and 0.7V with a scan rate of 0.1V / s. Figure 8 illustrates the selectivity of lectin-based cell capture using patient endoscopic brushings on the polymer-ITO platform (see example 10). a), b) Schematic depicting the endoscopic brushing of different tissue regions (BE and NE) to collect and test individual cell types. c), f) Depict the schematic of loading pure populations of each cell type separately onto lectin-functionalised polymer ITOs. d), g) Optical characterisation of cell capture / release of each cell type. Scale bars - 20μm), h) Simplified Randles circuit-based quantification of Rct values representing the lectin baseline, cell capture and cell release for each cell type tested. The error bars correspond to the standard deviation from the mean for n=3 devices. A one- way ANOVA was performed to assess significant differences in Rct values in e) and h), p>0.05, non-significant. Figure 9 shows mask designs for photolithography. a) Mask for gold patterning forming the gold electrode areas; b) Mask for patterning the polymer PEDOT:PSS-pNIPAAM with etching windows coinciding with the gold electrodes and contact pads. Figure 10 illustrates the functionalisation strategy for covalent coupling of lectin ECA onto the polymer-coated MEA. a) Schematic of each step in the functionalisation protocol. b) Graphical representation of each step followed by characterisation studies employed to track each layer of functionalisation.1- Bare gold electrode and brightfield microscopy with scale bar - 200μm, 2-polymer-coated electrode and AFM with scale bar - 1μm, 3- AuNP doping and SEM with scale bar – 500nm, 4- Lectin ECA-FITC coupling and fluorescence microscopy and 4- TIF- lifeact-RFP capture and fluorescence microscopy with scale bar 200μm. c), d) Represent the EIS measurements taken during each step of the functionalisation process with the Nyquist plot of raw data in e) and the simplified Randle’s circuit-based quantification of the Nyquist plot in f). Error bars correspond to the standard deviation from the mean for n=18. Pairwise t- test with Bonferroni correction indicated statistically significant differences (*** p ≤0.001) for all conditions relative to its previous measurement. Figure 11 Shows the results of mechanistic studies to evaluate the mechanism of cell release from the platform. a) 3D rendering of AFM data depicting the surface morphology of the polymer PEDOT:PSS-pNIPAAM at room temperature (RT) and post heating to 37°C. b) Representative 2D AFM with scale bar - 1μm and line plots of the polymer indicating the vertical heights of the polymeric chains at RT and at 37°C. c) Graphical depiction of the mechanistic characterisation experiment. d) Circuit-based quantification of Rct from the EIS measurements taken at each step during the experiment, starting with the polymer baseline. Error bars correspond to the standard deviation from the mean for n=18. Pairwise t-test with Bonferroni correction between the RT and post-heat condition for each step was carried out. Only the post-heat condition after cell capture indicated a significant difference *** p ≤0.001. Figure 12 shows cell capture homogeneity and sensitivity on the lectin-based bioelectronic sensor platform. a) EIS quantification demonstrating sensitivity and homogeneity of cell capture across the multiple electrodes of an MEA chip (n=6). b) EIS quantification demonstrating sensitivity and homogeneity of cell capture across three MEA chips (n=18). c) Representative images of optical monitoring of cell capture within the optical window. d) Linear regression analysis of cell capture across multiple MEA chips. Error bars in b), d) correspond to the standard deviation from the mean for n=18. Figure 13 shows the results of cell detachment studies on the bioelectronic platform. a) Schematic of the experiments conducted to evaluate thermal detachment efficiency (that is the release of captured target cells). Inset depicts the optical characterisation of TIF cell capture and release on the MEA platform with fluorescence microscopy. b), c) d) Nyquist, Bode and Rct plots representing the EC monitoring of cell capture and release across multiple MEA chips. Error bars represent standard deviation from the mean for n =18 electrodes. Pairwise t-test with Bonferroni correction was carried out. Both cell capture and post-release indicated a significant difference *** p ≤0.001. Figure 14 shows the results of target specific cell capture in highly mixed cellular sampling. a) Schematic representation of endoscopic brushings used to collect NE cells to which lectin ECA specific TIF cells were spiked in at different ratios. b) Different ratios prepared and tested independently to evaluate the platform’s selective capture capabilities. c) Representative optical microscopy images of the transparent window of the electrode for each ratio tested d) Rct quantification of EIS measurements indicating the selectivity of target cell capture in mixed sampling. Error bars correspond to the standard deviation from the mean for n=6 electrodes. Pairwise t-test with Bonferroni correction was carried out. Both ratios with TIFs spiked-in indicated a significant difference relative to the lectin baseline *p<0.05, *** p ≤0.001. Figure 15 shows the strategy for clinical testing of Cytosponge samples on the bioelectronic enrichment platform. Schematic depicting a microfluidic approach integrating several MEA arrays for processing Cytosponge sampling, and the multiple steps of clinical validation experiments carried out to evaluate columnar cell enrichment. Figure 16 shows EIS-based quantification of cells captured on the electrodes for each Cytosponge sample. Measurements provided for comparison of the lectin baselines, cells captured (after 1hr of processing) and cells released for Cytosponge sample collected from a healthy volunteer (left) a BE case with segment C0M4 (middle) and a BE case with segment C7M12. Error bars correspond to the standard deviation from the mean for n=6 electrodes collected across two MEAs (3 electrodes each). Pairwise t-test with Bonferroni correction across different conditions was carried out for each sample. Statistically significant increase in ΔRct values was observed for cells captured only for the two BE cases with *p<0.05, for BE segment C0M4 and *** p ≤0.001 for BE segment C7M12. Additionally, a significant decrease in ΔRct was observed for post release of captured cells for the BE case with C7M12 ** p ≤0.01. Figure 17 shows optical characterisation of cells captured within the electrode’s optical window during Cytosponge sample processing of BE cases. a) Graphical representation of two lectin-functionalised fluidic MEA devices, wherein the primary device (top) was used for brightfield evaluation of cell capture, and secondary device (bottom) used for on-chip immunofluorescence evaluation. b) MUC5AC IF staining carried out on cells captured in a parallel microfluidic MEA device. Scale bars are 100μm. Figure 18 shows representative images of optical characterisation of cells enrichment. Cytospin was performed on fixed cellular sample aliquots collected after the thermal detachment process during processing of the Cytosponge sample BEST2 / CAM / 0872. a) H&E staining of three groups of sample aliquots collected. Red circles indicate identified columnar cell clusters. b) MUC5AC IF staining of three groups of sample aliquots collected. Green – MUC5AC staining of columnar cells, Blue – DAPI nuclear staining. The scale bars indicated are 100 μm. Figure 19 shows RT-qPCR results representing nucleic-acid-based characterisation of cell enrichment. The relative gene expression of specific markers in aliquots of cellular samples collected during processing of the Cytosponge sample of the patient with long segment BE. Unpaired t-test was carried out across different sample fractions for each RNA biomarker. Significant increase in fold change values were observed for the enriched cell fraction for MUC5AC *p<0.05, and MUC2 ****,p<0.0001. Error bars correspond to the standard deviation from the mean for n=3. Figure 20 shows a Nyquist plot and the equivalent circuit. Figure 21 shows a Bode plot and equivalent circuit representing the magnitude (in blue) and phase (red). Figure 22 illustrates commonly employed types of EIS. DETAILED DESCRIPTION The present inventors looked to improve diagnosis of diseases and risk or diseases, by enriching cell samples for cells associated with said diseases. In particular, when the disease is cancer. Exemplary disease Oesophageal cancer presents a pressing global health concern, characterised by a high incidence and mortality rate, which poses significant challenges to healthcare systems worldwide. Oesophageal cancer manifests as a diverse disease encompassing two primary histological types: oesophageal squamous cell carcinoma (ESCC) and oesophageal adenocarcinoma (EAC). ESCC is often linked to tobacco and alcohol consumption, nutritional deficiencies, as well as specific environmental factors. It primarily affects the upper and middle sections of the oesophagus. In contrast, EAC primarily occurs in the lower part of the oesophagus and the gastro-oesophageal junction. It is closely linked to the gastroesophageal reflux disease (GERD) and its which may result is a further condition, Barrett's oesophagus (BE). BE is characterised by the oesophageal squamous epithelium transforming into a columnar epithelium, an adaptive response to prolonged gastro-oesophageal reflux of acid, bile, and other harmful substances. Normally, the damage caused by reflux is repaired through the regeneration of squamous cells. However, in some individuals, the squamous epithelium is gradually replaced by a differentiated columnar epithelium. This transformed tissue exhibits a glandular structure, consisting of a mosaic of gastric type epithelium and crypts resembling intestinal epithelium. Many cells types are indicative of BE, including gastric and intestinal cells types, goblet cells and / or columnar epithelium cells. The pathognomonic cell type (characteristic of BE, which can be used as a diagnostic of the condition) is the goblet cell. This condition is referred to as gastric and intestinal metaplasia (IM) respectively. Further cell dysphasia may lead to oesophageal cancer as described above. BE is recognised as a histological precursor of EAC. Therefore, when BE is diagnosed, patients generally enter into a surveillance or monitoring programme to assess progression to dysplasia and adenocarcinoma to facilitate early intervention. BE is typically detected through endoscopy and the histopathological examination of biopsy samples. The diagnosis is based on the presence of IM and various cell types discussed above. The current clinical management of BE involves controlling GERD, often experienced as heartburn, using acid-suppressing medications to reduce symptoms and to lower further damage to the oesophageal lining. In addition, individuals diagnosed with BE in may be offered endoscopic surveillance accompanied by biopsies at regular intervals. However, patients at risk are sometimes missed by these practices. Furthermore, endoscopy may be considered unpleasant and invasive by patients, leading to poor attendance and follow up. There is therefore an interest in non-endoscopic approaches for detection of BE and EAC. The main strategies involve either imaging-based assessment, such as using wireless capsule endoscopy, or cell-sampling devices such as balloon cytology, and the CytospongeTM. Cytosponge The CytospongeTMconsists of a small capsule attached to a string that is swallowed by the patient. Once in the stomach, the capsule dissolves to reveal a slightly abrasive polyurethane sponge that is then pulled back up through the oesophagus, a pan-oesophageal cell sample including ribbons of tissue and clumps of cells (See Figure 1). In usual clinical protocol, after sampling, the sponge is placed in a preservative fluid and subsequently processed into a formalin-fixed paraffin-embedded (FFPE) cell block in the processing laboratory (Figure 1 b, c). This material can be analysed to detect the presence of BE or its progression to dysplasia and cancer. Sections of the FFPE samples are cut and stained with H&E, and the IHC biomarker TFF3, to allow a direct comparison between the morphological appearances and biomarker status (depicted in Figure 1 d). In addition to BE-associated goblet cells, the expected cellular components from a Cytosponge sample will mainly include squamous epithelium from the oesophagus, and gastric type columnar epithelium from the stomach and / or hiatus hernia. Other cells that could be present include background immune cells, respiratory cells, tonsillar cells and isolated fungal spores. The present invention aims to solve the problem of excluding the unwanted cells from a cell sample, such as a Cytosponge sample, and enriching for preferred target cells which are indicative of a disease such as BE or EAC. Though using a Cytosponge cell sample to detect BE is used as an example herein, it will be apparent that the invention can be used on any cell sample, and any type of cell can be used as the target cell as long as a selective capture reagent can be used to bind to the target cell. Furthermore, the inventors unexpectedly found that the current invention enables the target cells to be released from capture undamaged, which enables further analysis of the target cells. This is another advantage over existing platforms for cell enrichment. Features of the invention The current invention is a bioelectronic cell enrichment platform. It uses a polymer-coated microelectrode array (MEA) for the selective capture, and possible quantification / further analysis, of target cells in a cell sample, and allows the retrieval of these captured cells, preferably live retrieval. The bioelectronic cell enrichment platform is used in vitro, and all the methods described herein are also in vitro. The bioelectronic cell enrichment platform of the current invention comprises a smart polymer, suitably in a film, and its use for the capture and release of target cells in a cell sample. When coated with microfabricated gold electrodes, the smart polymers are capable of noninvasively capturing and controllably releasing cells while, at the same time, monitoring these processes with conventional electrical measurements (see for example Saez et. al., as per the above). The base of the microelectrode array of the invention is referred to as a “substrate”. This substrate is an inert substance, preferably glass. Polymers The bioelectronic cell enrichment platform describe herein comprising; a microelectrode array (MCA) comprising a glass substrate coated with a conducting surface layer and a smart polymer, said smart polymer being coated with a capture substrate, said capture substrate being coated with a selective capture reagent, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises metal nanospheres, suitably gold nanospheres (AuNP). The term “conducting polymer” as used herein refers to a polymer with electrical conductive properties. Conductive polymers are organic polymers, typically containing a linear backbone with repeated units of conjugated monomers, with highly reversible redox behaviour and properties that resemble both metal and plastic materials. The most commonly used are polyaniline (PANI), polypirrole (PPy), poly(ethylenedioxithiophene) / poly(styrene sulfonate) (PEDOT:PSS), polythiophene (PTh) and polyfuran (PFu). Some polymers may require a process called "doping" to enhance their conductivity. Doping involves the introduction of chemical agents or dopants into the polymer matrix, which leads to the redistribution of charge carriers and the formation of charge carriers in the polymer structure. The doping process is crucial for some polymers because it introduces charge carriers (either positive or negative) into the polymer chain, allowing the material to conduct electricity. The choice of dopant can also influence the properties of the conducting polymer, such as its stability, conductivity, and responsiveness to external stimuli. “PEDOT” is a semiconductor formed by a chain of its EDOT monomers. PEDOT exhibits poor solubility in common solvents and a tendency to form aggregates limit its processability and stability. To overcome these challenges, PEDOT is doped with polystyrene sulfonate (“PSS” as referred to herein), a water-soluble polymer that enhances solubility, dispersibility, stability and conducting properties of the material . This forms the “PEDOT:PSS” as referred to herein. The chemical structure of PEDOT:PSS is shown below:- In order to improve the stability of PEDOT:PSS films in aqueous solutions, a cross-linking agent such as 3-glycidoxypropyltrimethoxysilane (GOPS) can be added to the mixture solution. The inclusion of GOPS has been found to enhance the mechanical and electrical stability of the films and reduce the swelling due to humidity (Donahue, M. J. et. al., Mater Sci Eng: R: Reports 140, 100546 (2020).). The combination of conducting polymers, such as PEDOT:PSS, with metal nanoparticles (also referred to herein as nanospheres), such as gold nanoparticles (Referred to herein as “AuNPs”), carbon nanotubes (referred to herein as “CNTs”), iron oxide nanoparticles (Fe2O3) and graphene has been shown to result in hybrid structures with enhanced properties and functionality. Nanostructures can improve the conductivity of the materials and provide larger surface area available for addition of interacting molecules ultimately resulting in significantly improved sensitivity (Dong Y. Z., et. al., Polymers (Basel) 12, (2020).). “Smart polymer” as used herein refers to stimuli-responsive polymers which mimic biopolymers and biological systems, and can respond to a variety of signals such as include temperature, ionic strength, solvent choice, the effect of radiation and applied fields as well as changes in pH, ion content, and chemical agents including ligands and enzymes. An example of a smart polymer is “PEDOT:PSS-pNIPAAM” (also referred to herein as “PEDOT:pNIPAAM”. These terms are used interchangeably herein). Poly(N-isopropylacrylamide) (referred to herein as “PNIPAAm”) is perhaps the most widely used smart polymer in the biomedical field owing to its ability to undergo temperature- dependent hydration and dehydration, causing hydrophilic and hydrophobic alterations. In an aqueous medium, PNIPAAm exhibits a temperature-dependent phase transition at ~32 °C, close to the body temperature, and can be modulated by incorporating hydrophobic and hydrophilic monomers. Exploiting this property, PNIPAAm is used in numerous biomedical applications including thermally modulated drug delivery systems, and in tissue engineering and cell culture substrates. Garcia-Hernando et. al., (Biosens Bioelectron 191, 113405 (2021)) first presented a bioelectronic platform for the capture and release of captured cells employing a smart polymer coating onto electrodes. The authors present for the first time, the use of a copolymer that combines the use of the thermo-responsive PNIPAAm with the electroactive material - PEDOT:PSS, to generate a smart surface for label-free and non-invasive monitoring of the capture and release of cells on gold electrodes by EIS. This is the PEDOT:PSS-pNIPAAM smart polymer as used herein. The PEDOT:PSS-pNIPAAM smart polymer can be layered with other polymers, especially insulating polymers, in the microelectrode array of the invention. A preferred insulating polymer to use is parylene. Parylene is the common name of a polymer whose backbone consists of para- benzenediyl rings −C6H4−connectedby 1,2-ethanediyl bridges−CH2−CH2−. It can be obtained by polymerization of para-xylylene H2C=C6H4=CH2. Suitably the microelectrode array of the invention comprises at least one, at least 3 or more than 2 layers of parylene. Parylene (or any insulating polymer) may be deposited on the MEA in a particular pattern. A mask may be used to achieve this pattern. The pattern may include gaps (as per the etching windows discussed below). Suitably PEDOT:PSS-pNIPAAM is coated with an insulating polymer, suitably parylene, using a spin method. That is spinning the substrate with the polymer or a prepolymer solution repeatedly to create layers. Layers may be baked or otherwise polymerised during the spin deposition process. One or more or all layers of insulating polymer, such as parylene, can be etched away, partially or completely, to form one or more etching window areas on the microelectrode array. These etching windows are for smart polymer deposition or electrical contact purposes. Selective capture reagent Suitably, the selective capture reagent selectively binds to cell surface markers. For example, the selective capture reagent may be an antibody which binds to cell-surface proteins, or a glycan which binds to cell-surface lectins. Suitably, the selective capture reagent is a lectin, A “lectin” as referred to herein is member of a class of proteins that bind to carbohydrates. Lectins possess the ability to recognise and bind specific carbohydrate structures, for example carbohydrate structures present on cell surfaces. Lectins can be used as an alternative to antibodies as recognition elements. Lectins can be classified based on their binding to different glycan structures. Some lectins bind to cancer-specific alterations in glycosylation. Suitably wherein said selective capture reagent is a functionalised lectin, suitably Erythrina Cristagalli lectin. Functionalisation of lectins is described in Example 2 below. This allows the lectins to attach to the surface of the polymers used in the invention, specifically the smart polymer. Preferably it is covalent functionalisation. Erythrina cristagalli lectin is a commercially available lectin of legume, plant origin. It is galactose-specific, binding to N-acetyllactosamine (LacNac). Suitably, the capture substrate comprises AuNP having a diameter of 20-100nm, suitably 30- 70nm, suitably 40-60nm, most suitably about 50nm or 50nm. The capture substrate can be part of the conducting surface or comprised within the conducting surface or preferably added as a further layer. Suitably, the smart polymer releases at least the captured target cells when heated, suitably when heated to 20-40°C, suitably when heated to 30-40°C, suitably when heated to 23-37°C, most suitably when heated to 32-37°C. In preferred embodiments the target cells are not disrupted or damaged by capture or release. In preferred embodiments, captured target cells are labelled, suitably with a fluorescent label, such as a labelled antibody. In some embodiments the smart polymer releases the capture substrate, selective capture reagent and captured target cells when heated, suitably when heated to 20-40°C, suitably when heated to 20-40°C, suitably when heated to 23-37°C, most suitably when heated to 32- 37°C. In preferred embodiments the target cells are not damaged, disrupted or damaged by capture or release. Released target cells are ideally alive and useable for further analysis. Further analysis can for example include PCR, sequencing, staining, microscopy, and counting. Suitably, the “conducting surface layer” comprises any conducting surface in a layer on the MEA substrate (suitably a glass substrate or a silicon substrate). Suitably, said conducting surface layer comprises one or more of gold, indium, tin oxide, graphene, carbon nanotubes or graphite. In preferred embodiments said conducting surface layer comprises gold. The conducting surface layer is preferably placed onto the substrate in a particular pattern. A mask may be used to achieve this pattern. The pattern may include gaps. The term “wafer” may also be used as an alternative to “layer”. Diseases Suitably, the selective capture reagent selectively binds to cells associated with a disease or indicative of disease or diagnostic or a disease. Said disease may suitably be cancer, a bacterial disease, a viral disease, an early stage disease, a late stage disease, a genetic disease, an autoimmune disease, H.Pylori or Streptococcus pneumoniae. In preferred embodiments the selective capture reagent selectively binds to cancer cells or cancer associated cells. In one embodiment the selective capture reagent binds to cells associated with oesophageal cancer cells such as oesophageal squamous cell carcinoma cells and oesophageal adenocarcinoma cells. Cells associated with a disease may be cells which are not specifically diseased cells themselves but are indicative of a disease (for example, only occur in patients who are suffering from a disease or are likely to suffer from a disease). They may be cells which occur at specific stages of a disease, for example at advanced (late) stages or early stages of a disease such as cancer. Cells indicative of a disease are cells which only occur in patients who are suffering from a disease or are likely to suffer (at risk) from a disease (or only rarely occur in patients who are not suffering from a specific disease / at risk of same disease). They may be cells which occur at specific stages of a disease, for example at advanced (late) stages or early stages of a disease such as cancer. These cells may be diagnostic of a disease. That is, the presence of these cells may be used to diagnose a patient with a disease. Suitably, the selective capture reagent selectively binds to cells associated with Barret’s oesophagus (BE). In preferred embodiments the selective capture reagent binds to gastric and intestinal cells types, goblet cells and / or columnar epithelium cells or any other cells associated with or diagnostic of BE. Suitably, the selective capture reagent selectively binds to cells which are indicative of a stage of a disease, such as early stage or late stage, or to cells which are indicative of the possibility of developing a disease. Suitably, the selective capture reagent selectively binds to cells which are diagnostic of a disease or diagnostic of a stage of a disease, such as early stage or late stage. The term “disease” as referred to herein may be any harmful deviation from the normal structural or functional state of a patient, preferably a human patient. Said disease may be of any origin, such as bacterial, viral, cancer, and / or autoimmune. Cells The aim of the present invention is to selectively capture target cells from a cell sample. The terms “cell sample” and “cellular sample” refers to the group of cells selected for analysis. The cell sample may be a cell-containing fluid such as blood, bone marrow or cerebrospinal fluid. Suitably, the cell sample can be obtained from a patient, from a medical or biological test (such as Cytosponge), from blood or other tissue. The cell sample initially contains a mix of different cell types. The term “target cells” as used herein refers to the type of cell or set of cells in a cell sample which are to be enriched because they are of interest to the researcher, due for example to their biological or indicative properties. Target cells may include immune cells and engineered immune cells. Suitably, an example target cell is columnar epithelium cells. The aim of the current invention is “enrichment” or “cell enrichment” of a cell sample. That is increasing the concentration of the target cells by selecting them from a cell sample containing mixed cell types. This is achieved by using a selective capture reagent selectively binds to target cells only. “Healthy” also known as “background” cells or “wildtype” cells are the non-target cells in a sample. Optical window Suitably, the microarray electrode comprises an optical window. Said optical window allows monitoring of cell capture and / or functionalisation of the selective capture reagent, suitably when labelling can be visualised, for example of fluorescent labels have been used. Said window may be used as an additional check for target cell binding and / or release. Suitably the optical window is found in the core of the microelectrode array. Methods The present invention further includes a method of selectively obtaining target cells from a cell sample, said method comprising the steps of (a) incubating a cell sample with the bioelectronic cell enrichment platform of the invention as described herein; (b) washing away unbound cells; (c) heating to release captured target cells. Suitably, the heating step (c) is to 20-40°C, suitably to 30-40°C, suitably to 23-37°C, most suitably 32-37°C. Suitably, the target cells are disease cells, cells associated with or indicative of a disease or cancer cells. The target cells may be labelled, such as with a fluorescent label. Suitably, there is an additional step of analysing the target cells. Analysing may include sequencing, PCR, microscopy, selective labelling, counting, fixing. Methods of the invention are carried out in vitro. Uses The present invention further includes use of the bioelectronic cell enrichment platform of the invention to identify cells associated with disease or risk of a disease. Uses of the invention are in vitro. Suitably, the disease or risk of disease is Barrett’s oesophagus or cancer. Suitably, the disease or risk of disease is a bacterial disease, a viral disease, a genetic disease, an autoimmune disease. In certain embodiments, the present invention may be used to enrich immune cell populations, including those relevant to cell therapy applications, provided suitable capture reagents for the desired cell surface markers are available. The invention further includes a bioelectronic cell enrichment platform comprising; a microelectrode array comprising a glass substrate coated with gold and at least two layers of insulating polymer, wherein multiple etching windows are created in the insulating polymer; wherein a smart polymer is layered on the gold inside one or more etching windows, wherein a selective capture reagent is covalently coupled to a capture substrate layer which is bound to the smart polymer surface, preferably electrostatically bound or covalently bound to the smart polymer surface, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises AuNP. In preferred embodiments, microelectrode array comprising a glass substrate coated with a conducting surface layer, preferably a patterned conducting surface layer, suitably a gold layer, two layers of insulating polymer, preferably parylene, and a smart polymer layer preferably PEDOT:PSS / pNIPAAm, The top layer or both layers of parylene are removed to pattern the smart polymer on the conducting surface layer. This creates etching window areas where the insulating polymer is partially, preferably completely, etched away for smart polymer deposition or contact purposes. Wherein a capture substrate, comprising a metal nanosphere, preferably a gold nanosphere (AuNP) is layered in one or more etching window areas. Wherein a selective capture reagent is coupled, preferably covalently, to the capture substrate layer which is bound to the smart polymer surface in the etching windows. Wherein the selective capture reagent is preferably a lectin, most preferably a functionalised lectin. Electrochemical Impedance Spectroscopy (EIS) EIS stands as a pivotal technique in biosensors, offering a non-intrusive and remarkably sensitive means of detecting and analysing biological molecules. In biosensors, where recognition elements like enzymes or antibodies interact with target biomolecules, EIS becomes indispensable for its ability to discern subtle alterations in the sensor's electrical characteristics. This high sensitivity allows for the detection of biomarkers at concentrations ranging from femtomolar to micromolar, crucial for early disease diagnosis and monitoring (Magar, H. S., Hassan, R. Y. A. & Mulchandani, A. Electrochemical Impedance Spectroscopy (EIS): Principles, Construction, and Biosensing Applications. Sensors (Basel) 21, (2021). 192. Bigdeli, I. K., Yeganeh). Notably, EIS facilitates label-free detection, minimising the need for additional chemical labels and enhancing the simplicity and efficiency of biosensing applications. EIS allows for the study of any intrinsic material property or specific processes that could influence the conductivity / resistivity or capacitance of an EC system(Bigdeli, I. K., Yeganeh, M., Shoushtari, M. T. & Zadeh, M. K. Electrochemical impedance spectroscopy (EIS) for biosensing. Nanosensors for Smart Manufacturing 533–554 (2021) ). For example, changes in the conductance of the electrode can be measured as a result of protein immobilisation and antibody-antigen reactions on the electrode surface (Hasan, M. R. et al. Recent development in electrochemical biosensors for cancer biomarkers detection. Biosens Bioelectron X 8, 100075 (2021), Cardoso, A. R. et al. An ultra-sensitive electrochemical biosensor using the Spike protein for capturing antibodies against SARS- CoV-2 in point-of-care. Mater Today Bio 16, 100354 (2022)). In this technique, a small sinusoidally varying potential (U) is applied, and the resulting current response (I) is measured (Lazanas, A. C. & Prodromidis, M. I. Electrochemical Impedance Spectroscopy─A Tutorial. ACS Measurement Science Au 2023, 162–193 (2022)). By varying the excitation frequency (f) of the applied potential over a range of frequencies, one can calculate the complex impedance. Therefore, EIS combines the analysis of both real and imaginary components of impedance, namely the electrical resistance and reactance, as shown in the equation below. This technique probes the electrode-solution interface and the redox couple to measure the following variables: • Rsol (the solution resistance includes resistances in the electrode material, the electrode contacts, and in the solution between the working and reference electrodes) • Rct (the resistance to charge transfer for the redox couple) • Cd (the double-layer capacitance) • Z (the impedance, which is the effective resistance of the electrochemical cell to the alternating current, including reactance and ohmic resistance) When plotting the real part of the impedance (Zreal) on the x-axis and the imaginary part (Zimag) on the y-axis, a "Nyquist Plot" is formed (Figure 20). Each point on the Nyquist plot represents an impedance value at a specific frequency, with Zimag being negative. On the x- axis, impedance on the right side of the plot corresponds to low frequencies, while impedances at higher frequencies are depicted on the left side. Additionally, on the Nyquist plot, impedance can be illustrated as a vector (arrow) with a length of |Z|. The angle formed between this arrow and the x-axis is referred to as the "phase angle” (Bigdeli, I. K.,.et. al., for Smart Manufacturing 533–554 (2021)). Suitable embodiments or alternatives described herein may also be considered as and described as preferable embodiments or alternatives. Another common way to present impedance results is through a Bode plot, which is more widely used in the engineering community compared to the Nyquist plot (Lazanas, A. C. & Prodromidis, M. I.. ACS Measurement Science Au 2023, 162–193 (2022)). A Bode plot consists of two separate logarithmic plots: magnitude versus frequency and phase versus frequency (Figure 21). One of the advantages of this plot is its ability to clearly display all the relevant information. For instance, the presence of a capacitor in parallel to a resistor, which is a significant circuit configuration in electrochemical impedance spectroscopy, can be observed as a peak in the phase shift on the plot (Lazanas as per the above). The Bode plot, in particular, allows for a more straightforward understanding of individual components. However, Nyquist plots are highly sensitive to small changes in the system. This sensitivity can be advantageous in detecting small variations in impedance, making Nyquist plots suitable for applications where precision is crucial (See Figure 21). EIS measurements can be fitted to an equivalent electrical circuit to extract quantitative information, which consists of the above measured variables connected to each other in different ways183. Parameters like solution resistance (Rs), charge transfer resistance (Rct), and double-layer capacitance (Cdl) are derived from the fits, providing insights into the electrochemical processes occurring at the interface. Randles circuit is presented as an example of this in Figure 21. EIS study of the electrode-electrolyte interface can be conducted in two modes: non-Faradaic and Faradaic response, both of which can be operated in a label-free configuration (Dorledo de Faria, R. A et. al., Int J Biosens Bioelectron Volume 5, (2019)). In the non-Faradaic mode of EIS (nf-EIS) (Figure 22), the interface can be ideally modelled by a resistance (Rs) in series with a capacitance (Cdl) representing the double layer capacitance. However, in reality, the interface response is non-ideal, so the heterogeneities can be accounted for by modelling Cdl using a constant phase element (CPE) (Garrote, B. L., et. al., ACS Sens 4, 2216–2227 (2019)). EXAMPLES Example 1 -MEA Mask Design and Fabrication Fabrication of the MEA involves the utilisation of photolithography and the parylene C lift-off method. This approach enables precise patterning of the smart polymer PEDOT:PSS- pNIPAAM in the targeted region with exceptional resolution. First, the masks for photolithography were designed which determined all dimensions of the functional features. Two layers of masks were created for each device fabrication using the software Clewin 5.4: a gold-patterning mask and a polymer-patterning mask. The gold patterning mask consisted of the gold features printed in black ink, and the PEDOT:PSS-pNIPAAM patterning mask consisted of transparent features for the polymer and contact areas. The final design of the two masks were printed by a micro-photomask printing company, Micro Lithography Services Limited. All the clean-room fabrication steps were carried out by Zixuan Lu at the Nanoscience Centre, University of Cambridge. The fabrication steps began with cleaning of the 4-inch glass wafers, which acted as the substrate for fabrication of the MEAs. Next the photolithography patterning of the photoresist was carried out, followed by deposition of the titanium-gold layer using the LEV-Lesker electron-beam evaporator (Kurt J. Lesker Company GmbH, Germany). To peel off the metal layer to create the required pattern, the wafers were immerged into Technistrp® Ni555 solution (Technic, France) for 24 hrs. After this metal lift-off step, the designed gold pattern remained on the wafer. The next step involved coating the gold patterned surface with two layers of parylene dimer (Specialty Coating Systems, USA). This was carried out using the parylene coater, PDS 2010 Labcoater 2 (Specialty Coating Systems). The next step of the process was to apply photolithography techniques to form windows in the parylene (the parylene etching) based on the second mask design for electrode areas with photoresist. The Mask Aligner MA / BA6 (SUSS MicroTec SE, Germany) was used to exactly align the features of the second mask with the gold patterns on the wafer. After the etching windows were formed, the two layers of parylene at the etching window area were completely etched away for smart polymer deposition or contact purposes. The plasma etching system applied was Plasma Pro 80 RIE (Oxford instruments). Example 2-Polymer formulation and spin coating PEDOT:PSS synthesis: 31 μL of 98% (3-glycidoxypropyl) trimethoxysilane (GOPS) (Merck, UK) was pipetted into 1 mL of a dispersion of PEDOT:PSS (1.3 % wt) (CleviosTM PH1000, Heraeus, Ossila, UK), and then put in an ultrasound bath for 20 mins before use. PEDOT:PSS / pNIPAAm synthesis: 1 mL of a PEDOT:PSS dispersion was mixed with 225 mg pNIPAAm, 15 mg 2,2-dimethoxy-2-phenylacetophenone (DMPA) and 15 mg of N,N’- methylenebis(acrylamide) (mBAAm), all purchased from Merck, UK, with continuous stirring for 30 mins. Then, 31 μL of GOPS was pipetted to the prepolymer mixture and stirred at room temperature for 30mins continuously before use. The molar relationship between PEDOT:PSS and pNIPAAm polymers was calculated to be 1:27. To create layers, the desired prepolymer solution was spun for 30 s at 1500 rpm / s, followed by 30s at 3500 rpm / s and baked at 70ºC for 2 mins over the electrodes. After that, the layers were photopolymerised using a UV lamp, for 1 min at 365 nm. Then, hardbake was applied to the layers for 2 h at 120ºC. Once the polymerisation was complete, the layers were rinsed with ethanol and DI water to eliminate the non-polymerised monomers, resulting in a thin film of the polymeric layer on the substrate. Example 3 -Lectin Functionalisation experiments (Adsorption vs AuNP-based covalent coupling) Initial proof of concept experiments involved the functionalisation of lectins on polymer-coated indium tin oxide (ITO) platform by a simple physical adsorption. To achieve this, 100μLof 50μg / mL lectin ECA-FITC (ECA = Erythrina Cristagalli Lectin. FITC = Fluorescein Isothiocyanate) was loaded onto the polymer coated substrates and left standing overnight at 4ºC. Post incubation, the lectin solution was removed, and washed three times with PBS (phosphate-buffered saline). The adsorption of lectins to the polymer-coated surface was monitored optically by fluorescence microscopy. Next, 1% BSA (bovine serum albumin) solution was used to block the surface for 30mins prior to incubation with cells. Functionalisation of lectins on the bespoke MEA platform involved covalent coupling of lectins onto AuNPs. The final optimised protocol employed is as follows – First, the polymer coated MEAs were plasma treated (100 W, 2 min, Air) to introduce hydrophilicity. Next, 50μLof 0.1M positively charged Poly-L-Lysine (PLL) (Merck, USA) and incubated for 30mins to induce a net positive charge on the surface. Next, lipoic-acid coated 50nm AuNPs (Nanocomposix, UK) were loaded onto the PLL-coated MEAs at 50ug / mL for 4 hrs to allow electrostatic binding to the polymer surface. Subsequently, EDC and sulfo-NHS (Sigma Aldrich) were weighed separately (at 1:2 ratio) and mixed into 1mL PBS, pH 7.4 just before loading onto the AuNP- coated device for 30mins at room temperature. Immediately after the incubation, the surface was washed with PBS once and loaded with FITC-Lectin ECA at 50μg / mL concentration. The solution was left standing overnight to allow for functionalisation and washed with PBS three times. The successful functionalisation of lectins onto the AuNP-coated polymer surface was monitored by fluorescence microscopy of the optical window, and by EIS measurements, which indicated an average delta Rct shift of about ohms (when compared to the polymer baseline). This corresponds to the increase in impedance signal upon attachment of lectins to the electrode surface. Example 4 -Atomic Force Microscopy (AFM) experiments AFM images were made. The polymer PEDOT:PSS-pNIPAAM was spin-coated onto a glass cover slip based on the protocol stated above and used for AFM-based characterisation. The experiments were carried out in Scanasyst mode using ScanasystFluid+ probes (Bruker, USA) with a nominal spring constant of 0.7 N m–1 and a resonant frequency of 150 kHz. Images were recorded at scan speeds of 1.5 Hz and tip–sample interaction forces between 200 and 300pN.To resolve the morphology of the polymer layers, 5 × 5 μm scans were generated. Scans were taken of the polymer at room temperature first, which were then heated to 37ºC and scanned again, to evaluate the modified surface morphology. Measurements of the peak height of the polymers were performed by taking cross-sections across different areas of interest using the Nanoscope analysis software (Bruker, USA). Example 5 -Microscopy-based optical characterisation Brightfield and fluorescence microscopy was carried out as part of the optical characterisation of cell capture and release on different lectin-functionalised polymer substrates. All experiments were carried out using the Zeiss Axio Observer with the 20× / 0.8, (Plan- Apochromat, Zeiss) objective. The brightfield, EGFP (488nm), DAPI (359nm) and RFP (588nm) channels were used in most experiments together with fixed laser intensities. Example 6 -Microfluidic-based dynamic cell enrichment The mask design for the MEAs allowed for integration with a commercial microfluidic channel. Post lectin functionalisation and the parylene lift-off, a 60μm height microfluidic channel with an adhesive bottom (μ-Slide I Luer, IBIDI, Martinsried, Germany) was fixed onto a rectangular chip consisting of 4 MEAs. The fluidic channel was manually aligned such that the width of the channel (5mm) encompassed all the 6 electrodes of each MEA. The commercial design of the channel would cover the contact pads on the MEA preventing contacts for EIS measurements. To overcome this, one side of the microfluidic slide was trimmed using a drill (RS Pro, Bench Grinder, UK) to expose the area covering the contact pads to enable measurements. The experiments with cellular sample processing on the microfluidic channel were carried out using a syringe pump (Harvard Apparatus Pump 33 DDS, United States) with reverse flow. The fluidic flow involved the sample inlet on the right side of the channel connected to diluted cellular sample via the Luer inlet. The fluidic outlet located on the left of the channel was connected to a 20mL Syringe (BD Plastipak™ 20mL syringes with BD Luer-Lok™, Switzerland). The flow rate used during the experiments to achieve cell capture were optimised to be between 20-50ul / min. During the first 1hr of sample processing lower flowrates were used to provide the best chance for target cell capture, after which the dual-mode sensing operation was carried out. Following EIS measurements, the cellular samples were flowed through the channel at a higher flow rate to allow complete processing of the sample in a reasonable timeframe. Example 7 -Electrochemical measurements All electrochemical measurements were performed using a PalmSens4, equipped with PSTrace 5.6 software (PalmSens BV, Houten, The Netherlands). A three-electrode system was employed during all EIS and CV measurements. The platinum served as an auxiliary electrode and Ag / AgCl served as a reference electrode in 1X PBS electrolyte containing 5 mM [Fe (CN)6]3− / 4− and 0.1 M potassium chloride (KCl) at a 0.1 V potential over the frequency range from 100 kHz to 1 Hz. The working parameters of Cyclic Voltammetry (CV) included a scan rate of 0.1V / s and the applied potential range was between −0.3 and 0.7 V. The voltammograms obtained were analysed using the PSTrace software. For EIS measurements carried out on indium tin oxide (ITO) substrates, 8mm plastic well was glued onto a localised circular area (6mm diameter) containing a thin film of the polymer PEDOT: pNIPAAM. EIS measurements were carried out as stated above, with the localised area within the well acting as the working electrode (WE). Pogo pins were used to contact the conducting ITO substrate outside of the well, which completed the circuit for EIS measurements. EIS measurements were similarly carried out on the polymer-coated MEA platform using a fixed plastic well covering the 6 electrode areas. Here, the contact pads designed with a specific spacing was used to align with that of the pogo pins to make contact across the electrodes in the MEA at once. However, each electrode was queried separately to obtain independent electrode measurements. For EIS measurements carried out on MEAs within the fluidic channel, the PBS electrolyte containing 5 mM [Fe (CN)6]3− / 4− was passed through the channel to completely cover all the MEAs along with the inlet / outlet. The two MEAs placed in the middle of the channel were queried by EIS. The RE and the CE were dipped on either the inlet or the outlet, depending on its proximity to the MEA being measured. The pogo pins were used similarly as stated above to make contact with the exposed contact pads to act as the WE and complete the circuit for EIS measurements. Example 8 -Clinical Validation Experiments Cytosponge sample collection: All patients were recruited following informed consent from Addenbrooke’s Hospital (University of Cambridge, UK), with prior peer-review and approval by the ethical committee (REC: 10-H03087-1). Cytospin™ of released cells: Different aliquots of cell populations from the clinical validation experiment were collected, kept on ice and centrifuged at 400 x g (4°C) for 5mins, and washed with 1mL ice-cold PBS. Cells were then centrifuged again (400 x g, 4°C) and fixed with 500μL4% paraformaldehyde (PFA) for 20 mins on ice. Cells were then centrifuged (400 x g, 4°C), to remove the PFA and cells were washed again with 1mL PBS and resuspended in PBS. The resuspended fixed cells were used with the Cytospin™ centrifuge with cytofunnels and cytoclips (Thermo Scientific, USA) for transferring them onto microscope slides. The resuspension volume was based on the cell counts, obtained via standard trypan-blue haemocytometer-based assay utilising the disposable C-slide haemocytometer (Curiosis, South Korea). Based on prior optimisation results, 200μLtubes containing fewer than 80,000 cells were used further for Cytospin-based transfer onto the glass slide. Cytospin receptacles were assembled, and the suspension of fixed cells were added to the cytospin slide centrifuge (Cytospin 3, Shandon Southern Instruments, Inc, Sewickley, PA, USA). Cells were centrifuged at 500rpm for 5 mins with medium acceleration onto SuperFrost Ultra Plus™ adhesion slides (Thermo Fisher Scientific Inc., USA) before being dried face up in the dark overnight at room temperature (22°C). Each sample was split and transferred onto two glass slides to carry out H&E staining and immunofluorescence-based evaluation. Haematoxylin & Eosin (H&E) staining of Cytospin slides: One set of Cytospin slides were stained with H&E to assess the morphology of the cells, and qualitatively assess enrichment. The staining was carried out as per a previously published protocol (Crowley, L. C., et. al., Cold Spring Harb Protoc 2016, 773–777 (2016)). Briefly, the slides were first washed with tap water (1min), and then dipped in a well containing Gill’s II haematoxylin (VWR, USA, code: 1.05175.0500) which stains the nuclei (for 2mins). The slides were then washed again with tap water and dipped in 1% aqueous eosin (eosin powder VWR code 34197) made up with 1% calcium carbonate (Sigma-Aldrich, Merck, USA, code: C4830) which stains cytoplasm. Following this, the slides were washed with tap water and dipped in series of wells containing ethanol prepared at 70%, 95% and two wells of 100%, in order to remove excess stain and reveal clear morphological features. Finally, the slides were dipped in two wells of Histoclear (1-3mins), and then mounted with a coverslip. IF (immunofluorescence) staining of Cytospin Slides: To further verify the columnar cell identity of the enriched cells, a second set of Cytospin slides were stained with the MUC5AC antibody (MA1-21907, Thermo Fisher,USA) using the manufacturer’s protocol. Briefly, the slides were treated with 100% methanol for permeabilisation for 30mins. Next the slides were blocked for non-specific binding with a blocking buffer (Dako Agilent, USA, code: X0909) for 30min at room temperature. Following this, the slides were incubated with the primary antibody for MUC5AC overnight at 4°C. The slides were then washed and incubated with a corresponding secondary antibody with a FITC fluorophore tag for 2hrs at room temperature (RT). The slides were subsequently washed in PBS and mounted using mounting medium with DAPI (Vectashield plus antifade, 2B Scientific, UK). MUC5AC staining was imaged the following day using ZEISS Axio Observer microscope. On-chip IF staining directly on MEAs: To perform on-chip IF staining of captured cells, an aliquot of the Cytosponge sampling was loaded onto a parallel microfluidic MEA chip previously functionalised with lectin ECA. After 1hr of sample processing, the 4% PFA was loaded completely into the channel and incubated for 30mins to fix the cells for staining. Following fixation, the same IF protocol as stated above was carried out for staining the captured cells with MUC5AC. RT-qPCR based evaluation of RNA from clinical sample aliquots: Nucleic acid extraction and RT-qPCR experiments were carried out. RNA was extracted from the cells retrieved from the bioelectronic enrichment platform using the Qiagen AllPrep DNA / RNA Micro kit (Qiagen UK, Cat. No. / ID: 80284) following the manufacturer’s protocol. 120 ng of RNA per sample was used in reverse transcription to generate cDNA using Qiagen QuantiTect® Reverse Transcription kit (Qiagen UK, Cat. No. / ID: 205311) as per the manufacturer’s protocol. qPCR was performed using Qiagen QuantiNova Probe PCR kit (Qiagen UK, Cat. No. / ID: 208252) following the manufacturer’s protocol.0.8 ng cDNA was used in one qPCR reaction assuming 100% conversion efficiency, whereby 120 ng cDNA was synthesised from 120 ng RNA. For each sample, three technical replicates were performed for each of the following genes using the relevant primer set: MUC5AC, MUC2, CDX2, CDH17, TFF3, KRT8. GAPDH and β2M were used as housekeeping genes. The relative gene expression was calculated using the ΔΔCT method263. All primers used are stated in Table 1. Table 1. Primer sets for RT-qPCR studies.

[0002] Results: Evaluating the hybrid polymer PEDOT: pNIPAAm for lectin-based cell enrichment – proof of concept studies A previous study demonstrated PEDOT:PSS-pNIPAAM to be an effective hybrid polymer for electrochemical monitoring of cell capture and their heat triggered release. However, cell capture was achieved by a non-specific adhesion of cells via fibronectin adsorbed onto the surface of the hybrid polymer. In this results section, the inventors show initial proof of concept experiments to demonstrate the applicability of PEDOT:PSS-pNIPAAM for lectin-based specific cell capture and thermal release. Figure 5 depicts the proposed schematic of the mechanism of cell capture and release based on heat-induced conformational change of the smart polymer. First, optical evaluation of the proposed mechanism was carried out using lectin-functionalised polymers coated onto a glass slides followed by the evaluation of a dual-mode optical- electrochemical platform for monitoring the capture-release mechanism on lectin- functionalised ITO (Indium tin oxides) substrates. Optical characterisation of lectin ECA coated polymer on a glass slide To visualise the lectin-based cell capture mechanism, ECA lectin was functionalised onto polymer coated glass slide. While not represented below, appropriate negative and positive controls were used to validate this experiment involving functionalisation of a blocking agent bovine serum albumin (BSA) and fibronectin to facilitate adhesion respectively. The cell line TIF LifeAct-RFP was used for all proof-of-concept experiments to evaluate cell capture on lectin-functionalised polymer. To assess the influence of pNIPAAM on thermal-induced cell detachment, PEDOT:PSS (conductive polymer) and PEDOT:PSS-pNIPAAM (smart polymer comprising conducting and thermo-responsive polymer respectively), were spin-coated independently onto glass slides. Next, FITC-tagged lectin ECA was adsorbed onto both polymers overnight, which was followed by a 30min incubation with BSA. The slide surface was then washed with PBS and imaged by brightfield / fluorescence microscopy. The functionalisation of lectins on the polymer surface was confirmed by observation of green fluorescence on glass slides. Next, about 500,000 TIF cells were incubated on both slides for 1hr, washed and imaged by brightfield and fluorescence microscopy. It could be observed that the RFP-tagged TIF cells were attached on both types of polymer- coated slides (Figure 6). The mechanism for cell capture observed here indicates a specific cell surface marker-based attachment to lectins present on the polymer surface. This was further confirmed using a hybrid polymer coated slide with no lectin coating (negative control), where no attached cells were observed after 1hr of incubation. To test the effect of heat on each polymer and their attached cells, both PEDOT:PSS and PEDOT:PSS-pNIPAAM coated slides were kept on a hot plate set to 37°C for 15mins, washed with PBS and imaged. It was evident from optical microscopy that cells attached on the PEDOT:PSS slide maintained their attachment to the lectin-coated surface (Figure 6a), while most of the cells in the PEDOT:PSS-pNIPAAM coated slide were observed to be easily washed away (Figure 6b). In terms of the mechanism for cell detachment, it was previously demonstrated to be based on the release of the adsorbed fibronectin by thermal actuation of the polymer which becomes more hydrophobic246. Since lectins are similarly just adsorbed onto the polymer surface like fibronectin, it is believed that a similar mechanism is at play here. The ECA lectin conjugated with FITC (indicated by green fluorescence in the FITC channel in Figure 6) is likely being released due to the thermal actuation of the polymer. This is also evidenced by the decrease in the FITC fluorescence intensity in the post-heated hybrid polymers. Example 9 -Dual-mode characterisation of lectin ECA coated polymer on indium tin oxide substrates Next, the inventors sought to test the lectin-based selective cell capture-release mechanism on a dual-mode sensing platform that could allow for simultaneous optical and electrochemical monitoring. This provides several advantages – the electrochemical monitoring offers a direct, quantitative method to assess the amount of target cells bound to the lectins; the ability for optical characterisation presents an avenue for further antibody-based staining assessment to confirm the presence of cell-specific markers; and finally, it presents a mutual confirmation of captured target cells adding to the robustness of the sensor system thus improving clinical translation. The simplest way to test the dual-mode sensing approach was using indium tin oxide (ITO) coated substrates, which presents as both an optically transparent and a conducting substrate. Figure 7 demonstrates the proof of concept dual-mode sensing platform using ITOs for lectin- based cell capture / release on the hybrid polymer. 20mm x 20mm square glass substrates coated with a conducting ITO material were obtained from Ossila, UK and spin-coated with the polymer PEDOT:PSS-pNIPAAM. A mask of specific dimension (6mm diameter circle) was used to localise / standardise the area of measurement. Lectin ECA adsorption onto the polymer coated ITO, cell capture, and thermal release was carried out as stated previously (Figure 7a). However, due to the transparent and conductive nature of the substrate, each step of the experiment was monitored both optically (by brightfield microscopy) and electrochemically (by EIS and CV). Figure 7b depicts the configuration of the ITO-based platform, wherein a plastic well of 8mm diameter is glued onto the polymer-coated ITO. Brightfield microscopy of the well area (in PBS) was carried out as an optical confirmation of lectin-based capture of TIF cells, and their subsequent detachment from the surface when subjected to heat (at 37°C) was quantified. The EC measurements in the well were carried out in PBS containing 5mM FeCN redox probe using a three-electrode system. Figure 7c-f presents the various representations of the EC data collected. The raw electrochemical impedance data is represented by the Nyquist and Bode plots in Figure 7c, and Figure 7d respectively. Impedance-based assessment indicated an expected trend of increased signal at the low frequencies upon cell capture, which was diminished post-release. This shift at low frequencies in a faradaic regime indicates the interactions of the redox probe with the electroactive surface. Capture of cells on the surface presents a blocking effect, which leads to an increase in the magnitude of impedance. Similarly, the detachment of cells by thermal actuation led to a decrease in impedance indicated by a shift in the lower frequency. This trend was maintained across the phase and magnitude Bode plots and the Nyquist plot. The electrochemical monitoring of cell capture-release was further made clear by the quantification of the Nyquist plot using a simplified Randall’s circuit (Figure 7e) which represents the charge transfer resistance of the ITO electrode surface. The quantified Rct values for the lectin functionalised baseline, cell capture, and post-release was 3.7kΩ, 13.9kΩ and 9.7kΩ, respectively. This provides greater insights into the proposed mechanism indicating an increasing and then decreasing trend for impedance measurements for the capture and release of cells respectively. Finally, cyclic voltammetry (CV) was also investigated as a potential strategy for EC evaluation of the capture-release mechanism. Figure 7f presents the corresponding CV measurements where the quantified oxidation peak heights for the lectin functionalised baseline, cell capture, and post-release were 450μA, 200μA, and 250μA respectively. This provides further evidence for the blocking-effect on charge transfer after cell capture which tapers off post-release, thus corroborating the EIS measurements. These findings present the first proof of concept for a dual-mode sensing platform for lectin-based capture and thermal release of cells on the hybrid polymer PEDOT:PSS-pNIPAAM. Post thermal release, some isolated cells and cellular debris were observed to still be attached on the ITO surface which was further corroborated by the EIS signal for ‘cells released’ showing a signal higher than the lectin baseline. This suggests that there is a need to assess the release efficiency from the platform. The cell capture saturation, viability of released cells and the thermal detachment efficiency of the platform are discussed in further detail below. It is noted that temperature is known to alter electrochemical measurements. This means that measurements taken at room temperature cannot be directly compared with measurements taken at 37°C. To avoid this, all electrochemical measurements post thermal actuation of the polymer were carried out after leaving the device at room temperature for 30mins, to allow it to recalibrate to the temperature at which the baseline and cell capture measurements were taken (22°C). At this stage, the dynamic polymer’s conformation was expected to return to its original (hydrophilic) state allowing for a more like-for-like comparison of signals collected previously. This Example illustrates the ability of the platform described herein to selectively capture cells from complex matrices. This makes it suitable for liquid biopsy workflows, including rare cell enrichment applications. Example 10-Selective lectin-based cell capture from endoscopic brushings on polymer- ITO substrates All data presented so far is based on capture of TIF cells, which were identified to be specific to the ECA lectin. ECA demonstrates specificity for only to the columnar cells arising from gastric or Barrett’s tissue, while no specific binding was observed in normal squamous oesophagus tissue (NE tissue). To prove the specificity of cell capture, and to assess the clinical applicability of the polymer-based platform, endoscopic brushings from localised tissues (NE and BE) were tested on the lectin-functionalised polymer-ITO platform. Figure 8 demonstrates the selectivity of cell capture, and the verification of the dual-mode sensing platform for lectin-based cell capture / release on the hybrid polymer. Cells collected from endoscopic brushings of NE and BE tissue were washed with PBS and counted (using a haemocytometer) before loading onto the ITO platform (Figure 8a, b). Equal number of cells for each brushing were incubated on the lectin-functionalised polymer-ITO platform (Figure 8c, f). From the optical characterisation of the well after 1hr of incubation, it could be seen that some of the BE cells were bound to the ITO surface (Figure 8d), while none of the NE cells were attached (Figure 8g). This selective lectin-based binding corroborates previous biopsy staining results of lectin ECA. To perform the electrochemical characterisation, simplified Randall’s circuit-based quantification of Nyquist data was chosen since it clearly depicts the trend of EIS signal observed during the experiment. Figure 8e, h, presents quantification of impedance signals from three lectin functionalised ITOs used to test each brushing sample. Complementing the optical findings, the EIS measurements showed a modest increase in the mean Rct value only for the BE brushing sample with an increase of 1.9kΩ, relative to the lectin functionalised baseline. Interestingly, a small decrease in the mean Rct was observed in the case of NE brushing sample of 0.6kΩ. Owing to the relatively high rate of variability observed in the replicate devices, these shifts were not found to be significantly different. One potential reason for the low shifts in impedance measurements could be the relatively low number of cells loaded. For this study, the endoscopic brushings from each tissue yielded about 600,000 cells which were matched to have equal numbers of BE and NE cells (by counting on the Vicell cell counter) and evenly split across three devices. Therefore, the resulting number of cells incubated in this study was about 1 / 3rd that of the previous study with the cell line, which was more readily available. However, this also reveals the lack of sensitivity of the ITO-based platform for monitoring cell captured when the loading numbers are lower than 500,000 (as in the case of the previous experiment). Example 11- Design and fabrication of a bespoke microelectrode array (MEA) Bespoke masks for gold patterning and polymer patterning were designed, printed, and used for fabrication. Figure 9a depicts the gold patterned mask of a 4-inch wafer consisting of multiple 24 MEAs. Each MEA region contained 6 circular electrodes with a diameter of 1.2mm. These dimensions were chosen based on the previous study employing PEDOT:PSS- pNIPAAM. Each electrode was connected to its respective square contact pad and assigned a number. The contact pads used were 1mm x 1mm, with the separation between each pad also being 1mm. This was optimised to align with gold pogo pins, which were used to make contact with the pads. The distance between the array of electrodes and the contact pads was optimised to either glue a plastic well with 8mm diameter and 0.5mm thickness, or a microfluidic channel with 5mm channel width. Out of the 6 circular electrodes, three electrodes (numbers 1,3,5) were integrated with a transparent circular window at its core with a diameter of 500μm. This new design was implemented to allow for dual-mode optical-electrochemical monitoring of cell capture-release. The other three electrodes (numbers 2,4,6) were regular circular electrodes of the same diameter, but without the optical window at its core. In addition to the 6 circular electrodes, the right side of each array design consisted of a mask for an OECT array with 3 channels and a large rectangular planar gate. This OECT array was not used in the context of this application but integrated for different projects within the group. Two types of gold electrodes, one with and without an optical window at its core was included in the design, to assess the possible alteration of EIS signal due to the presence of the optical window. Initial characterisation experiments of the baseline measurements for each type of electrodes revealed no major differences in the Rct values (not represented here). This indicated that future mask designs could consist of MEAs with all electrodes having the optical window, to provide a better optical representation of the captured cells. Figure 9b depicts the second mask design of etching windows for the hybrid polymer film deposition on each electrode within the array. This mask was used to pattern two layers of parylene – a widely employed biocompatible polymer coating used as an insulating material. The top layer of the parylene could be lifted off after spin coating of the conducting polymer to create patterned conductive polymer regions at the exposed sites. The dimension of the exposed etching windows was designed to match that of the gold circular electrode, which determine the shape and size of the conductive polymer layer. These polymer-coated regions are directly used for the impedance-based analysis. The etching window at the contact pads was added to exposing the gold contacts to connect to the pogo pin, as the working electrode. The implementation of the parylene layer was also used to aid in the localised functionalisation of lectins on the electrode areas, which is further detailed in the section below. Polymer spin coating and functionalisation Figure 3a,b depicts an MEA chip consisting of the six circular gold electrode array that was cut and used for each experiment. The inset further depicts the design of a single bare gold electrode with the new optical window design at its core. In this application, the MEA was spin- coated with the hybrid conducting and thermo-responsive polymer PEDOT:PSS-pNIPAAM, to create a platform for bioelectronic cell enrichment. The thin film coating of the PEDOT:PSS- pNIPAAM provided a transparent polymeric layer at the core of the electrode, allowing for optical monitoring of surface binding events. The conductive property of the polymer further ensured that the entire circular surface of the electrode including the transparent core, were a connected electroactive surface. The additional thermo-responsive property of the coated polymer was used to achieve the thermal release of captured cells on the electrode. To begin the MEA chip preparation, the surface of the chip was cleaned and treated with O2 plasma to increase the hydrophilicity on the surface and enable efficient spin coating. Upon spin coating of the hybrid polymer, the surface was UV cross linked and hot-baked. The patterned parylene layer in addition to aiding in localised spin-coating of the polymer layer, presented an opportunity to also achieve localised lectin functionalisation exclusively on the electrode regions. Figure 3c depicts the step-by-step process starting with the spin-coating of the hybrid polymer PEDOT:PSS-pNIPAAM onto the entire MEA chip. Before lifting off this polymer spin-coated parylene surface, the inventors sought to make use of this layer to further achieve localised functionalisation of lectins onto the polymer-coated gold electrode surface. A series of functionalisation steps was carried out to attach lectin ECA onto the chip’s surface (Section “Functionalisation steps” above). After the successful attachment of FITC-lectins onto the surface, confirmed as before using fluorescent microscopy, the upper parylene layer was peeled off, to reveal six patterned gold electrode surfaces coated with the polymer PEDOT:PSS-pNIPAAM and now functionalised with lectins. This localised functionalisation of lectins enabled the specific attachment of cells exclusively onto the electroactive surface, thus enhancing the sensitivity of measurements. Alternatively, if the parylene layer was to be lifted off prior to the functionalisation procedure, it would limit the cell detachment efficiency since cells attached to the lectins outside of the hybrid polymer coated electrode regions, would fail to be released by thermal actuation. In doing so, the efficiency of a potential enrichment application would be significantly enhanced, as detailed below. Figure 3d demonstrates the three-electrode system employed for EC monitoring. A plastic well of 8mm was glued onto the MEA chip containing lectin functionalised electrodes. The patterned gold contact pads with specific labels were used to contact pogo pins and act as the WE. A single electrode was connected at each time to take individual electrode measurements. External Ag / AgCl reference and platinum counter electrodes were dipped into the electrolyte solution within the plastic wells to complete the circuit. A representative faradaic mode EIS circuit was used to quantify the EIS Nyquist data as stated previously. Functionalisation steps Previous experiments on lectin-based cell capture were carried out through physical adsorption of lectins onto the polymer surface This was initially attempted on the fabricated MEA chip, as a simple method of functionalisation. However, in many instances, the parylene lift off procedure followed by stringent wash steps, led to a heterogenous and patchy coating of lectins. Previous experiments testing endoscopic brushing samples on polymer adsorbed with lectins, also indicated the inability of this approach to bind clusters of cells or cell groups which are clinically relevant for diagnosis. To overcome these limitations, a new functionalisation strategy was optimised and implemented to achieve homogenous and efficient coating of lectin ECA on the polymer surface. AuNPS have been previously used to increase surface area for better functionalisation and thus lead to improved sensitivity. Shown in Figure 10a are different steps performed to design conductive surface that enable controllable attachment of lectins to conductive polymer through Au NPs. Briefly, the polymer PEDOT:PSS-pNIPAAM was spin coated onto the MEA as previously stated, with the parylene coat still on (step 1). The polymer coated MEA was plasma treated to introduce a net negative charge on the surface to enable electrostatic attachment of positively charged PLL (step 3). Next, this surface was coated with negatively charged AuNP-lipoic acid (with a COOH group)(step 3) and subsequently modified with lectin ECA through amide bond formation using EDC-NHS strategy (step 4). Characterisation of each step of the functionalisation process was carried out using different modalities represented in Figure 10b. Before polymer coating, a brightfield image of the bare Au electrode was obtained. Spin coated PEDOT:PSS-pNIPAAM, polymer was characterised by AFM, with rough polymer surface clearly shown. SEM was employed to characterise AuNPs attachment, followed by fluorescence microscopy after the attachment of FITC-tagged lectin ECA and the binding of TIF Lifeact-RFP cells. Figure 10c and d depict the raw EIS Nyquist and the corresponding quantification of the Rct values respectively for each step of the functionalisation process. The quantification of the Rct, plotted in Figure 10d, presents the bar plot comparison with error bars representing the standard deviation in signal from 3 MEA arrays consisting of 6 electrodes each (n=18). First, an increase in impedance signal is observed after spin-coating of the hybrid polymer. The conducting polymer PEDOT:PSS is normally expected to enhance the volumetric capacitance of the electrode area and thus decrease the Rct value. However, since the hybrid polymer consists of chains of the non-conducting pNIPAAM, this is expected to result in a relatively higher impedance than the bare gold electrode. Following the functionalisation protocol, after incorporation of AuNP-lipoic acid on the polymer, the Rct value is observed to drop drastically below that of the bare gold electrode baseline. This indicates a strong conductive influence of the charged metal nanoparticle (see for example Chinnadayyala, S. R. et al. App Sci 2019, Vol.9, Page 3269, 326 (2019).) The covalent coupling of lectin ECA via EDC-NHS chemistry, as expected, result in an increase in the Rct value, further demonstrating the high-sensitivity of the electrodes to monitor surface binding events of biomolecules. Finally, the capture of TIF lifeact-RFP cells produced a drastic increase in the impedance signal owing to the ‘blocking’ of the electrode surface by relatively large cell monolayer. A one-way ANOVA was performed pairwise across each group, revealing statistically significant differences among all conditions, with the exception of the comparison between the polymer and lectin baselines. The dual-mode detection capability shown in this Example enables sensitive identification of biological targets, with potential applications in pathogen detection where combined optical / electrochemical verification. Example 12 - Mechanistic studies of functionalisation, actuation After the cells were captured on lectin functionalised MEA, the next step of the bioelectronic enrichment platform was their thermal-induced release to retrieve the cells for downstream molecular assays. The release mechanism previously stated on the ITO platform was clear. The adsorbed lectins on the polymer surface were released by thermal actuation of the polymer, leading to the cells being detached. With the new functionalisation strategy stated in the previous section, it was important to characterise the mechanism of detachment of the cells. Figure 11a shows a 3D rendering of AFM data of the polymer PEDOT:PSS-pNIPAAM at room temperature and at a temperature of 37°C. The conformational change that the polymer undergoes from being hydrophilic at RT to its transition to becoming hydrophobic at temperatures above its LCST of 32C was clearly observed by AFM. The ‘pillar-like’ polymer chains observed at RT seemed to relatively flatten out upon heat exposure. Line plots depicting the peak height presented in Figure 11b are based on the 2D AFM data of the hybrid polymer at RT and at 37C. The vertical peak height of the pillar-like polymer chains was estimated to be 30.8nm, while the quantification of vertical peak height of the same polymer heated to 37°C was lowered to about half its original value at 14.9nm. This further confirms the conformational changes that the polymer undergoes after heat exposure. Based on these findings, it was confirmed that the detachment of cells attached to the polymer surface was based on the thermal actuation of the polymer leading to a significant conformational change. The specific layer of functionalisation being detached, leading to cell release was still however unclear. Figure 11c depicts the mechanistic study experiments carried out wherein the polymer-coated MEA was subjected to heat after each functionalisation step. EIS measurements were taken after each step in the functionalisation protocol before and after heat exposure to assess its influence on the measurement (Figure 11d). Based on the heat-exposure data, it was concluded that neither the AuNPs nor the lectins were being entirely released from the surface, as this would have led to a marked changes in impedance values, or a significant decrease in fluorescent intensity, neither of which was observed. However, upon heating the surface with cells attached to the surface, the heat actuated polymer conformation change seemed to still release the attached cells (in other words, the target cells were selectively released). Example 13 -Sensitivity Establishing the limit of detection (LoD) and the dynamic range of sensing is paramount for any biosensor platform, since it directly defines the scope for its clinical application. To find the limit of EC detection of captured cells on the MEA, EIS signals were obtained from increasing number of cells bound on lectin functionalised electrodes. Based on previous results of Cytosponge pathology image analysis, an adequate sample is estimated to contain anywhere from a total of 1-4 million cells. Within this sample, the columnar cells, which are the cell group of interest for detection of BE, make up only about 1- 10% of the sampling (for long segment BE). This puts an estimated required dynamic range of the electrochemical sensor to be between 10,000 to 400,000 cells. Since the designed MEA consists of 6 electrode areas which are coated with the polymer and functionalised with lectin ECA. The inventors assessed the impedance of individual electrode measurements independently which collectively provided the estimate of total cell capture sensitivity of each MEA. Figure 12a presents the impedance analysis of an MEA chip indicating an even spread of cells attached across the 6 electrodes, with limited variability observed between the middle (electrode number 3, 4) and outer electrodes (numbers 1,2,5,6). This relatively limited variability presents further evidence for the homogeneity of lectin functionalisation across the MEA chip and presents a simple analytical pathway for estimating the total cells attached on each chip. An increasing number of TIF cells were loaded onto the lectin functionalised MEA starting from 1,500 cells until 300,000 cells. Each incremental cell loading was incubated for 1hr before imaging / EIS measurements. Figure 12b represents the results of EIS evaluation conducted on 4 separate lectin functionalised MEA chips, each consisting of 6 electrodes. Based on the assumption of homogenous cell binding across the 6 electrodes, the average impedance signal of each MEA chip was matched to one sixth the total number of cells loaded on each array. The error bars denote the relatively low inter-chip variability which tends to increase with incremental cell loading numbers. Figure 12c shows representative images of electrode 3 (E3) from an MEA chip at each step of the experiment corresponding to cell loading for each electrode. As expected, an increasing number of cells are observed to attach to the electrode with each incremental cell loading step. The corresponding cumulative Rct values obtained from EIS measurements taken at each cell loading step revealed a linear relation between the delta Rct values and the cell loading numbers (Figure 12d). Using linear regression analysis, a limit of detection of 140 cells / electrode and a linear dynamic range between 200 to 40,000 cells / electrode was established. This provides the final MEAs linear dynamic range of sensing to be between 1,200 to 240,000 cells. These results indicate the potential applicability of the bioelectronic enrichment platform for Cytosponge sampling. To account for potential high degree of columnar cell groups present in the sampling (which is often observed in case of long segment BE cases), simply incorporating multiple arrays to process the sampling was envisaged and implemented using a microfluidic channel. Example 14 -Thermal cell detachment efficiency The efficiency of the developed enrichment platform relies on its capacity to recover cells captured on the polymer surface, a process that is not guaranteed to be 100%. This example evaluates the performance of the developed MEA-based platform was the effectiveness of thermally releasing captured cells from the surface. First, a massive excess of about 650,000 cells (counted by ViCell counter, Thermo Fisher, USA) were incubated on the lectin functionalised MEA chip for 1hr to establish the limits of the platform in terms of cell capture. After the incubation step, the unbound, washed cells were collected and counted. The counts indicated the saturation of cell capture on the MEA chip being ~250,000 cells. This was calculated based on the washed unbound cell counts being ~400,000. Based on these results, the efficiency of cell release by thermal actuation was next sought to be properly assessed. In a simple experiment, depicted in Figure 13a, a lectin functionalised MEA chip was incubated with a fixed number of cells (200,000 cells) that were previously counted by the ViCell cell counter. This experiment also provided an assessment of cell viability, which was observed to be 98% before loading onto the surface. After the incubation, unbound cells were washed and collected for counting. The captured cells were imaged by fluorescence microscopy to confirm cell attachment (Figure 13a, inset). Next, EIS measurements were taken which showed a significant increase in impedance values indicating the ‘blocking effect’ of cells attached on the surface. Once the successful capture of cells was monitored and confirmed both optically and electrochemically, the platform was subjected to heat by using a pre-heated PBS wash buffer (kept at 37°C). This approach was tested as an alternative to hot plate since it reduces the thermal stress subjected to the captured cells, and allows for a simpler, and more easily operable method of cell detachment. The detached cells were then collected by washing the MEA surface with the pre-heated PBS and stored for counting and viability assessment. The cell detachment was again confirmed both optically and electrochemically, wherein the impedance values reduced to a significant amount in comparison to that observed upon cell capture. The collected washed unbound cells and the detached captured cells were counted on the ViCell counter. Interestingly, no unbound washed cells were detectable which indicates a near 100% cell capture. This could be because the number of cells loaded were within the saturation established for cell capture on the lectin-functionalised MEA chip. Replicate experimental data of the detached cell counts from 3 different MEAs indicated an estimate range of the efficiency of thermal release to be between 78-85%. This points to the fact that a minority of the captured cells remain attached to the polymer surface even after thermal exposure. This could potentially be further improved by additional cycles of thermal actuation of the polymer after it reverts to room temperature. Assessment of cell viability of the detached cells using the ViCell counter always indicated a viability of about 95%. This presents an important milestone performance for the platform since it indicates the applicability of the released cells for a numerous downstream cellular and molecular characterisation assay. Additionally, this confirms the biocompatibility of FeCN as a redox probe to perform EIS measurements. The EIS measurements of thermal release indicate a drop in impedance as previously observed in the case of ITOs (Figure 13b, c, d). Comparing the Rct values post-release with that of cells captured, an average drop of 65.2% was observed. This new Rct value is not expected to provide an accurate estimate of the number of cells still attached on the surface, since cellular debris and altered conformation state of the polymer is known to affect the signal. The drop in Rct is meant to be used as a quality control check and could be used to quickly ascertain if the platform if performing to its intended specifications (i.e achieve about 82% release of captured cells). Example 15-Target specific cell capture in mixed sampling Lectin-based cell specific capture was first demonstrated on the ITO platform stated above. Pure cell groups obtained from endoscopic brushings from each tissue type were used to assess the platform’s specificity. Since encountering a pure cell population from an adequate Cytosponge sampling is unlikely, it was sought to test the MEA platform’s specificity, under heterogenous sampling conditions. Figure 14a represents the cell-line spike-in experiment performed to evaluate the platform’s selectivity. NE cells were first collected from a patient endoscopic brushing sample, which were then evenly split into three vials (Figure 14b). Instead of using endoscopic brushings of BE tissue, the inventors sought to first test the selectivity in a more controlled manner, and as such chose to spike-in TIF cells (as a positive control cell known to bind specifically to lectin ECA). One of the vials was spiked in with 25% of TIF cells, the next vial was spiked in with 5% of TIF cells and the final vial contained a pure population of NE cells (as a non-specific negative control). These spike-ins were chosen to represent the low and high end of the dynamic sensing range established from previous sensitivity experiments. Each vial was incubated on a separate lectin-functionalised MEA. Figure 14c shows the optical characterisation of cell capture within the transparent window of the electrode after 1hr incubation with the sample vial and a wash step to remove unbound cells. Representative electrode image is provided from each MEA which were incubated with different ratios of cell mixtures. Figure 14d presents the corresponding Rct values obtained from EIS measurements taken from each of the MEAs. The EIS measurements taken from each individual electrode was measured, and the average Rct value obtained for each array was plotted, with an error bar representing the standard deviation from the mean of 6 electrodes. It could be observed that the MEA incubated with a pure sample of NE cells, showed no indication of cell attachment on any of the electrodes, thus corroborating the findings within the optical window in Figure 14c (where no cells were attached). The MEA incubated with 5% of TIF cells indicated a slight shift in Rct values, while no cells were found within the optical window. This highlights the sensitivity of the electrochemical platform to detect a small minority of target cells within a large background of healthy oesophageal squamous cells. Finally, the MEA incubated with 25% TIF cells showed a drastic shift in Rct, expectedly indicating relatively higher cell attachment on the electrodes within the MEA. However, a higher degree of variability of cell attachment was also observed across the 3 MEA chips, relative to measurements made with pure cell populations. The spike-in ratios also fall within the realm of a potential Cytosponge sampling constitution which presents a preview of the platform’s performance under real-world conditions. These results demonstrate the platform’s ability to capture target cells when mixed in with high degree of background healthy cells. However, this was still a controlled experiment wherein single cell suspension of ECA lectin specific TIF cells were loaded at known ratios. The next section presents the clinical testing of the bioelectronic enrichment platform on patient Cytosponge sampling representing a proof-of-concept evaluation of the platform under real- world conditions. . Clinical validation of the bioelectronic enrichment platform This section presents the results of testing the bioelectronic enrichment platform on Cytosponge sampling from patients with BE and a healthy control. For clinical sample testing, a fluidic strategy was employed where several MEAs in series were attached to a single channel fluidic system with an inlet and outlet to increase the surface area for cell capture. Several layers of evidence were collected during each sample testing, starting with EC quantification of captured cells, their optical characterisation through the optical window, followed by a series of downstream molecular characterisation assays post cell detachment from the MEA platform. Example 16- Fluidic-based dynamic cell enrichment on Cytosponge sampling Previous sections reported the performance of the bioelectronic enrichment platform using replicate data of one MEA chip consisting of 6 lectin-functionalised electrodes (see example 15). To process an entire Cytosponge sampling with 1 to 4 million cells, there was a need to increase the surface area by incorporating multiple MEAs. Doing so would shorten the time required for sample processing, owing to the increased surface area for cell capture provided by 12 electrodes functionalised with lectins. The μ-Slide I Luer fluidic channel was obtained from IBIDI, GmbH and attached on a chip consisting of 4 MEAs (Figure 15) through the provided adhesive side. However, only the two MEAs in the middle of the channel were functionalised with lectins. This was done to limit the effects of fluidic stress on MEAs closer to the inlet and outlet. Also, taking EIS measurements of selected electrodes from two MEAs was deemed to be more feasible, to enable a reasonable assay time. Before implementing the fluidic-based enrichment system on the MEAs, preliminary testing of dynamic cell capture in a fluidic channel was tested on lectin-coated polymer-glass slides. These optimisation experiments helped to ascertain the flow rate and sample dilution required to effectively process the Cytosponge sampling. Additionally, the advantages of the AuNP- based functionalisation strategy, compared to a physical adsorption-based approach was further observed with efficient dynamic cell capture in a fluidic system. For clinical testing of the bioelectronic enrichment platform, Cytosponge samples were processed and flowed through the fluidic MEA platform previously functionalised with the ECA lectin. Samples collected from each patient were immediately processed into a single cell suspension using a quick digestion step. This reduced the extent of cell clumping which is a frequently observed characteristic of the sampling. The processed cellular samples were then diluted and split into 3-5 tubes (depending on the cell counts), with each tube being run for 20mins on the bioelectronic enrichment platform through the fluidic channel. Figure 15 also depicts the workflow for the clinical validation experiments. Several independent techniques were employed to assess the platform’s performance on clinical sampling which could be split into two categories – that is, on chip cell capture monitoring, and post-release cellular and molecular characterisation. Target cells captured on the MEAs after 1hr of sample processing were assessed by the previously established dual-mode optical-electrochemical characterisation method, by EIS and brightfield / fluorescence microscopy. Next, a series of cellular and molecular characterisation was carried out on aliquots of cell samples obtained during the experiment. One set of cellular sample aliquots were fixed and stained with H&E and IF staining of MUC5AC to verify the tissue of origin. Another set of aliquots were used to perform nucleic acid-based characterisation including RT-qPCR. Example 17 -Electrochemical characterisation of captured cells on chip As part of the clinical validation study, the inventors sought to test and compare cases of dysplastic BE with NDBE and controls. Characteristics of the patients enrolled for this study is presented below in Table 2. Four patients with varying history of BE were recruited. This included two BE cases with different segment lengths (as per the Prague classification) and a healthy volunteer with no history of BE. One patient with an endoscopic history of ‘indefinite for dysplasia’ (IND) failed to swallow the Cytosponge, which led to three patient samples being tested on the bioelectronic enrichment platform. Table 2. Patient characteristics for clinical testing of the bioelectronic enrichment platform. Indefinite for Dysplasia – IND. Figure 16 represents the EIS data of captured cells on 2 MEAs (3 electrodes each) for every patient tested. Each measurement was taken in redox electrolyte (FeCN) introduced in the fluidic channel promptly after cell capture and release was confirmed optically. For capture of target cells from the Cytosponge, diluted samples (in PBS) were run for 1hr in the fluidic channel on the lectin functionalised MEAs. The thermal detachment of cells was achieved by introducing a wash step with pre-heated PBS (kept at 37°C). Figure 16 reports the EIS measurements taken on the microfluidic MEA platform at baseline (lectin ECA functionalised), at cell capture (i.e after processing the sample through the fluidic channel for 1hr) and after thermal detachment for each patient sample. Comparing the quantification obtained for cells captured on the three samples tested, a trend could be observed wherein the impedance signal correlated with the presence / severity of BE. The ΔRct values indicated that the number of cells captured on the longer segment BE sampling to be highest, followed by that of shorter segment BE and then healthy volunteer sampling, where a modest increase in signal was observed compared to the baseline. Post-thermal detachment, the impedance signal was observed to drop, although not usually back to its baseline measurement which is consistent with previous findings. To take EC measurements, the fluidic inlet and outlet were disconnected from the syringe pump. The RE, and CE were dipped either in the inlet or the outlet based on proximity to the array being measured, with the circular electrode in the MEAs (with attached cells) acting as the WE. This change in the placement affecting the proximal distances across the three electrodes led to a discrepancy in impedance measurement, from those measured with a static well on a single MEA chip (see for example Zhang, F. et al. Biotechnol Bioeng 111, 1931– 1939 (2014)).. Therefore, absolute quantification based on previously established standard curves was not possible. However, the relative quantification of EIS measurements in this preliminary clinical validation study were obtained within 2hrs of Cytosponge sampling, which indicated significant differences in impedance signal between BE cases and the healthy volunteer sampling. These results highlight the potential for making a relatively quick determination of either the presence or length of BE. By collecting data from a larger cohort of samples, an impedance threshold- based approach could be used to make a determination of clinically relevant BE (>3cm length). These preliminary results indicate that the EC measurements independently could present an attractive alternative to the traditional pathology-based processing workflows which tend to take a significantly longer amount of time and resources. Example 18 -Optical characterisation of captured cells on chip The EIS measurements reported above provides a relative quantification of cells captured on each patient sampling. To supplement these findings, and to verify the identity of the cells captured as columnar cell groups, the inventors performed optical characterisation of the MEAs. The presence of the optical window within the electrode offers an opportunity for additional layer of optical confirmation of cell capture on the MEAs, in addition to EIS measurements. While the circular window only provides a snapshot of the entirety of cells captured on the electrode surface area, it presents a glimpse into the type of cells being attached which could be evaluated further optically. Figure 17b (top panel), depicts representative brightfield images of cells captured on the lectin- functionalised electrode from BE patient sampling after 1hr of running the sample on the MEAs. Two interesting findings from these images emerge. First, the general morphology of the cells captured seem to resemble the gastric columnar cell group structure with groups of tall cuboidal cells, which is distinct from the squamous cells which are flat and sheet-like. This indicated in the first instance, the ability of the platform to capture cellular clusters dynamically as they’re flowing through the channel which has been a challenge for several microfluidic cell capture devices historically. Second, the relative number of cells observed seemed to corroborate the impedance measurement data, wherein long segment BE indicated more densely packed cell groups attached within the optical window, while the number of cells attached on the short segment and healthy volunteer was markedly lower. However, further sample testing in a larger cohort is required to verify these findings. A key finding from initial optimisation experiments was that cells captured on the lectin- functionalised MEA remained attached to the surface post fixation with 4% PFA (not represented here). This could therefore allow for direct on-chip staining, which resembles some of the capabilities of current CTC-capture devices wherein captured cells are confirmed optically by means of IF staining with cell-specific antibodies such as EpCAM, CD45 etc. Exploiting this finding, on-chip staining was performed on cells captured on the platform to verify their columnar identity. To achieve this, MUC5AC was chosen for IF staining of captured cells on chip, as a known marker of columnar cells of the gastric tissue. However, the PFA fixation step could affect the polymer actuation mechanism and the EIS measurements. Therefore, an aliquot of the Cytosponge sampling was run on another parallel lectin-functionalised array to capture columnar cells in the aliquot, and verify their identity by on-chip IF staining with the MUC5AC antibody. Figure 17b (bottom panel) depicts the IF staining of MUC5AC carried out directly on chip, on the cells captured on a parallel lectin functionalised MEA. The staining images confirms the presence of captured cells to be in fact columnar cells, which could have origins either in the gastric cardia or Barrett’s gastric metaplasia. This demonstrates another potential downstream diagnostic assay that could be carried out on the bioelectronic enrichment platform, to test cell-specific biomarkers, in addition to the quantitative estimates provided by EIS. Further staining with IM-specific biomarkers such as MUC2 or TFF3 could additionally provide insights into the capture of these cell types from the Cytosponge sampling. Example 19 -Post release cellular morphology and biomarker characterisation – IF, H&E The on-chip dual-mode optical-electrical monitoring of captured cells was conducted after 1hr of processing the Cytosponge sample on the platform. After these evaluations, additional sample processing on the channel was carried out for another 2-3 hrs to allow for maximum capture of target cell groups from the Cytosponge sample. The time for processing depended on the number of cells obtained from the Cytosponge which generally increased with the length of BE sample. After the diluted Cytosponge sample was completely processed on the platform, the captured cells were thermally detached by performing a wash step with pre- heated PBS (kept at 37°C). A new syringe was loaded to separately collect the detached cells which were later retrieved and split into aliquots that were either fixed (with 4% PFA) and stored at 4°C for cellular staining or stored directly in -80°C freezer for nucleic-acid based estimation. Figure 18 depicts the different cellular sample aliquots collected during the clinical testing experiment. Three sets of cellular material were collected and stored for characterisation by either cellular or nucleic acid-based methods. This included the unprocessed Cytosponge sampling representing the mixed heterogenous sampling, the washed unbound cells representing the non-specific healthy background squamous cells, and the target cells detached from the platform, representing potentially the enriched columnar cells from the Cytosponge. The PFA-fixed aliquots of each of these cellular samples were transferred onto a glass slide by the Cytospin technique as described above. Figure 18 shows the representative H&E and MUC5AC staining carried out on samples collected from the long segment BE case. The H&E staining in Figure 18a depicts the morphological features of cells present in each of the cellular samples collected during the experiment. A qualitative observation of columnar cell enrichment is provided when comparing the slide representing the unprocessed Cytosponge with that of detached target cells. In the enriched sample, columnar cell groups (indicated with red circles) were observed to be less crowded with squamous cells as compared to the unprocessed Cytosponge sampling. The middle panel representing the washed unbound cells from the platform was used to evaluate the extent of columnar cells which were missed being captured by the lectin-functionalised electrodes. Based on morphological features alone, only squamous cells were observed to be present, indicating minimal loss of target cell capture. Figure 18b depicts the MUC5AC staining that was carried out to validate the morphological assessment. A confirmation of the presence of columnar cell groups is presented with the positive staining of MUC5AC in both the unprocessed and enriched columnar fractions. No green signal arising from MUC5AC binding is observed in case of washed squamous cell fraction, which instead indicates the presence of a large cellular clump of normal healthy squamous cells. Comparing the unprocessed and the enriched cell fractions, it can be noticed that the number of background squamous cells (indicated by blue DAPI-stained cells without any MUC5AC staining) is considerably lower, which can be seen to present better resolution for detection of columnar cells. This further presents a qualitative marker-based indication of columnar cell enrichment from the platform. Further testing of the Cytospin slides with markers such as MUC2 could indicate specific enrichment of IM cells from the Cytosponge sampling. Example 10 -Post-release RNA characterisation – RT- qPCR Assessment of gene expression by RT-qPCR was conducted to critically evaluate the physical enrichment of the nucleic acid fraction of columnar cells from the Cytosponge sampling. An aliquot of cellular samples collected during the clinical validation experiments were stored for nucleic acid-based characterisation. The stored aliquots were used to simultaneously extract DNA and RNA using the Qiagen Allprep extraction kit. The extracted RNA samples from the sample aliquots were then queried by RT-qPCR using pre-optimised primers for several genes of interest. Two mucin genes were included to ascertain the enrichment of columnar cells arising primarily from gastric cardia (MUC5AC) and those from Barrett’s IM (MUC2). Additionally, several other IM-specific genes were included in the assay to test potential enrichment in gene expression. Figure 19 represents the relative gene expression values for all gene markers tested on extracted RNA material from cellular sample aliquots. The sample aliquots used for this analysis was derived from processing of the Cytosponge sample of patient with long segment BE (BEST2 / CAM / 0872). First, it can be observed that all gene markers tested indicated minimal expression on the washed squamous cell fraction, indicating that very few of the target cells are being missed by the platform. Mucins are an integral set of biomarkers in the oesophageal lining, which has been demonstrated to be abundantly expressed during BE development (see for example Bhat, S., Bashir, N. & Mir, S. A. Int J Res Med Sci 7, 1282–1287 (2019). And Dixon, J. et al.. Am J Gastroenterol 96, 2575–2583 (2001).). The gene expression data presented in Figure 19 corroborated a fold change of gene expression in the enriched sample compared to the unprocessed Cytosponge sampling, which indicate a statistically significant difference for MUC5AC (p value – 0.0294) and for MUC2 (p value < 0.0001). Especially, the increased expression of MUC2 in the enriched fraction provides the first indication of the platform’s potential to enrich specific genes of interest which could be particularly useful for downstream sequencing-based approaches. Current investigations are exploring the analysis of methylation sequencing data derived from these cellular fractions, aiming to provide deeper insights into the efficacy of the platform's enrichment strategy for identifying methylation-based biomarkers. It is imperative to conduct further testing on clinical samples with varying histories of BE and dysplasia to compare the relative expression patterns across the three sample fractions and conclusively establish this point. These preliminary results show promise in achieving a physical enrichment of nucleic- acid markers using the proposed platform, thereby offering potential for enhanced downstream sequencing-based analysis. This improvement could be greatly beneficial for the determination of dysplasia in patients, with greater clinical relevance. Any publication cited or described herein provides relevant information disclosed prior to the filing date of the present application. Statements herein are not to be construed as an admission that the inventors are not entitled to antedate such disclosures. All publications mentioned in the above specification are herein incorporated by reference. Various modifications and variations of the invention will be apparent to those skilled in the art without departing from the scope and spirit of the invention. Although the invention has been described in connection with specific preferred embodiments, it should be understood that the invention as claimed should not be unduly limited to such specific embodiments. Indeed, various modifications of the described modes for carrying out the invention which are obvious to those skilled in engineering, cellular biology and molecular biology or related fields are intended to be within the scope of the following claims.

Claims

CLAIMS 1. A bioelectronic cell enrichment platform comprising; a microelectrode array comprising a glass substrate coated with a conducting surface layer and a smart polymer, said smart polymer being coated with a capture substrate, said capture substrate being coated with a selective capture reagent, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises metal nanospheres, suitably gold nanospheres (AuNP).

2. The bioelectronic cell enrichment platform according to claim 1, wherein said selective capture reagent is a lectin, suitably wherein said selective capture reagent is a functionalised lectin, suitably Erythrina Cristagalli lectin.

3. The bioelectronic cell enrichment platform according to any preceding claim, wherein the capture substrate comprises AuNP having a diameter of 20-100nm, suitably 30-70nm, suitably 40-60nm, most suitably 50nm.

4. The bioelectronic cell enrichment platform according to any preceding claim, wherein the smart polymer releases at least the captured target cells when heated, suitably when heated to 30-40°C most suitably when heated to 32-37°C.

5. The bioelectronic cell enrichment platform according to any preceding claim, wherein the conducting surface layer comprises one or more of gold, indium, tin oxide, graphene, carbon nanotubes or graphite, suitably wherein said conducting surface layer comprises gold.

6. The bioelectronic cell enrichment platform according to any preceding claim, wherein the selective capture reagent selectively binds to cells indicative of disease, diagnostic or adisease or associated with a disease, suitably wherein the selective capture reagent selectively binds to cancer cells or cancer associated cells.

7. The bioelectronic cell enrichment platform according to any preceding claim, wherein the microelectrode array comprises an optical window.

8. The bioelectronic cell enrichment platform according to claim 6, wherein the selective capture reagent selectively binds to cells associated with Barret’s oesophagus.

9. The bioelectronic cell enrichment platform according to any preceding claim, further integrated with a microfluidic system.

10. A method of selectively obtaining target cells from a cell sample, said method comprising the steps of (a) incubating a cell sample with the bioelectronic cell enrichment platform of any of claims 1- 8; (b) washing away unbound cells; (c) heating to release captured target cells.

11. The method of claim 10 wherein heating step (c) is suitably to 30-40°C most suitably to 32-37°C.

12. The method of claim 10 or 11 wherein the target cells are disease cells, cells associated with or indicative of a disease, cancer cells, specific immune cells, gastric and intestinal cells types, goblet cells, columnar epithelium cells pathogenic bacterial cells, H.Pylori or Streptococcus pneumoniae.

13. The method of any of claims 10-12 wherein there is an additional step of analysing the target cells.

14. The method for detecting target cells using the platform of any one of claims 9-13, further comprising the steps of: (i) applying an alternating current voltage to the microelectrode array; (ii) measuring electrochemical impedance spectroscopy (EIS) signals before and after cell capture; (iii) correlating impedance changes to cell capture efficiency.

15. Use of the bioelectronic cell enrichment platform of any of claims 1-9 to identify cells associated with disease or risk of a disease.

16. Use according to claim 15 wherein the disease or risk of disease is Barrett’s oesophagus or cancer.

17. A bioelectronic cell enrichment platform comprising; a microelectrode array comprising a glass substrate coated with a conducting surface layer and at least two layers of insulating polymer, wherein multiple etching windows are created in the insulating polymer; wherein a smart polymer is layered on the conducting surface layer inside one or more etching windows, wherein a selective capture reagent is covalently coupled to a capture substrate layer which is electrostatically bound to the smart polymer surface, wherein said smart polymer comprises PEDOT:PSS / pNIPAAm; wherein said capture substrate comprises metal nanospheres, suitably gold nanospheres (AuNP).

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