Cytochrome c oxidase deficiency detection in human fibroblasts using scanning electrochemical microscopy

Significance

Cytochrome c oxidase deficiency is an incurable mitochondrial disease, but symptoms in young infants can be managed if an early diagnosis is made, improving the quality of life of affected individuals. Currently, this disorder is diagnosed through muscle biopsies which are invasive, expensive, and time-consuming, leading to a delay in diagnosis. Our study proposes the quantification of cytochrome c oxidase (COX) activity in living fibroblasts using scanning electrochemical microscopy. Our results confirm N, N, N′, N′-tetramethyl-para-phenylene-diamine as electroactive indicator for COX activity and provide a proof of concept for the successful detection of COX deficiency.

Abstract

Cytochrome c oxidase deficiency (COXD) is an inherited disorder characterized by the absence or mutation in the genes encoding for the cytochrome c oxidase protein (COX). COX deficiency results in severe muscle weakness, heart, liver, and kidney disorders, as well as brain damage in infants and adolescents, leading to death in many cases. With no cure for this disorder, finding an efficient, inexpensive, and early means of diagnosis is essential to minimize symptoms and long-term disabilities. Furthermore, muscle biopsy, the traditional detection method, is invasive, expensive, and time-consuming. This study demonstrates the applicability of scanning electrochemical microscopy to quantify COX activity in living human fibroblast cells. Taking advantage of the interaction between the redox mediator N, N, N′, N′-tetramethyl-para-phenylene-diamine, and COX, the enzymatic activity was successfully quantified by monitoring current changes using a platinum microelectrode and determining the apparent heterogeneous rate constant k₀ using numerical modeling. This study provides a foundation for developing a diagnostic method for detecting COXD in infants, which has the potential to increase treatment effectiveness and improve the quality of life of affected individuals.

Cytochrome c oxidase (COX), also known as complex IV, is a multimeric protein composed of fourteen subunits that act as the terminal electron acceptor of the electron transport chain, which is responsible for coupling oxidative phosphorylation with ATP production (1, 2). Found in the inner membrane of the mammalian mitochondria (2), three of the subunits of this transmembrane protein are encoded by mitochondrial DNA while the others are encoded by nuclear DNA (3). Mutations in mitochondrial or nuclear genes encoding COX lead to the incorrect assembly of COX, resulting in functional defects which cause a rare heterogenous disorder called cytochrome c oxidase deficiency (COXD) (4). This disorder dramatically affects vital organs such as the heart, brain, liver, and skeletal muscles, causing a range of degenerative diseases, from lethal infantile myopathy to Leigh syndrome and hepatic encephalopathy (4–6). Being the most prevalent among all mitochondrial disorders (30% of the cases) (7, 8), which affect 1 in 5,000 people worldwide (9), COXD is often fatal in childhood, with only few individuals surviving adolescence (4, 10). Mutations in thirty possible nuclear and mitochondrial genes are known to cause COXD (4), which includes alterations in the SCO1 nuclear-encoded gene. SCO1 encodes the SCO1 metallochaperone protein (11, 12), which mediates copper transport from subunit COX17 to COX2, allowing the assembly of the CuA copper site in COX and ultimately enabling the proper biosynthesis of the protein (3, 11). Therefore, mutations in SCO1 indirectly impair the proper assembly of COX, leading to the biosynthesis of nonfunctional COX molecules. COXD caused by SCO1 mutation results in clinical phenotypes such as hepatic failure (10), fatal encephalopathy (4), and ketoacidosis (7).

Currently, COXD is mostly diagnosed by collecting skin and muscle biopsy samples from patients which are subject to a variety of enzymatic assays and histochemical methods including spectrophotometric, immunofluorescent, and cytochrome c oxidase/succinate dehydrogenase double-labeling (COX/SDH) assays (13–17). These methods present several drawbacks. For instance, COX/SDH is often limited by its inability to detect COXD at low levels due to this method’s dependence on staining (18). Immunofluorescence-based methods require expensive antibodies to detect COX activity, which significantly increases the cost of the technique (16, 19). Additionally, these assays are labor-intensive, time-consuming, and often require invasive muscle biopsies from young patients (16, 18, 20). To minimize invasive procedures, studies have proposed COXD diagnosis using epithelial cells collected with buccal swabs (21, 22) and lymphocytes isolated from patient blood samples (23), with the use of detection methods such as spectrophotometry (21), dipstick immunocapture assays (22) and high-resolution respirometry (23). Although these approaches are less invasive than biopsies, some of these methods are time-consuming, as they require protein or cell isolation procedures, whereas others provide only qualitative results. This might present sensitivity issues since the detection solely relies on the visual observation of test stripes, which could lead to false negative results. Exploring alternative approaches that may lead to inexpensive, quick, and highly sensitive detection methods in the future is therefore crucial to establish more efficient COXD diagnosis.

Electrochemical methods have revealed an outstanding potential to be used as diagnostic assays to detect many disease biomarkers. Specifically, scanning electrochemical microscopy (SECM) stands out as an electroanalytical technique that can detect trace levels of biomolecules with high spatial and temporal resolution (24–28). In SECM, a microelectrode is scanned across an underlying surface monitoring an electrochemical current while recording its lateral tip position, and the effect of electrode properties has been studied in the past (29). Unlike other scanning probe techniques, SECM not only reveals the topography of samples in a solution but also enables the detection and visualization of local variations in their electrochemical reactivity. The high sensitivity of this method and the micrometric dimensions of the sensor make SECM a particularly attractive technique to study biological samples, such as single living cells and tissues (24, 30, 31).

Biological applications of SECM have been described, including studies of cancer (32–35), cardiac disorders (36, 37), oxidative stress (38, 39), bacterial metabolism (40–42), and cell interactions in biofilms (43, 44). The ability of SECM to detect metabolites locally makes it an ideal tool to discover biomarkers and metabolic redox indicators at the single-cell level (39, 45, 46), which has the potential to enhance the understanding of cellular mechanisms involved in the onset and progression of diseases. The use of SECM for the study of biological cells and tissues has achieved impressive advances in the last two decades. For instance, Hu and colleagues have recently utilized platinized carbon nanoprobes to dynamically quantify reactive oxygen and nitrogen species inside single phagolysosomes in living macrophages to better understand the cellular phagocytosis process (38). In a promising proof-of-concept study, Thomas and coworkers were able to measure the production of oxygen by living algae cells, in situ and in real time, using scanning photoelectrochemical microscopy and micro-optical-ring gold electrodes as powerful detection probes (45). On the other hand, the regeneration of muscle tissue was recently investigated by Bironaite and colleagues using SECM, and their findings revealed that this electroanalytical method can be used to locally stimulate specific subpopulations of skeletal muscle–derived mesenchymal stem/stromal cells to improve their regeneration potential and functioning, while measuring their redox activity (47). These are only a few examples for SECM’s versatility and potential as a valuable tool to investigate a range of biological processes.

Herein, we propose a sensitive electroanalytical method based on SECM to quantify COX activity in living primary fibroblasts, which are known for being a model cell line used to study mitochondrial disorders associated with COX (48). Our detection approach uses the redox mediator N, N, N′, N′-tetramethyl-para-phenylene-diamine (TMPD). TMPD is commonly used as the substrate in the oxidase test to determine the presence of cytochrome c oxidases in bacteria (49, 50), and has been previously employed to quantify bacterial cytochrome c oxidase activity using electrochemistry (51). Herein, SECM measures fibroblasts’ ability to oxidize TMPD to TMPD^+•, which is mediated by COX. The oxidized species diffuse through the mitochondrial outer membrane and cross the lipid cell bilayer to reach the microelectrode, where it is reduced. The generated current response is a convoluted signal that is influenced by the cell’s reactivity, but also by the cell’s topography (52). To decouple the effect of topography and reactivity on the detected current signal, a numerical modeling approach described previously is employed (33, 53). This methodology enables the extraction of an apparent heterogeneous rate constant and thereby quantifying COX activity in living fibroblasts. By validating TMPD as a disease indicator by means of SECM, we demonstrate SECM’s application for disease detection. This is a report on an electrochemical method devoted to investigating COXD, and the presented results have the potential to serve as a foundation for the future development of electroanalytical assays for COXD diagnosis.

Results and Discussion

COX Redox Mechanism and TMPD Electrochemistry.

As shown in Scheme 1A, under normal aerobic conditions, COX catalyzes the electron transfer from reduced cytochrome c to molecular oxygen, resulting in water and oxidized cytochrome c, while transporting one proton per electron from the matrix to the intermembranous mitochondrial space (2). Simultaneously, cytochrome c oxidoreductase or complex III (CRE) pumps two protons per electron while catalyzing the electron transfer from ubiquinol (QH₂) to oxidized cytochrome c, restoring cytochrome c to its reduced form. Cytochrome c can only transfer one electron at a time; hence, to produce one water molecule, cytochrome c must undergo reduction twice (2).

Scheme 1.

Schematic representation of cellular and electrochemical reactions. (A) COX (violet) and cytochrome c reductase (orange) complexes under normal aerobic conditions in a mammalian cell. (B) Proposed mechanism of complex IV under TMPD concentration > 0.4 mM; TMPD (T), TMPD⁺ (T^+·), OMM: outer mitochondrial membrane, IMS: intermembrane space, IMM: inner mitochondrial membrane, MM: mitochondrial matrix. (C) SECM measurements at 12 μm above the surface of a single fibroblast cell in 1 mM TMPD in feedback mode.

To enable the electrochemical detection of COX activity using SECM, we used TMPD as the redox mediator due to its well-known interaction with this enzyme (49–51). In the presence of TMPD concentrations higher than 0.4 mM, electron transfer reactions can occur directly from TMPD to COX, without the involvement of cytochrome c (54, 55). Scheme 1B proposes the cellular uptake of the membrane-permeable redox mediator TMPD (54). TMPD crosses the plasma cell membrane barrier, as well as the mitochondrial outer membrane, interacting with COX located in the inner mitochondrial membrane. TMPD undergoes oxidization to TMPD^+•, transferring one electron to COX to reduce oxygen to produce water (56). Just as in the case of cytochrome c, two TMPD molecules are required to produce one water molecule. COX can transport one proton per electron into the intermembranous space, allowing ATP synthesis to continue through the ATP synthase. Once oxidized, TMPD^+• passes through the outer mitochondrial membrane and the cell plasma membrane to interact with the surface of the microelectrode as it passes over the cell, as seen in Scheme 1C. A reduction potential applied at the electrode reduces TMPD^+• back to TMPD, creating a feedback loop, which produces an increase in electrochemical current during SECM imaging.

To understand TMPD’s role as a redox mediator for single-cell SECM studies, it was electrochemically characterized using cyclic voltammetry. TMPD undergoes two electron transfer steps, generating TMPD^+• first, and TMPD²⁺ in the second electron transfer step (57). As shown in Fig. 1A, TMPD oxidizes at +200 mV to produce TMPD^+• and is subsequently oxidized to TMPD²⁺ at +600 mV. Reduction potentials are observed at +200 mV and −200 mV. The effect of electrolyte composition was studied in Dulbecco’s Modified Eagle’s medium (DMEM), in the presence and the absence of fetal bovine serum (FBS), and 0.1 M KCl (SI Appendix, Fig. S1). Although similar voltammetric profiles were observed for TMPD in DMEM media with and without FBS (SI Appendix, Fig. S1), all SECM experiments were performed in the absence of serum, because of potential electrode fouling over time (58). The increase in the cathodic current at potentials more negative than −0.3 V (vs. Ag/AgCl) observed in Fig. S1 is most likely related to the reduction of oxygen dissolved in the media. To avoid the possible signal interference by oxygen, SECM measurements on living fibroblasts were performed by applying a potential at the microelectrode that corresponds to the mediator’s first electron transfer step (+200 mV for TMPD oxidation to TMPD^+• and −200 mV for TMPD^+• reduction back to TMPD). The electrochemical behavior of TMPD was validated at varying scan rate values (10 to 300 mV s⁻¹) using a Pt macroelectrode (diameter = 3.0 mm), with the potential being swept from −200 to +200 mV. The anodic peak current varied linearly with the square root of the scan rate, therefore confirming that the oxidation of TMPD to TMPD^+• consists in a diffusion-controlled process. This result is in full agreement with the literature reporting TMPD redox behavior on metallic surfaces (51).

Fig. 1.

(A) Voltammetric profile of 1.0 mM TMPD at a 25-μm platinum microelectrode in 0.1 M KCl solution. Reduction and oxidation potentials of TMPD are shown by red dashed lines (respectively). (B) Cyclic voltammograms in 1.0 mM TMPD at a 3-mm Pt macroelectrode in 0.1 KCl solution at different potential scan rates. The inset shows the relationship between the anodic peak current and the square root of the potential scan rate.

Electrochemical Monitoring of TMPD Interaction with Living Fibroblasts.

The two fibroblast cell lines used in this work included a control cell line (Control) of healthy fibroblasts, capable of expressing a fully assembled and functional COX enzyme. SCO1 cells carry the SCO1 gene mutation that impairs the proper biosynthesis of COX (3, 11). Both cell types present a similar morphology, characterized by an elongated shape with a fibrous aspect (SI Appendix, Fig. S2). To validate the cell lines of choice, expression levels of fully assembled COX and SCO1 proteins in the Control and SCO1 cell lines were quantified by Western blot analysis. The SDS-PAGE result displayed in Fig. 2A confirms the reduced level of SCO1 protein (29 kDa) in SCO1 cells. VDAC1 (35 kDa) was used as a loading control and was detected equally in both cell lines. Accordingly, blue native page gel electrophoresis (BN-PAGE) results (Fig. 2B) show that cells carrying the SCO1 mutation present reduced levels of fully assembled COX (Complex IV) due to decreased levels of SCO1 protein (9). Complex I was used as a loading control and was detected equally in both fibroblast lines. The decreased levels of SCO1 and COX proteins in SCO1 cells, therefore, validate SCO1 as a COXD cell line. Differential electrochemical signals are therefore expected for COX activity in Control cells compared to SCO1 deficient fibroblast cells.

Fig. 2.

Western blot results of mitoplasts isolated from Control and SCO1 fibroblast cells. (A) Fibroblasts fractionated by SDS-PAGE and blot were subjected to antibodies against SCO1. VDAC1 was used as a loading control. Molecular weight (MW) protein ladder is indicated on the left. (B) Fibroblasts fractionated by 1D BN-PAGE and blot were subject to antibodies against COX (complex IV). Complex I was used as a loading control.

Prior to SECM measurements, cells were exposed to DMEM basic media (no FBS) in the presence of 1 mM TMPD for 15 min. Cell viability was not affected by TMPD exposure according to a trypan blue exclusion cell viability assay in the literature, where it has been reported previously that cell exposure to TMPD for 15 min does not affect viability (59). However, the absence of FBS in the cell culture media is known to affect both cell viability and morphology (58). By conducting a cell viability study over 90 min, we were able to confirm that the viability and morphology of Control and SCO1 fibroblast lines remained unaffected for the first 60 min (SI Appendix, Fig. S3), and no external signs of stress were observed through the SECM-integrated optical microscope. This ensures that fibroblasts remained viable during all SECM line scans.

Approaches have been described for the study of living cells using SECM, including the use of depth scans (47, 60–63) and constant-height lateral scans performed in the x, y-direction (39, 45, 64–66) Lateral scans significantly reduce the risk of making physical contact between the electrode and the biological cell. Depending on the imaging system of an SECM instrument, stopping an approach above a cell, especially when using a colored redox mediator, such as TMPD, can be very challenging. The presented methodology proposes an alternative to vertical cell scans to protect the physical integrity of the bioentities, and to avoid mechanical stimuli influences on the cellular redox activity and response.

Three-dimensional SECM images (Fig. 3) of single Control and SCO1 fibroblasts show a differential current behavior between the two cell lines, qualitatively. In this representative example of an experimental dataset, larger current signals from a Control cell (Fig. 3 A and B) can be seen compared to the current obtained from a SCO1 cell (Fig. 3 C and D). Three-dimensional images were obtained in 1 mM TMPD at a 50 µm s⁻¹ scan velocity, using a microelectrode biased at the potentials of +200 mV (oxidation of TMPD) or −200 mV (reduction of TMPD^+•). It should be noted that a solution of TMPD prepared in the presence of oxygen naturally presents a ratio of 3:1 of reduced and oxidized TMPD (35), respectively. When a reduction potential of −200 mV is applied, the electrode detects the ability of the living cells to oxidize TMPD to TMPD^+•, which is reduced back to TMPD at the microelectrode (Scheme 1). During SECM constant height imaging of living cells, the measured signal is always a convolution of topographical contributions and reactivity-related signals (26, 37). Topographical contributions affect the current as the tip-to-substrate distance changes. For example, larger cell features on the substrate result in a decrease in current, because of the hindered diffusion of the redox active species toward the electrode. Cavities on the surface, in contrast, would result in an increase in current, as the tip-to-substrate distance increases. Reactivity describes the regeneration of the redox mediator by the living cell. As shown in Fig. 3 A and C, as the microelectrode passes over the cells, a clear increase in the electrochemical current signal was observed. In this case, the reactivity of the cells is able to overcompensate for the topographical contributions, resulting in an overall increase in current when the electrode passes the cell located on the petri dish. Similarly, when an oxidation potential of +200 mV is employed (Fig. 3 B and D), the electrode detects the ability of the living cells to reduce TMPD^+• to TMPD, which is then oxidized to TMPD^+• at the microelectrode. Interestingly, depending on the applied potential at the microelectrode, SECM is able to monitor the activity of both cytochrome c oxidase and reductase.

Fig. 3.

SECM 3D imaging of single living fibroblasts in basic DMEM media containing 1 mM TMPD at a 50 μm s⁻¹ scan velocity. Microelectrode polarized at +200 mV and −200 mV to scan Control (A and B) and SCO1 fibroblasts (C and D).

Fig. 4A demonstrates SECM line scans in TMPD across Control and SCO1 cells at varying scan velocities (10 to 100 µm s⁻¹). It is evident that, as the scan velocity is increased, the current measured at the microelectrode also increases because of greater mass transport caused by forced convection (53). Thereby, electrochemical currents that originate from contributions by topography are more susceptible to scan velocity changes, than contributions from reactivity. This forms the basis of the previously reported numerical model, capable of decoupling topography from reactivity (33).

Fig. 4.

SECM line scans across single living Control (A and B) and SCO1 (C and D) fibroblasts in basic DMEM media containing 1 mM TMPD, with scan velocities ranging from 10 to 100 μm s⁻¹ and the microelectrode biased at +200 V (A and C) and −200 V (B and D). Graphs show normalized currents (tip currents divided by the initial current observed at the beginning of each scan). Insets in A and C show optical micrographs of the studied cells, and the electrode edge path during line scan acquisition is represented by the black dashed lines.

In comparison, Control cells exhibit higher electrochemical currents than the SCO1 fibroblasts, both at +200 mV (CRE activity) and −200 mV (COX activity), which is in agreement with the phenotype of each cell line. However, if cells exhibit differential topographies, the assessment of cellular activity based on pure faradaic currents is inaccurate, as cells of different shapes and sizes will have differential contributions to the measured electrochemical current from topography and reactivity. Numerical modeling is an effective strategy to decouple topography from reactivity and to extract an apparent heterogeneous rate constant k₀, which indicates the cell’s ability to regenerate a redox mediator in solution, as demonstrated previously (33, 53, 67).

Extraction of the Apparent Heterogeneous Rate Constant (k₀).

Using numerical modeling, TMPD oxidation and TMPD^+• reduction occurring at the microelectrode after interaction with COX and CRE in living fibroblasts can be utilized to extract the apparent heterogeneous rate constant (k₀), representing the biological cell kinetics. This numerical approach was described in detail in previous studies (53, 67), and is based on the extraction of a substrate’s kinetic rate from electrochemical data obtained using SECM constant height imaging at slow scan velocities (<100 µm s⁻¹). The numerical COMSOL model considers experimental parameters such as the microelectrode size, steady state current, scan velocity, diffusion parameters of redox species, as well as cellular features, including the cell’s shape and size, to quantify the redox environment of a cell. This mathematical approach fits the theoretical results from the substrate kinetic rates varying from 10⁻⁸ to 10⁻¹ m s⁻¹ to the normalized current detected at the microelectrode as the function of scan velocities. Because the microelectrode current depends on the scan velocity of the tip, the diffusion coefficient, and the tip-to-sample distance, the kinetic rates can be extracted from the normalized shear Peclet number (P_s) (53).

A visualization of the electrode and fluid movement within the simulation is presented in SI Appendix, Fig. S6. When a microelectrode passes a living cell, the cell-surrounding fluid is forced to pass in between the electrode tip and the body of the cell. Because of the limited amount of space for the fluid to pass, the concentration of the redox species in between the cell is temporarily increased. The height of a cell thereby affects the measured current more than the reactivity of the cell, which remains constant. Therefore, the faster the scan velocity, the more fluid is pushed between the microelectrode and the cell. By simulating the scanning profiles for several scan velocities, an average kinetic rate k₀ can be extracted. Fig. 5 shows the normalized tip current as a function of P_s for the Control and SCO1 fibroblasts (Fig. 5 A and C, respectively). For both COX and CRE activity, the SCO1 values are lower than the Control values, confirming slower kinetic rates in the SCO1 fibroblasts (Fig. 5 B and D).

Fig. 5.

Summary of the numerical modeling results for the analysis of Control and SCO1 fibroblasts using SECM. (A and B) COX activity and (C and D) CRE activity. (A and C) The colored lines represent the simulated normalized peak currents for several substrate and sample kinetics ranging from 10⁻⁸ to 10¹ m s⁻¹. The current signal for Control (C65, square) is higher than the one for SCO1 fibroblast (circle). (B and D) The normalized current is taken at low P_s (<1) as a function of kinetic k₀.

Validation of Results through NaN₃ COX Inhibition Bioassay.

The results of SECM and the numerical modeling were validated by performing a spectrophotometric bioassay for the determination of COX activity in Control and SCO1 fibroblasts. This assay is based on the capture of COX enzyme present in whole cell extracts by COX-specific antibodies immobilized on the wells of a microplate, followed by the addition of a cytochrome c solution to each well (n = 3). The decrease in the absorbance at 550 nm (absorption wavelength of cytochrome c) indicated enzymatic activity. The bioassay results (Fig. 6B) show that SCO1 fibroblasts naturally presented a significantly lower COX activity compared to Control cells (P < 0.01), which is in full agreement with the literature (12). The treatment of Control and SCO1 cells with 5.0 mM NaN₃, a well-known COX inhibitor, resulted in a pronounced decrease in COX activity due to enzymatic inhibition for both cell lines (P < 0.01). Interestingly, the enzymatic activity of COX in Control fibroblasts dropped to the approximate level of SCO1 cells after treatment (P > 0.01), whereas the activity of SCO1 cells decreased even further in the presence of the inhibitor (P < 0.01). Here, it must be mentioned that the incubation of cells in the presence of 5 mM NaN₃ did not affect cell viability significantly over the incubation period (P < 0.01) (SI Appendix, Fig. S4), as reported in previous studies (68, 69). Therefore, the observed effects can be exclusively attributed to the azide’s strong inhibition activity toward COX. The azide anion of NaN₃ binds to the enzyme metal sites heme a₃ and Cu_B, causing a drastic impairment in the enzymatic function (70, 71). The decrease in COX activity in the presence of NaN₃ is also seen in SECM line scans (Fig. 6A and SI Appendix, Fig. S5) and numerical modeling (Fig. 6B).

Fig. 6.

COX inhibition assay. (A) SECM line scans across Control fibroblasts at varying scan velocities in basic DMEM media without serum containing 1.0 mM TMPD after incubation of cells in 5.0 mM NaN₃ for 6 h. Microelectrode biased at −200 mV for SECM line scans. (B) Percent changes observed in k₀ (SECM; n = 4) and absorbance (bioassay; n = 3) after treatment of Control and SCO1 cells with 5.0 mM NaN₃ for 6 h (*P < 0.10; ***P < 0.01).

The average estimated k₀ values obtained for COX activity from untreated and NaN₃-treated Control and SCO1 cells are displayed in Table 1, along with the k₀ values for CRE activity (n = 4 individual cells in four different cell culture dishes). Quantitatively, the average COX kinetic rate for SCO1 cells was found to be significantly lower than the one obtained from the Control fibroblasts (P < 0.10), which is in clear agreement with the phenotype of the cells analyzed in this study. After the NaN₃ treatment, the COX k₀ value of both Control and SCO1 cells significantly decreased (P < 0.10), as a result of the enzyme inhibition by azide anions (SI Appendix, Fig. S7). The difference in the average k₀ values of untreated Control and SCO1 cells was found to be nonsignificant when the CRE activity was analyzed (P > 0.10). The mutation in the SCO1 gene in SCO1 fibroblasts affects the biosynthesis of COX, with no direct effect on the expression of the CRE enzyme. However, the observable trend of reduced activity of CRE in SCO1 cells is not surprising as the activities of the enzyme complexes in the electron transport chain are intrinsically connected (72, 73). Importantly, the agreement of results obtained by the proposed SECM method, and the NaN₃ spectrophotometric bioassay confirms the validity of TMPD as a disease indicator for COXD using SECM.

Table 1.

Summary of the apparent heterogeneous rate constant (k₀) values determined for untreated and NaN₃-treated Control and SCO1 cell lines from SECM line scans results

Cell line	k0 COX (× 10−5 m s−1)		k0 CRE (× 10−5 m s−1)
	Untreated	NaN3-treated
Control	3.49 ± 1.45	0.411 ± 0.129		4.16 ± 2.83
SCO1	2.10 ± 0.912	0.959 ± 0.0708		2.35 ± 1.52

Conclusions

In this study, we present SECM as a promising and effective method to detect and quantify COX activity in living fibroblasts as a biomarker of COXD. The presented electroanalytical approach was developed using fibroblasts as model cell lines and was based on quantifying current variations resulting from the intracellular interaction of the redox mediator TMPD with COX. An apparent heterogeneous rate constant for the turnover of TMPD/TMPD^+• by single fibroblast cells was extracted using numerical modeling. The study of the cell reaction kinetics enabled the successful decoupling of the effects of cell reactivity and cell topography on the overall bioelectrochemical response, hence giving insight into the differential COX activity between Control and SCO1 deficient cells. TMPD/TMPD^+• regeneration was found to be considerably slower in the SCO1 cell line compared to healthy Control cells, making this SECM approach a reliable method for COX activity detection in human cells. The presented proof of concept presents a rapid cell analysis procedure, which avoids nonideal experimental conditions, such as temperature fluctuations due to prolonged cell imaging. Furthermore, this approach can be conducted with basic SECM components and does not require specialized modules, such as shear force. Therefore, our study shows that electrochemistry coupled with numerical modeling presents a powerful combination that enables the differentiation of normal and dysfunctional forms of COX that result in the development of COXD. Validating this proof of concept in cell samples that can be obtained through minimally invasive methods in the future has the potential to create biosensing systems for COXD clinical diagnostics.

Materials and Methods

All chemicals were purchased from Sigma-Aldrich, and all media and materials for cell culture and related assays were purchased from Thermo-Fisher Scientific unless otherwise stated.

Cell Culture and Sample Preparation for SECM Analysis.

Control and SCO1 skin fibroblast cell lines were acquired from Montréal’s Children’s Hospital cell bank and provided by Dr. Eric Shoubridge (McGill University). These two primary cell lines were immortalized as described previously (74), and cultured in DMEM supplemented with 10% heat-inactivated FBS, in tissue culture flasks at 37 °C and 5% CO₂. After reaching 80 to 90% confluency, cells were harvested. To this end, cells were washed with 0.0067 M phosphate-buffered saline (PBS, pH 7.0 at 37 °C) and detached from the surface of the flask using 0.05% trypsin-EDTA solution at 37 °C. For SECM analysis, around 40,000 cells were seeded in 10 × 35 mm² petri dishes and incubated overnight under the conditions described above.

Western Blot Assay.

Following procedures previously described (12, 75, 76), fully assembled COX was detected by subjecting mitoplasts from Control and SCO1 fibroblasts to BN-PAGE. Mitoplasts were initially isolated using a mitochondrial isolation kit (89874, Thermo-Fisher Scientific). For BN-PAGE analysis, sedimented mitochondria were resuspended in 0.75 M aminocaproic acid and 50 mM Bis-Tris buffer at pH 7. 10% n-dodecyl-B-D-maltopyranoside was then added to the suspension and incubated on ice for 30 min. The mixture was centrifuged at 16,000×g for 30 min before 5% Coomassie Brilliant Blue stain in 0.5 M aminocaproic acid was added to the supernatant. For first dimension separation, a 6 to 15% native polyacrylamide gradient gel was used. The blot was subjected to antibodies against COX (ab14744, Abcam) and loading control complex I (ab110412, Abcam). SDS-PAGE was performed as previously described by Booy et al. (77), to detect SCO1 protein in Control and SCO1 fibroblasts. This blot was subjected to antibodies against SCO1 (PA554460 Invitrogen) and loading control VDAC1 (MABN504 MilliporeSigma). HRP-conjugated goat anti-rabbit (12348) and HRP-conjugated goat anti-mouse (12349) antibodies (Millipore Sigma) were used for chemiluminescent detection. Proteins were visualized using a ChemiDoc System (Bio-Rad).

Cell Viability Assay.

To determine the viability of fibroblasts in the presence of TMPD, trypan blue exclusion assays were conducted for Control and SCO1 cell lines. Cells were exposed to 1 mM TMPD in basic DMEM media (no FBS) for varying time intervals of 10, 30, 60, and 90 min. The medium was removed; cells were washed with PBS and then harvested after treatment with trypsin. Subsequently, fibroblasts were incubated in the presence of trypan blue (1:1) at room temperature for 3 min (78). Percent cell viability was determined using a Countess 2 Automated Cell Counter (Thermo Scientific). The same experimental protocol was used to evaluate cell viability of Control and SCO1 cells after their incubation for 6 h in 5 mM of the COX inhibitor NaN₃.

SECM Measurements.

All electrochemical experiments were performed using an ElProScan ELP3 system, with POTMASTER software (version V2 × 66) and an ElProScan Controller ESC 3 (HEKA Elektronik). A three-electrode setup consisting of an Ag/AgCl pseudoreference electrode, a 0.5-mm-diameter platinum wire auxiliary electrode, and a 25-µm diameter platinum microelectrode (RG = 2.8, HEKA Elektronik) was used. Prior to any measurement, the platinum microelectrode was polished using 0.05 µm alumina slurry (Buehler) on an automated electrode polisher (LaboPol-20, Struers Inc.). The electrochemical behavior of TMPD in basic DMEM cell media (no FBS) was investigated using cyclic voltammetry, with measurements carried out at a potential scan rate of 100 mV s⁻¹.

All SECM imaging measurements were carried out in feedback mode. TMPD (1 mM) was used as a redox mediator and was dissolved in basic DMEM media (no FBS). Prior to cell exposure, the TMPD solution was sterile-filtered using a 0.22 µm pore syringe filter (Millipore Sigma). Cells were incubated in the TMPD solution for 15 min at 37 °C before SECM data acquisition. Target fibroblasts were identified using an optical microscope located underneath the SECM stage. Prior to commencing line scans, a tip-to-substrate distance of 12 µm was achieved through a negative feedback approach curve over plastic in close proximity to a living cell of interest (53). All biological samples for this study were scanned at constant height with scan velocities ranging from 10 to 100 μm s⁻¹. Line scans were performed across a distance of 250 μm with a step size of 1 μm, resulting in the collection of 251 data points per line scan. All measured tip currents were normalized by the initial current observed at the beginning of each scan (I_T/I_{T, i}). During SECM data collection, dimensions and morphology of cells were constantly monitored using the SECM-integrated optical microscope. To ensure cell viability, SECM measurements were collected in under 30 min in the presence of TMPD redox mediator. Line scans were carried out with untreated Control and SCO1 living fibroblasts and with the same cell lines after treatment in the presence of 5.0 mM sodium azide (NaN₃) as a COX inhibitor. A NaN₃ solution prepared in PBS buffer was directly added to DMEM cell media (10% FBS), and cells were incubated for 6 h at 37 °C and 5% CO₂. To further validate SECM results, the enzymatic activity of COX was determined for untreated and inhibitor-treated Control and SCO1 cells using a spectrophotometric COX enzyme activity assay (Complex IV Human Enzyme Activity Microplate Assay kit, Abcam), which was performed according to the manufacturer’s instructions.

Statistical Analysis.

All experiments were performed in quadruplicates (unless stated otherwise), and all results are expressed as mean ± SD. Two-sample one-tailed t tests performed at 90% and 99% confidence levels were used to determine the existence of statistically significant differences between the results obtained for Control and SCO1 cell lines. All statistical analyses were performed using Origin 2019.

Supplementary Material

Appendix 01 (PDF)

Data, Materials, and Software Availability

All study data are included in the article and/or SI Appendix.