Effects of 2,3-dimercaptosuccinic acid modified Fe₂O₃ nanoparticles on microstructure and biological activity of cardiomyocytes

Abstract

Iron oxide nanoparticles (IRONs) have been widely applied in clinical magnetic resonance imaging and in vitro cardiac tissue engineering. However, the underlying effects of IRONs on the microstructures and biological activity of cardiomyocytes remain a controversial issue. In this study, IRONs were modified with 2,3-dimercaptosuccinic acid (DMSA-IRONs) to increase the hydrophobicity. The effects of DMSA-IRONs on the microstructures and biological activity of cardiomyocytes were systematically investigated. DMSA-IRONs were internalized by cardiomyocytes and mainly distributed in the cytoplasm in a dose-dependent manner. Live/dead assay and MTT assay demonstrated that DMSA-IRONs did not affect the survival of cardiomyocytes. The distribution pattern of F-actin and cell shape were also not changed by the incubation of DMSA-IRONs. Further investigations of the mechanism proved that DMSA-IRONs did not interfere with the formation of cell adherens junctions, gap junctions and intracellular Ca²⁺ transients of cardiomyocytes. However, the intracellular reactive oxygen species (ROS) concentration in cardiomyocytes with incubation of DMSA-IRONs was significantly reduced due to its peroxidase-like activity. In conclusion, DMSA-IRONs are of good biocompatibility and potentially beneficial for cardiomyocytes by decreasing the intracellular ROS. The study here provides a basic investigation of the DMSA-IRONs application in clinical cell-imaging and cardiac disease therapy.

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1. Introduction

Iron oxide magnetic nanoparticles (IRONs), which have large surface areas and quantum size effects, were considered as a single magnetic domain exhibiting superparamagnetic phenomenon.¹ Their specific physico-chemical, thermal and mechanical properties made them promising for biomedical applications, such as clinical magnetic resonance imaging for tissue and organ imaging, drug delivery and tracing implanted cells in vivo.^2–4 Previous studies have described that the iron ion could be released from the iron oxide core back into hemoglobin due to the break-up of IRONs in the acid environment within the lysosome.⁵ Recently, ferumoxytol (FerahemeTM), an ultra-small superparamagnetic iron oxide, was allowed to be applied as a supplement of iron for patients with anemia by the FDA.

The heart is a vital organ for a person, and heart injuries, such as myocardial infarction, remain one of the main causes of morbidity and mortality in the world.⁶ In the USA, approximately 500 000–700 000 die of ischemic heart disease per year, and the annual incidence rate of myocardial infarction was approximately 6/1000.⁷ Due to the physico-chemical characteristics, IRONs have been applied in cardiac tissue engineering in vitro and in clinical research of injured myocardium therapy.^8–12 In magnetic force based tissue engineering, IRONs were used to impregnate polymers to endow them with a magnetic response capacity, which provided a stimulating microenvironment for the construction of engineered cardiac patch in vitro.¹³ In terms of the effects of IRONs on cardiomyocytes in vitro, it was convincing that IRONs had few impacts on the cardiac mitochondrial respiratory chain complex activities, as well as on the cardiac differentiation of embryonic stem cells.^14,15 Mahmoudi et al. reported that IRONs could alter the cardiomyocytes gene expression profiles associated with cell proliferation.¹⁶ In vivo, IRONs were shown to be promising for tracking the implanted stem cells in infarcted myocardium, and no adverse effect was observed on the therapeutic capacity of the stem cells by IRONs.¹⁷ Besides, IRONs were proved to be positive for infarcted myocardium remodeling when as MRI contrast agent.¹⁸

The unitary biological activity of cardiomyocytes is based on their microstructures. N-Cadherins, which locate in the cytoplasm membrane and the sites of cell–cell interfaces, organize into adherens junctions to involve the mechanical modulation of cardiomyocytes.¹⁹ Meanwhile, gap junctions that are composed of connexin 43 provide the chance of direct electric coupling and transmission of action potentials of cardiomyocytes.²⁰ Though IRONs were used in heart imaging and cell transplantation widely, the underlying effects of IRONs on the microstructures and biological activity of cardiomyocytes remain largely unknown.

IRONs without any surface coating exhibit strong hydrophobic interactions to form large clusters, which present a higher magnetic moment than individual nanoparticles, and the magnetic dipolar interaction occurs between clusters to induce chain formation.²¹ Previously, the aggregations of IRONs were reported to evoke cytotoxicity,²² and therefore, suitable surface modifications of IRONs are often indispensable for biomedical applications. It was confirmed that DMSA with a carboxy group could form strong carboxylic chelating bonding with IRONs to increase the stability of an IRONs aqueous solution over broad ranges of pH and salt concentrations.^23,24 In this study, IRONs were modified with DMSA to increase their hydrophobicity (DMSA-IRONs). Cardiomyocytes were incubated with various concentrations of DMSA-IRONs and it was found that DMSA-IRONs have good biocompatibility with cardiomyocytes. Moreover, DMSA-IRONs rarely affected the microstructures, or electrophysiological performance of the cardiomyocytes, but decreased their intracellular reactive oxygen species (ROS) level.

2. Materials and methods

2.1. Materials

0.1% trypsin solution, nuclear fast red, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), phalloidin-FITC, DAPI and nitroblue tetrazolium salt (NBT) were obtained from Sigma-Aldrich (St. Louis, MO, USA). Dulbecco's modified Eagles medium (H-DMEM), fluo-4 acetoxymethylester and dihydroethidium (DHE) were purchased from Invitrogen/Gibco (Eugene, OR, USA). A Live/Dead Viability/Cytotoxicity Kit was obtained from Life Technologies (Eugene, OR, USA). The primary antibodies, N-cadherin, α-actinin and connexin 43, were obtained from AbCam (Cambridge, MA, USA), and the primary antibody GAPDH was purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). DMSA, di-azo-aminobenzene (DAB) and H₂O₂ were obtained from Aladdin (Shanghai, China). FITC-labelled goat anti-mouse IgG and Cy3-labelled goat anti-rabbit IgG were purchased from Boster (Wuhan, China), and PVDF membranes were obtained from Roche (Roche, Germany). The lysis buffer was obtained from Tiandz (Beijing, China).

2.2. Preparation of DMSA-IRONs

IRONs (γ-Fe₂O₃) were prepared by chemical coprecipitation, and the stable ferrofluid was then obtained via surface modification with DMSA, as reported.^23,25 Briefly, 100 mL mixed solution of FeCl₃·6H₂O (0.01 M) and FeSO₄·7H₂O (0.006 M) was modulated to pH 9 by ammonia solution (1.5 M). The resulted magnetite was washed with deionized water 5 times and 75% ethanol 2 times by magnetic separation. After dispersion in water, the pH of the magnetite solution was adjusted to 3.0 by acidification with 0.1 M HCl. Then the solution was oxidized in air under 100 °C for 1 h. The obtained γ-Fe₂O₃ nanoparticles were modified with DMSA, according to a previous study, at a 100 : 9.2 mol ratio (Fe : DMSA).²³ Subsequently, the products were washed repeatedly by magnetic separation. Finally, the resulting colloidal solution was mixed with a mannitol solution and lyophilized overnight. The morphology of DMSA-IRONs was observed by transmission electron microscopy (TEM) using JEM-2000EX. And the hydrodynamic diameter of the rehydrated DMSA-IRONs was determined by photon correlation spectroscopy (Malvern Zetasizer 3000, Malvern Instruments Co.) at 25 °C.

2.3. Isolation of primary cardiomyocytes and cell culture

The experimental protocol was approved by the Institutional Animal Care and Use Committee of the Academy of Military Medical Sciences. Cardiomyocytes were isolated using our established method.²⁶ In brief, cardiomyocytes were isolated from 1 day-old Sprague-Dawley (SD) rats. The ventricles were minced and digested with 0.1% trypsin solution. And the purified cardiomyocytes were obtained through pre-plating isolated cells on the tissue culture dishes for 2 h. H-DMEM with 15% fetal bovine serum was used to culture the collected cardiomyocytes. For all the following biological evaluations, the cardiomyocytes were cultured for 7 days and were treated with the final concentration of 0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹ DMSA-IRONs for 24 h.

2.4. Prussian blue staining

After incubation with DMSA-IRONs, the cardiomyocytes were washed 3 times with PBS (0.1 M, pH = 7.4), and subsequently fixed with 4% paraformaldehyde/PBS for 30 min. The fixed cardiomyocytes were incubated with 10% potassium ferrocyanide in 20% hydrochloric acid for 30 minutes, and counterstained with nuclear fast red.

2.5. Live/dead viability assay

Cell viability was measured by a live/dead cell assay using a Live/Dead Viability/Cytotoxicity Kit. After treatment with DMSA-IRONs, the cardiomyocytes were washed with PBS followed by the addition of 2 μM calcein acetoxymethyl (calcein AM) and 4 μM ethidium homodimer-1 (EthD-1). After incubation for 30 min at room temperature in the dark, the cardiomyocytes were visualized through a fluorescence microscope (Olympus).

2.6. MTT assay

To assess cell viability and/or metabolic activity, tetrazolium dye, MTT, was used, which could be reduced into formazan in living cells by mitochondrial succinate dehydrogenase and determined by absorbance. After removal of the culture medium containing DMSA-IRONs, the cardiomyocytes were incubated in serum-free H-DMEM with MTT (0.5 mg mL⁻¹) for another 4 h. Formazan crystals were dissolved with dimethylsulfoxide 200 μL. The absorbance was determined through a microplate reader (DYNATECH MR7000 instruments) at 570 nm. The relative cell viability (%) was calculated by

[A]_test/[A]_control × 100.

2.7. Immunohistological staining

After incubation of DMSA-IRONs, the cells were collected for immunofluorescent staining. Cardiomyocytes were labelled with primary antibody N-cadherin and the Cy3-labelled goat anti-mouse IgG as the secondary antibody, and phalloidin-FITC was detected at last. Cell nuclei were stained by DAPI. The nuclear shape index [NSI = (nucleus) width/length] and the cell shape index [CSI = 4π × area/(perimeter)²] were calculated by ImageJ2x software.²⁷

For the double labelling of sarcomeric actinin (α-actinin) and connexin 43, the cardiomyocytes were incubated with antibody α-actinin and antibody connexin 43 as the first primary antibodies, and FITC-labelled goat anti-mouse IgG and Cy3-labelled goat anti-rabbit IgG as the secondary antibodies. The cell nuclei were stained by DAPI. And the samples were observed under a Zeiss LSM 510 microscope.

2.8. Western blotting

Cardiomyocytes were solubilized to prepare the total protein. For SDS-PAGE separation, total proteins (100 μg) were loaded and transferred to a PVDF membrane incubated with antibodies against connexin 43 and N-cadherin. As control, the expression of GAPDH was evaluated. The analysis of band intensities was performed with ImageJ2x software.

2.9. Determination of cardiomyocyte intracellular calcium

The treated cardiomyocytes were rinsed with Tyrode's solution and were labelled with 10 μmol L⁻¹ fluo-4 acetoxymethylester (Fluo-4 AM) and 0.1% Pluronic F-127 for 45 minutes at 37 °C and 5% CO₂. Using a confocal laser microscope (Nikon Eclipse Ti-E confocal microscope), Fluo-4 AM was excited at 488 nm. The images were acquired at 10 frame per s, 256 × 256 pixels, and subsequently the fluorescence intensity was analyzed with the Volocity software, and the signals were normalized to basal cell fluorescence after fluo-4 loading (F₀). Intracellular Ca²⁺ was calibrated by the following pseudo-ratio equation:

[Ca²⁺]_i = K_d × (F/F₀)/(K_d/[Ca²⁺]_i-rest + 1 − F/F₀)

with K_d = 1100 nmol L⁻¹ and [Ca²⁺]_i-rest = 100 nmol L⁻¹.²⁸

2.10. In vitro peroxidase-like activity assay

The peroxidase-like activity of DMSA-IRONs was carried out, as previously reported, at room temperature.²⁹ Various concentrations of DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹) were dispersed in the reaction buffer 0.05 M Tris–HCl (pH = 7.5), 500 mM H₂O₂ and DAB substrate were then added into the mixture (DMSA-IRONs + DAB + H₂O₂ + reaction buffer groups). 30 min later, the results of reaction were recorded. The following groups served as negative controls: (1) H₂O₂ + DMSA-IRONs + reaction buffer; (2) DAB + DMSA-IRONs + reaction buffer; (3) DAB + reaction buffer.

2.11. Measurement of intracellular ROS

Intracellular ROS was determined by dihydroethidium (DHE) staining, which could interact with ROS to produce oxyethidium to emit a bright red color through reacting with nucleic acids.³⁰ The cardiomyocytes were exposed to DMSA-IRONs and incubated with DHE for 15 min in the dark at room temperature. To induce the oxidative stress, cardiomyocytes incubated with DMSA-IRONs for 24 h were treated with 100 mM H₂O₂ for 1 h followed with DHE incubation, as detailed above. After washing with PBS three times, the cells were examined under fluorescence microscopes (Olympus). The intensity of the fluorescence was determined by ImageJ2x software.

As another assay to verify the relative intracellular ROS compared with the control cardiomyocytes, nitroblue tetrazolium salt (NBT), was used.⁴ After incubation with DMSA-IRONs, the cardiomyocytes were cultured for another 6 h with fresh medium containing NBT (1 mg mL⁻¹) at 37 °C and 5% CO₂. And then to each well was added 95 μL lysis solution (0.04 N HCl in isopropanol). After shaking at 1200 rpm for 30 min under ambient temperature, 105 μL 10 N KOH was added, and the absorbance was obtained at 620 nm. To induce the oxidative stress, after 24 h of cultivation with DMSA-IRONs the cardiomyocytes were treated with 100 mM H₂O₂ for 1 h, and following this a NBT assay was used to determine the intracellular ROS, as detailed above.

2.12. Statistical analysis

Average and standard deviations were obtained from three separate experiments. All data are expressed as mean ± standard error of mean (SEM) unless indicated otherwise and analyzed using one-way analysis of variance (ANOVA) using the Tukey's post hoc test. Data was considered statically significant at P < 0.05.

3. Results

3.1. Characterization of DMSA-IRONs

Bare IRONs are instable in physiological aqueous solutions and have a tendency to degrade in biological systems.^31,32 In this case, we modified IRONs by coating with DMSA in this study. DMSA-IRONs showed good hydrophilic dispersion in water solution. The average diameter of iron core in TEM image was 7.9 ± 0.8 nm (Fig. 1A). And the hydrodynamic diameter of DMSA-IRONs measured by photon correlation spectroscopy (PCS) was 99.8 nm (Fig. 1B). For photon correlation spectroscopy, nanoparticles with hydration shells interact with each other in colloid solutions, and, therefore, the diameters of the nanoparticles determined by PCS appear bigger than by TEM. The suitable size and good hydrophilic dispersion of DMSA-IRONs show they are promising for further biomedical application. Therefore, it is necessary to evaluate the biological effects of DMSA-IRONs on cardiomyocytes at the microstructural and biological activity levels prior to clinical application.

Fig. 1 (A): TEM image of DMSA-IRONs; and (B): the hydrodynamic diameter of rehydrated DMSA-IRONs.

3.2. The internalization of DMSA-IRONs by cardiomyocytes and cell viability

Cardiomyocytes were incubated with DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹) for 24 h. The elongated cardiomyocytes could be observed in all groups exposed to DMSA-IRONs (Fig. 2A). The distribution of DMSA-IRONs in the cardiomyocytes was visualized by Prussian blue staining, which stained the iron oxide cores in blue. The results in Fig. 2B indicate that DMSA-IRONs could be strongly absorbed and formed massive agglomerates of cardiomyocytes after exposure to DMSA-IRONs. The internalized DMSA-IRONs in the cardiomyocytes were enhanced with increasing concentrations of DMSA-IRONs, which means that DMSA-IRONs could be internalized in a dose-dependent manner. Moreover, the internalized nanoparticles were mainly distributed in the cytoplasm around the cell nucleus, but not in the cell nucleus.

Fig. 2 (A): Observation of cardiomyocytes under phase contrast microscope; and (B): Prussian blue staining of cardiomyocytes treated by DMSA-IRONs for 24 h and the relative magnified areas. Black arrows indicate the location of intracellular iron with a blue color in the cardiomyocytes. (A and B): Scale bar = 50 μm, for magnified figures, scale bar = 25 μm.

Next, we examined the effects of the internalized DMSA-IRONs on the survival of cardiomyocytes. The results of a live/dead assay showed that most of the cardiomyocytes were alive (green), only a few of the cells were dead (red) after exposure to DMSA-IRONs for 24 hours, and no statistically significant effects of DMSA-IRONs on the viability of cardiomyocytes was observed (Fig. 3A). Consistently, the results of the tetrazolium dye (MTT) assay, which could be reduced by mitochondrial succinate dehydrogenase in living cells into formazan, also confirmed the great biocompatibility of DMSA-IRONs (Fig. 3B).

Fig. 3 (A and B): Live/dead staining of cardiomyocytes with incubation of DMSA-IRONs for 24 h (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹) and the analysis of the live/dead assays. A green color denotes live cells, and a red color denotes dead cells. (C): MTT assay of cardiomyocytes with incubation of DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹) for 24 h. Scale bar = 50 μm.

3.3. Analysis of DMSA-IRONs on the cytoskeleton arrangement and N-cadherin expression in cardiomyocytes

We further investigated the effects of internalized DMSA-IRONs on the microstructures of cardiomyocytes. Microstructures of cardiomyocytes, including actin filaments, adherens junctions composed of N-cadherin, affect the cell shape and function. The visualization of the cytoskeleton by labelling of the F-actin microfilament and N-cadherin could reveal the effects of DMSA-IRONs on the mechanotransduction structures of cardiomyocytes. The results in Fig. 4A indicate that the stress fibers of F-actin were distributed through the whole cytoplasm of cardiomyocytes and were connected to the cell membrane with exposure to DMSA-IRONs or not. Comparing with the control groups, few changes of F-actin arrangement could be observed in cardiomyocytes after exposure to DMSA-IRONs. The immunofluorescent results show that N-cadherin was mainly distributed in the cytoplasm membrane, and no apparent differences in the distribution of N-cadherin could be observed between the cardiomyocytes treated by DMSA-IRONs and the control cardiomyocytes (Fig. 4A). Moreover, the western blotting results also verified that the expression of N-cadherin was not affected by incubation of DMSA-IRONs (Fig. 6).

Fig. 4 (A): The double immunofluorescence of phalloidin (green) and N-cadherin (red) of cardiomyocytes under different concentrations of DMSA-IRONs; the nuclei were stained with DAPI (blue). Scale bar = 30 μm. (B): Nuclear shape index (NSI) between cells treated with DMSA-IRONs and the control cells is calculated. (P > 0.05); the cell shape index (CSI) is calculated between the cardiomyocytes with exposure of DMSA-IRONs and the control cells. (P > 0.05).

The cell shape index (CSI) was calculated by ImageJ2x software which could visualize the apparent cell shape. When the CSI approaches 0, this means the cell shows an elongated shape, while if the CSI approaches 1, this means the cell shows a more circular in morphology.³³ And the results indicate that when cardiomyocytes were treated by DMSA-IRONs, the shape of the cardiomyocytes was not affected (Fig. 4B). The nuclear shape index (NSI) indicates the morphology of the cell nucleus; when NSI approaches 1, the nucleus is circular, and when NSI gets close to 0, the nucleus is elongated. There was no significant difference in NSI between cells treated with DMSA-IRONs and untreated cells (P > 0.05) (Fig. 4B).

3.4. Effects of DMSA-IRONs on the expression of connexin 43 of cardiomyocytes

It has been proved that the spontaneous contraction of cardiomyocytes also depends on the cell–cell communication.³⁴ Connexin 43 composes gap junctions which provide the chance of directing the electric coupling of cardiomyocytes.²⁰ To certificate the effects of DMSA-IRONs on electric coupling, we determined the distribution and expression of connexin 43 in cardiomyocytes. Connexin 43 is localized in the plasma membrane, especially in the cell–cell contact sites (Fig. 5), which means that gap junctions could form in adjacent α-actinin positive cardiomyocytes in either the control group or the DMSA-IRONs treated groups. No apparent differences in the distribution of connexin 43 among the cardiomyocytes in all groups could be observed (Fig. 5). Meanwhile, a western blotting assay confirmed that the expression of connexin 43 in cardiomyocytes was not affected in by exposure to DMSA-IRONs (Fig. 6).

Fig. 5 The double immunofluorescence of α-actinin (green) and connexin 43 (red) in cardiomyocytes treated with DMSA-IRONs; the nuclei are stained with DAPI (blue). The white arrows indicate the distribution of connexin 43 in cell–cell interfaces of the cardiomyocytes. Scale bar = 40 μm.

Fig. 6 Western blotting of N-cadherin and connexin 43 of cardiomyocytes exposed to different levels of DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹); the GAPDH expression acted as internal control.

3.5. Contractive activity and electrophysiological performance of cardiomyocytes

Previous studies confirmed that the above microstructures partially determined the spontaneous contraction of cardiomyocytes.^35,36 Consistent with the microstructures results, the spontaneous contraction of electromechanical cardiomyocytes was not affected significantly by DMSA-IRONs treatment (ESI video A–D†).

Considering the important electrical conductivity of cardiomyocytes and the few changes of connexin 43 in cardiomyocytes with exposure to DMSA-IRONs, the impacts of DMSA-IRONs on the electrophysiological performance were further assessed by examining the spontaneous calcium transients in cardiomyocytes. That Ca²⁺ fluctuations of cardiomyocytes can be observed and rhythmic Ca²⁺ increases within cardiomyocytes indicates the electrical conduction of cells (Fig. 7A and B). The amplitude of Ca²⁺ transients determined in the control cardiomyocytes was 159 ± 15 nmol L⁻¹; while the amplitudes of cardiomyocytes exposed to 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹ DMSA-IRONs were 231 ± 97 nmol L⁻¹, 150 ± 39 nmol L⁻¹, and 152 ± 21 nmol L⁻¹, respectively (Fig. 7C). There was an insignificant difference between the cardiomyocytes of the control groups and the DMSA-IRONs treated groups. The results reveal that the incubation of DMSA-IRONs does not influence the spontaneous electrical activity of cardiomyocytes. The little influence of DMSA-IRONs on Ca²⁺ fluctuations in cardiomyocytes is convincing of the safety of DMSA-IRONs on contraction of the cardiomyocytes.

Fig. 7 (A): Confocal laser microscopy pictures showing intracellular Ca²⁺ transients of diastole and systole cardiomyocytes; (B): relative absolute intracellular Ca²⁺ levels in cardiomyocytes under different dosages of DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹). (C): Average intracellular Ca²⁺ levels in cardiomyocytes treated by different dosages of DMSA-IRONs (0, 25 μg mL⁻¹, 50 μg mL⁻¹ and 100 μg mL⁻¹).

3.6. Peroxidase-like activity of DMSA-IRONs in vitro and anti-oxidant in cardiomyocytes

ROS content in the microenvironment usually causes damage to cell function and survival. Recently, the peroxidase-like activity of IRONs to break up H₂O₂ was reported.³⁷ To determine the peroxidase-like activity of DMSA-IRONs, we evaluated the catalyzing capability of DMSA-IRONs in the presence of H₂O₂.

Comparing with negative control groups (H₂O₂ + DMSA-IRONs, DMSA-IRONs (100 μg mL⁻¹) + DAB, DAB), the results reveal that DMSA-IRONs reacted with DAB to produce a dark brown color in a dose-dependent manner in an H₂O₂ environment (Fig. 8A). Therefore, the results confirmed that DMSA-IRONs have a peroxidase-like activity that diminishes ROS content in vitro.

Fig. 8 (A): The detection of the peroxidase-like activity of DMSA-IRONs in vitro using DAB as the substrates; (B): representative images and quantification analysis of the intracellular ROS of cardiomyocytes which is detected by DHE staining (red). * = P < 0.05, scale bar = 25 μm; (C): NBT assay used for the determination of the intracellular ROS of cardiomyocytes treated with DMSA-IRONs. * = P < 0.05; (D): intracellular ROS of the cardiomyocytes in an H₂O₂ microenvironment detected by DHE staining (red). The representative images and quantification analysis of intracellular ROS. * = P < 0.05, scale bar = 25 μm; (E): relative intracellular ROS levels of cardiomyocytes exposed to DMSA-IRONs in an H₂O₂ microenvironment through a NBT assay. * = P < 0.05.

IRONs are supposed to scavenge intracellular ROS to promote the survival, proliferation and differentiation of cells.³⁷ To determine the impact of DMSA-IRONs on the intracellular ROS content, cardiomyocytes were examined by both DHE assay (Fig. 8B) and NBT assay (Fig. 8C). With the treatment of DMSA-IRONs, the ROS content of the cardiomyocytes decreased compared with the control group. Meanwhile, the intracellular ROS levels of cardiomyocytes with an incubation of IRONs (25–100 μg mL⁻¹) remained similar (Fig. 8B and C). The results of the DHE and NBT assays indicate that DMSA-IRONs scavenged the intracellular ROS content of cardiomyocytes. Therefore, the decreased ROS content ensured the survival and cell function of the cardiomyocytes. Consistent with the results of the MTT, the live/dead assay, the quantitation of ROS confirmed good biocompatibility of DMSA-IRONs.

To further prove the peroxidase-like activity of DMSA-IRONs in cardiomyocytes, the cardiomyocytes were exposed to H₂O₂, and then the intracellular ROS content was analyzed through DHE staining. The treatment of H₂O₂ increased the intracellular ROS levels of cardiomyocytes, and, comparing with control groups, the cardiomyocytes with intracellular DMSA-IRONs diminished the intracellular ROS levels significantly (Fig. 8D). Meanwhile, the NBT assay further confirmed the peroxidase-like activity to diminish intracellular ROS content in the presence of H₂O₂ (Fig. 8E).

4. Discussion

Magnetic nanoparticles are widely used as contrast agents and for biomedical research. Therefore, it is necessary to confirm the potential effects of IRONs on cells before clinical application. In this study, we reveal the positive effects of supplemented DMSA-IRONs on cardiomyocytes, including cell microstructures and biological activities.

Desirable surface modifications of IRONs with suitable materials, such as non-polymeric organic stabilizers, polymeric organic stabilizers and inorganic materials, could endow them with various physico-chemical properties and may cause specific biological effects.¹ Comparing with the positively charged IRONs–NH₂ and bare IRONs, negatively charged IRONs–COOH has been reported to have a significantly different influence on cell compartments and to alter specific genes involved in cell proliferation.¹⁶ DMSA-IRONs with carboxyl (–COOH) groups were proved to have negative charge in the surface of IRONs.³⁸ In this study, DMSA-IRONs demonstrated negligible toxicity to cardiomyocytes, which was consistent with previous findings that DMSA-modified IRONs were biocompatible with fibroblasts,²³ macrophages,³⁹ and proliferation and differentiation of stem cells.⁴⁰

We observed that the internalized DMSA-IRONs was mainly distributed in the cytoplasm but not in the nuclei (Fig. 2). It was reported that the distribution of IRONs was mainly confined to the cytoplasmic vesicles with no presence in the mitochondria, nucleus or endoplasmic reticulum.^41–43 Furthermore, IRONs are considered to be degraded in lysosome to release iron ions due to the low pH environment,^44,45 and an excess of iron ion was reported to affect the biological activity and levels of intracellular ROS, which may pose effects on the cell plasma membrane and nucleus.⁴¹

The microstructures of cardiomyocytes, including actin filaments, adherens junctions composed of N-cadherin and gap junctions composed of connexin 43, affect cell shape and function of the cardiomyocytes. The F-actin microfilament cytoskeleton was reported to be involved in a variety of cell processes, including the determination of cell morphology, locating of cellular components, the interactions of cell–cell and cell–substrate and the transduction of cell signals.⁴⁶ Neuhuber et al. reported that actin filaments are involved in maintaining cell shape and actin-related signal pathways, such as cell death and migration.⁴⁷ In the evaluation of DMSA-IRONs on actin filaments, no significant changes of F-actin and cell shapes of cardiomyocytes were observed after exposure to DMSA-IRONs, which is consistent with previous reports.^16,48 Meanwhile, the cytoskeleton could anchor with cell adherens junctions, and previous studies indicated that N-cadherin connected with F-actin through β-catenin to modulate the mechanical stress.⁴⁹ Through the double immunofluorescence of F-actin and N-cadherin, the colocalization of N-cadherin and F-actin was not affected by the incubation of DMSA-IRONs (Fig. 4). The normal expression and distribution of N-cadherin supports the formation of adherens junctions and mechanical transduction between cardiomyocytes.⁵⁰

Gap junctions (composed of connexins) between neighboring cardiomyocytes are used for the direct communication and passage of small metabolites and ions.⁵¹ Importantly, gap junctions support the direct electric coupling and transmission of action potentials of cardiomyocytes.²⁰ The normal distribution and expression of connexin 43 in cell–cell junctions of cardiomyocytes without disturbance by DMSA-IRONs is supposed to support various ion transmissions of the cardiomyocytes. Moreover, the assembly of gap junctions is supported by the formation of adherens junctions between cardiomyocytes.⁵² Therefore, the normal formation and distribution of adherens junctions and gap junctions of the cardiomyocytes in a microenvironment with DMSA-IRONs guarantees the interactions between cardiomyocytes. Moreover, intracellular Ca²⁺ transients further confirm the few effects of DMSA-IRONs on the electrical performance of cardiomyocytes (Fig. 6). Our results are consistent with former studies; Au et al. found that IRONs labelling did not impact the intracellular Ca²⁺ levels in embryonic stem cell-derived cardiomyocytes.¹⁴ The small influence of DMSA-IRONs on cell microstructures is convincing of the safety of DMSA-IRONs on cardiomyocytes and supports the normal functional performance of cardiomyocytes.

In terms of the peroxidase-like activity of IRONs, it was supposed to derive from the surface ferrous content of nanoparticles.⁵³ Meanwhile, the affinity between IRONs and substrates is related to the catalytic activity; the higher the affinity of IRONs towards the substrates, the higher is the catalytic activity produced.⁵⁴ Moreover, the electrostatic affinity derived from the superficial charge of IRONs with different surface coatings could enhance catalytic activity.⁵⁴ On the effects of intracellular IRONs with different superficial charge, negatively charged IRONs–COOH could down-regulate the lipid peroxidation related gene compared with the bare IRONs and the positively charged IRONs–NH₂.¹⁶ The peroxidase-like activity of IRONs was supposed to derive from the intact iron core but not the iron ion,⁵⁰ and the break-up of IRONs in the lysosome has been convincing.⁵⁵ It has been proved that the surface modification could decrease the degradation of intracellular IRONs,^1,32 and also it was proved that magnetic nanoparticles reduced the intracellular glutathione due to the peroxidase mimetics.⁵⁶ Therefore, we supposed that the intact DMSA-IRONs in cardiomyocytes kept the peroxidase-like activity to decrease intracellular ROS.

In infarcted myocardium, ROS would be abundantly produced and injure the surrounding myocardium. Florian et al. proved the positive effects of IRONs on ventricular remodelling in patients with myocardial infarction.¹⁸ The peroxidase-similar activity of DMSA-IRONs might exert positive effects to improve the remodeling of infarcted myocardium. The exact mechanisms of DMSA-IRONs on diminishing the intracellular ROS content in cardiomyocytes need further investigation.

5. Conclusions

DMSA-IRONs could be internalized by cardiomyocytes in a dose-dependent manner and have great biocompatibility to cardiomyocytes under the concentration of 0–100 μg mL⁻¹. The microstructures, including actin filaments, N-cadherin composed cell adherens junctions and connexin 43 composed gap junctions of cardiomyocytes, were not affected by DMSA-IRONs. In biological activity, DMSA-IRONs did not affect the intracellular Ca²⁺ transients of cardiomyocytes significantly. Importantly, DMSA-IRONs with peroxidase-like activity were proved to diminish the intracellular ROS levels of cardiomyocytes. This study provides a basic investigation of effects of DMSA-IRONs on cardiomyocytes prior to clinical application.

Acknowledgements

This work was supported by the Key Program of the National Natural Science Foundation of China (no. 31030032), the National Natural Science Funds for Distinguished Young Scholar (no. 31125013), the National High Technology Research and Development Program of China (no. 2012AA020506) and the National Natural Science Foundation of China (no. 31100697).

Notes and references

1. A. K. Gupta and M. Gupta, Biomaterials, 2005, 26, 3995–4021.
2. A. Verma and F. Stellacci, Small, 2010, 6, 12–21.
3. N. Singh, G. J. Jenkins, R. Asadi and S. H. Doak, Nano Rev., 2010, 1, 12–21.
4. S. J. Soenen, U. Himmelreich, N. Nuytten and M. De Cuyper, Biomaterials, 2011, 32, 195–205.
5. D. L. Thorek, A. K. Chen, J. Czupryna and A. Tsourkas, Ann. Biomed. Eng., 2006, 34, 23–38.
6. R. K. Iyer, L. L. Chiu, L. A. Reis and M. Radisic, Curr. Opin. Biotechnol., 2011, 22, 706–714.
7. J. P. Karam, C. Muscari and C. N. Montero-Menei, Biomaterials, 2012, 33, 5683–5695.
8. R. Dash, J. Chung, T. Chan, M. Yamada, J. Barral, D. Nishimura, P. C. Yang and P. C. Simpson, Magn. Reson. Med., 2011, 66, 1152–1162.
9. O. M. Mykhaylyk, N. O. Dudchenko and A. K. Dudchenko, Ukr. Biokhim. Zh., 2005, 77, 80–92.
10. T. Y. Tang, R. R. Moustafa, S. P. Howarth, S. R. Walsh, J. R. Boyle, Z. Y. Li, J. C. Baron, J. H. Gillard and E. A. Warburton, Eur. J. Vasc. Endovasc. Surg., 2008, 36, 53–55.
11. J. V. Terrovitis, J. W. Bulte, S. Sarvananthan, L. A. Crowe, P. Sarathchandra, P. Batten, E. Sachlos, A. H. Chester, J. T. Czernuszka, D. N. Firmin, P. M. Taylor and M. H. Yacoub, Tissue Eng., 2006, 12, 2765–2775.
12. K. Shimizu, A. Ito, J. K. Lee, T. Yoshida, K. Miwa, H. Ishiguro, Y. Numaguchi, T. Murohara, I. Kodama and H. Honda, Biotechnol. Bioeng., 2007, 96, 803–809.
13. Y. Sapir, B. Polyak and S. Cohen, Nanotechnology, 2014, 25, 014009.
14. K. W. Au, S. Y. Liao, Y. K. Lee, W. H. Lai, K. M. Ng, Y. C. Chan, M. C. Yip, C. Y. Ho, E. X. Wu, R. A. Li, C. W. Siu and H. F. Tse, Biochem. Biophys. Res. Commun., 2009, 379, 898–903.
15. Y. Baratli, A. L. Charles, V. Wolff, L. Ben Tahar, L. Smiri, J. Bouitbir, J. Zoll, F. Piquard, O. Tebourbi, M. Sakly, H. Abdelmelek and B. Geny, Toxicol. In Vitro, 2013, 27, 2142–2148.
16. M. Mahmoudi, S. Laurent, M. A. Shokrgozar and M. Hosseinkhani, ACS Nano, 2011, 5, 7263–7276.
17. Y. Amsalem, Y. Mardor, M. S. Feinberg, N. Landa, L. Miller, D. Daniels, A. Ocherashvilli, R. Holbova, O. Yosef, I. M. Barbash and J. Leor, Circulation, 2007, 116, I38–I45.
18. A. Florian, A. Ludwig, S. Rosch, H. Yildiz, S. Klumpp, U. Sechtem and A. Yilmaz, Int. J. Cardiol., 2014, 173, 184–189.
19. M. Takeichi, Curr. Opin. Cell Biol., 1995, 7, 619–627.
20. R. M. Shaw and Y. Rudy, Circ. Res., 1997, 81, 727–741.
21. Y. Lalatonne, L. Motte, J. Richardi and M. P. Pileni, Phys. Rev. E: Stat., Nonlinear, Soft Matter Phys., 2005, 71, 011404.
22. M. Mahmoudi, A. Simchi, M. Imani, M. A. Shokrgozar, A. S. Milani, U. O. Hafeli and P. Stroeve, Colloids Surf., B, 2010, 75, 300–309.
23. M. Auffan, L. Decome, J. Rose, T. Orsiere, M. De Meo, V. Briois, C. Chaneac, L. Olivi, J. L. Berge-Lefranc, A. Botta, M. R. Wiesner and J. Y. Bottero, Environ. Sci. Technol., 2006, 40, 4367–4373.
24. N. Fauconnier, J. N. Pons, J. Roger and A. Bee, J. Colloid Interface Sci., 1997, 194, 427–433.
25. Y.-k. Sun, M. Ma, Y. Zhang and N. Gu, Colloids Surf., A, 2004, 245, 15–19.
26. Y. S. Zhao, C. Y. Wang, D. X. Li, X. Z. Zhang, Y. Qiao, X. M. Guo, X. L. Wang, C. M. Dun, L. Z. Dong and Y. Song, J. Heart Lung Transplant., 2005, 24, 1091–1097.
27. J. Kim, K. S. Choi, Y. Kim, K. T. Lim, H. Seonwoo, Y. Park, D. H. Kim, P. H. Choung, C. S. Cho, S. Y. Kim, Y. H. Choung and J. H. Chung, J. Biomed. Mater. Res., Part A, 2013, 101, 3520–3530.
28. C. Mauritz, K. Schwanke, M. Reppel, S. Neef, K. Katsirntaki, L. S. Maier, F. Nguemo, S. Menke, M. Haustein, J. Hescheler, G. Hasenfuss and U. Martin, Circulation, 2008, 118, 507–517.
29. K. Fan, C. Cao, Y. Pan, D. Lu, D. Yang, J. Feng, L. Song, M. Liang and X. Yan, Nat. Nanotechnol., 2012, 7, 459–464.
30. M. M. Tarpey, D. A. Wink and M. B. Grisham, Am. J. Physiol.: Regul., Integr. Comp. Physiol., 2004, 286, R431–R444.
31. T. R. Pisanic II, J. D. Blackwell, V. I. Shubayev, R. R. Finones and S. Jin, Biomaterials, 2007, 28, 2572–2581.
32. S. J. Soenen and M. De Cuyper, Nanomedicine, 2010, 5, 1261–1275.
33. R. G. Thakar, F. Ho, N. F. Huang, D. Liepmann and S. Li, Biochem. Biophys. Res. Commun., 2003, 307, 883–890.
34. S. B. Geisler, K. J. Green, L. L. Isom, S. Meshinchi, J. R. Martens, M. Delmar and M. W. Russell, J. Biomed. Biotechnol., 2010, 2010, 624719.
35. J. Li, M. D. Levin, Y. Xiong, N. Petrenko, V. V. Patel and G. L. Radice, J. Mol. Cell. Cardiol., 2008, 44, 597–606.
36. M. Noorman, M. A. van der Heyden, T. A. van Veen, M. G. Cox, R. N. Hauer, J. M. de Bakker and H. V. van Rijen, J. Mol. Cell. Cardiol., 2009, 47, 23–31.
37. D. M. Huang, J. K. Hsiao, Y. C. Chen, L. Y. Chien, M. Yao, Y. K. Chen, B. S. Ko, S. C. Hsu, L. A. Tai, H. Y. Cheng, S. W. Wang and C. S. Yang, Biomaterials, 2009, 30, 3645–3651.
38. Y. Ge, Y. Zhang, J. Xia, M. Ma, S. He, F. Nie and N. Gu, Colloids Surf., B, 2009, 73, 294–301.
39. J. Gu, H. Xu, Y. Han, W. Dai, W. Hao, C. Wang, N. Gu and J. Cao, Sci. China: Life Sci., 2011, 54, 793–805.
40. J. Liu, L. Wang, J. Cao, Y. Huang, Y. Lin, X. Wu, Z. Wang, F. Zhang, X. Xu and G. Liu, Nanoscale, 2014, 6, 9025–9033.
41. M. Calero, L. Gutierrez, G. Salas, Y. Luengo, A. Lazaro, P. Acedo, M. P. Morales, R. Miranda and A. Villanueva, Nanomedicine, 2014, 10, 733–743.
42. B. H. Kenzaoui, M. R. Vila, J. M. Miquel, F. Cengelli and L. Juillerat-Jeanneret, Int. J. Nanomed., 2012, 7, 1275–1286.
43. H. Xu, W. Dai, Y. Han, W. Hao, F. Xiong, Y. Zhang and J. M. Cao, J. Nanosci. Nanotechnol., 2010, 10, 7406–7410.
44. E. Okon, D. Pouliquen, P. Okon, Z. V. Kovaleva, T. P. Stepanova, S. G. Lavit, B. N. Kudryavtsev and P. Jallet, Lab. Invest., 1994, 71, 895–903.
45. D. Pouliquen, J. J. Le Jeune, R. Perdrisot, A. Ermias and P. Jallet, Magn. Reson. Imaging, 1991, 9, 275–283.
46. M. Derangeon, N. Bourmeyster, I. Plaisance, C. Pinet-Charvet, Q. Chen, F. Duthe, M. R. Popoff, D. Sarrouilhe and J. C. Herve, J. Biol. Chem., 2008, 283, 30754–30765.
47. B. Neuhuber, G. Gallo, L. Howard, L. Kostura, A. Mackay and I. Fischer, J. Neurosci. Res., 2004, 77, 192–204.
48. A. Hanini, A. Schmitt, K. Kacem, F. Chau, S. Ammar and J. Gavard, Int. J. Nanomed., 2011, 6, 787–794.
49. J. L. Bruses, PLoS One, 2013, 8, e62435.
50. S. Satomi-Kobayashi, T. Ueyama, S. Mueller, R. Toh, T. Masano, T. Sakoda, Y. Rikitake, J. Miyoshi, H. Matsubara, H. Oh, S. Kawashima, K. Hirata and Y. Takai, Hypertension, 2009, 54, 825–831.
51. A. L. Harris, Prog. Biophys. Mol. Biol., 2007, 94, 120–143.
52. H. Zhu, H. Wang, X. Zhang, X. Hou, K. Cao and J. Zou, Mol. Cells, 2010, 30, 193–200.
53. L. Gao, J. Zhuang, L. Nie, J. Zhang, Y. Zhang, N. Gu, T. Wang, J. Feng, D. Yang, S. Perrett and X. Yan, Nat. Nanotechnol., 2007, 2, 577–583.
54. F. Yu, Y. Huang, A. J. Cole and V. C. Yang, Biomaterials, 2009, 30, 4716–4722.
55. A. S. Arbab, L. B. Wilson, P. Ashari, E. K. Jordan, B. K. Lewis and J. A. Frank, NMR Biomed., 2005, 18, 383–389.
56. Y. Ma, Z. Zhang, C. Ren, G. Liu and X. Chen, Analyst, 2012, 137, 485–489.

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Footnotes

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c4ra11079j

‡ These authors contributed equally to this work.

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