Spectrometer

Fourier transform infrared spectroscopy (FTIR) was performed to identify and determine functional groups of composite and epoxy structures in the infrared (IR) region between 400 cm ${}^{-1}$ and 4000 cm ${}^{-1}$ . The absorption spectra were obtained with a resolution of 4 cm ${}^{-1}$ and 64 scans in each assay. The equipment used was a Thermo Scientific spectrometer, using the OMNIC Spectra software.

The electronic states and chemical composition of NFs were identified by XPS. The samples were prepared by drop casting on glass substrates and allowing drying in the air. XPS measurements were performed on a spectrometer (Thermofisher Scientific, USA) equipped with a monochromatic aluminum source (Al-Kα, hν = 1486.7 eV) using a spot of 400 µm and a hemispherical analyzer, at a take-off angle of 0°. Survey scans were acquired at a pass energy of 200 eV and a step of 1 eV while the narrow scans were acquired at a pass energy of 50 eV and a step of 0.1 eV. A “dual beam” flood gun was used for charge compensation. Data acquisition and interpretation were processed with the Thermo Avantage software, using a peak-fitting routine with Shirley background and symmetrical 70/30% Gaussian/Lorentzian peak shapes and a Doniach–Sunjic type function to fit the Pt metal asymmetric peaks. The homogeneity of samples was checked by measurement on up to 4 different points.

The FT-IR scans were not only used to evaluate the functional groups of graphene and cement-based composite but also to monitor the hydration process of the cement mortars. FT-IR scans are generally used to test for functional groups of graphene, but they can also be used to determine changes in cement mortars’ hydration products. Therefore, in order to study the effect of TiO₂-RGO and GO on cement mortars’ hydration products, in this paper, the composition of samples was investigated by Fourier transform infrared (FT-IR) spectroscopy using a spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) over the wavenumber from 400 to 4000 cm⁻¹.

Transmission electron microscopy (TEM) images were obtained using a HT 7100 microscope (Hitachi, Japan) operated at an acceleration voltage of 120 kV. High-resolution TEM and energy dispersive X-ray spectroscopy (EDS) studies were carried out in JEM-2100F (JEOL, Japan) and Titan G2 ChemiSTEM Cs Probe (FEI, USA) operated at an acceleration voltage of 200 kV. The metal contents in catalysts were determined using inductively coupled plasma-optical emission spectroscopy (ICP-OES, PerkinElmer, Optima 7300DV, USA). X-ray diffraction (XRD) patterns were obtained with a D2 phaser X-ray diffractometer (Bruker, USA). X-ray photoelectron spectroscopy (XPS) was carried out using a spectrometer (Thermofisher Scientific, USA) with Al Kα X-ray (1486.6 eV) as the light source. All spectra were aligned using the C-1s peak at 284.5 eV as reference.

Cytotoxicity was measured by using the LDH assay. Chondrocytes were seeded at a density of 5 × 10⁴ cells/well in 24-well plates, cultured for 24 h, and then treated with SNP (0.5 mM) and 2-APB (100 μM) for 12 h. Firstly, the medium was aspirated (50 μl per well) from 24-well plates into 96-well plates for detection of LDH release and chondrocytes lysed with Triton X-100 for 30 min to obtain maximum LDH release. Then medium was aspirated (50 μl per well) from 24-well containing Triton X-100 into 96-well plates and 50 μl reagent added from the Cytotoxicity Detection Kit (Roche Diagnostics) to each well. The 96-well plate was preincubated for 30 min in the dark, and absorbance detected at 620 and 492 nm using a spectrometer (Thermo Fisher Scientific, United States). LDH release was determined based on subtracting absorbance at 620 nm from that at 492 nm. LDH from lysed cells using Triton X was the maximum LDH release and used as a standardizer to obtain the relative release of LDH. Relative LDH release = LDH release/maximum LDH release.