Su 70 scanning electron microscope

A commercial ZnO nanopowder (<100 nm) (Sigma-Aldrich, Saint Louis, MO, USA) was used. The structure of the powder was characterized by X-ray diffraction (XRD) using a Panalytical X’Pert Pro Diffractometer (PANalytical B.V., Almelo, The Netherlands) with Cu Kα1 radiation (λCu = 0.154056 nm). The XRD patterns were recorded in the range of 10–80° of 2q values, with a 0.02° step and time per step of 3 s. The powder specific surface area was accessed by gas adsorption (BET isotherm) in a Micromeritics-Gemini V2380 surface area analyser (Micromeritics, Norcross, GA, USA). Scanning electron microscopy (SEM) was performed to analyse the powder morphology using a Hitachi SU-70 Scanning Electron Microscope (Hitachi High-Tech, Tokyo, Japan). The particle surface charge was also accessed by measuring the Zeta potential of ZnO aqueous suspensions at different pHs in Coulter Delsa 440 SX equipment (Beckman Coulter, Indianapolis, IN, USA).

A ZnO nanopowder, supplied by Sigma-Aldrich, Saint Louis, MO, USA, referenced 544906, was used. The powder was characterized in terms of its crystallography, morphology, particle size distribution, and particle surface charge in aqueous suspensions, with a set of techniques that are described below.
For the crystallographic characterization, X-ray diffraction (XRD) using a X’Pert Pro Diffractometer, PANalytical B.V., Almelo, Netherlands, with Cu K_α1 radiation (λCu = 0.154056 nm) was applied. The morphology of the particles was inspected by scanning electron microscopy (SEM) performed in a Hitachi SU-70 Scanning Electron Microscope, Tokyo, Japan, and the specific surface area of the powder was accessed by gas adsorption (BET isotherm) in a Micromeritics-Gemini V2380 surface area analyzer, Micromeritics, Norcross, GA, USA. The particle size distribution (PSD) was determined in a water medium by laser diffraction through a Coulter LS-200 device, Beckman Coulter, Brea, CA, USA, and the particle surface charge was also accessed by measuring the Zeta potential of ZnO aqueous suspensions, at different pH levels, in a Coulter Delsa 440 SX device, Beckman Coulter, Indianapolis, IN, USA.

Infrared (IR) spectra were measured with a Fourier transform infrared (FT-IR) spectrophotometer (Spectrum One B, Perkin Elmer Co., Waltham, MA, USA). The viscoelasticity of the vegetable oil reaction products was measured with an Advanced Rheometric Expansion System (ARES, Rheometric Scientific, Piscataway, NJ, USA). The viscoelasticity measurements were conducted at 25 °C with a plate–plate type rheometer in a frequency sweep mode with the frequency oscillating from 0.1 to 1000 rad/s. Microencapsulation was conducted with a NZ-1000 mechanical agitator (Eyela, Tokyo, Japan). Images of the microcapsules and coatings were obtained with a BX-51 microscope (Olympus, Tokyo, Japan). The microcapsule mean diameter was analyzed from 300 data sets using a CC-12 CCD camera (Olympus, Tokyo, Japan) built into the microscope and TS image analysis software (Olympus, Tokyo, Japan). A SU-70 scanning electron microscope (Hitachi, Tokyo, Japan) was used to observe the microcapsules and coating surfaces.

The P. dentata cells were fixed using 2.5% to 3% concentration of glutaraldehyde with the pH ranging between 6.8 and 7.4; the pH was maintained using phosphate buffer solution 40 . The P. dentata cells were placed in the glutaraldehyde fixative solution for about 30 minutes to ensure complete infiltration of the glutaraldehyde solution into the P. dentata cells. The above fixed P. dentata cells were subjected to gradient dehydration using different concentrations of ethanol (50%, 60%, 70%, 80%, 90%, and 100%). All the subjected P. dentata samples were sequentially soaked in each concentration starting from 50% to 100% for at least 15 minutes, later these samples were preserved in -80 °C freezers until its use. The P. dentata samples were removed from the -80 °C freezer and placed in a freeze dryer (LABCONCO freezone 12, Kansas City, MO, USA) for 24 hours. The samples were then sputtered with gold or carbon and later observed using HITACHI SU-70 Scanning Electron Microscope for imaging, respectively.

SEM samples were prepared as described previously^{34 (link)}. Briefly, samples were fixed in 2.5% paraformaldehyde in 0.1 cabodylate buffer (Electron Microscopy Sciences), dehydrated using an ethanol gradient, incubated in hexamethyldisilazane (Electron Microscopy Sciences), and air-dried overnight. Samples were then coated with 8 nm thick iridium using a sputtering tool (Cressington 208) and imaged using a Hitachi SU-70 scanning electron microscope.

Paraffin embedding of adult flies fixed in phosphate buffered saline (PBS) formalin for 30 min. was performed under vacuum to permit penetration of the medium, followed by 10 um sectioning and staining with von Kossa/methylene green (black stain in upper panels) or for calcium with Alizarin Red (red stain in lower panels) using standard protocols (4X and 40X magnification). Alizarin Red staining of dissected Malpighian tubules was performed in PBS containing 0.1% Triton X100 and 20 mg/ml Alizarin Red dye, followed by bright field and epi-fluorescent micrographs of life Malpighian tubules from flies expressing GFP in principal or stellate cells at 360/670 nm and 360/488 nm (excitation/emission wavelength), respectively. For scanning electron microscopy Malpighian tubules were dissected in PBS containing 5 mg/ml proteinase K to gently remove epithelium and to expose mineral deposits inside the tubule lumen as previously described^{18 (link)}, air-dried on carbon support, coated with 5 nm chromium, and imaged using a Hitachi SU70 scanning electron microscope, followed by energy-dispersive X-ray-spectroscopy (EDX) microanalysis an individual microsphere.