Acidity is an important physicochemical property of atmosphericaerosols, but its effects on the ROS formation from SOA have beenpoorly characterized. By applying the electron paramagnetic resonancespin-trapping technique and the Diogenes chemiluminescence assay,we find highly distinct radical yields and composition at differentpH values in the range of 1–7.4 from SOA generated by oxidationof isoprene, α-terpineol, α-pinene, β-pinene, toluene,and naphthalene. We observe that isoprene SOA has substantial hydroxylradical (•OH) and organic radical yields at neutralpH, which are 1.5–2 times higher compared to acidic conditionsin total radical yields. Superoxide (O2•–) is found to be the dominant species generated by all types of SOAsat lower pH. At neutral pH, α-terpineol SOA exhibits a substantialyield of carbon-centered organic radicals, while no radical formationis observed by aromatic SOA. Further experiments with model compoundsshow that the decomposition of organic peroxide leading to radicalformation may be suppressed at lower pH due to acid-catalyzed rearrangementof peroxides. We also observe 1.5–3 times higher molar yieldsof hydrogen peroxide (H2O2) in acidic conditionscompared to neutral pH by biogenic and aromatic SOA, likely due toenhanced decomposition of α-hydroxyhydroperoxides and quinoneredox cycling, respectively. These findings are critical to bridgethe gap in understanding ROS formation mechanisms and kinetics inatmospheric and physiological environments.Reactive oxygen species(ROS) play a critical role in the chemicaltransformation of atmospheric secondary organic aerosols (SOA) andaerosol health effects by causing oxidative stress  The subsequent reactions of •OH withprimary or secondary alcohols can lead to the generation of superoxidevia decomposition of α-hydroxyperoxy radicals.23 Limited studies have investigated the effects of pH onROS formation. Enami24 reported that lowerpH promotes the decomposition of α-hydroxyhydroperoxides leadingto the formation of H2O2. Tong et al.25 showed significant enhancement in radical formationwith highly acidic (pH 0–1) conditions in the mixtures of isopreneSOA and mineral dust, which could be due to enhanced Fenton(-like)reactions in the presence of transition metals in the dust. Our recentstudy demonstrated substantial formation of organic radicals fromiron-facilitated decomposition of organic peroxides contained in SOAin surrogate epithelial lining fluid with a pH of 7.4.26 As there have been few systematic investigationsof pH effects on ROS formation, it is still highly uncertain how differentpH values would affect ROS formation from SOA, and the underlyingchemical mechanism is poorly understood.Reactive oxygen species(ROS), including hydroxyl radicals : highly acidic (1.0), moderately acidic(2.5–3.5), and neutral (7.4) conditions. We observed that pHimpacts yields and the composition of ROS depending on SOA types:isoprene and α-terpineol SOA are found with significantly higherROS formation at neutral pH; α-pinene, β-pinene, tolueneand naphthalene SOA generate more superoxide in acidic conditions.Further, we revealed using model compounds that the radical generationby organic peroxide decomposition can be suppressed under lower pH.In contrast, acidic conditions consistently promote H27 was used togenerate SOA particles from •OH photooxidation ofisoprene , α-terpineol , α-pinene , β-pinene, toluene ,and naphthalene . Detailed proceduresof SOA formation can be found in our recent studies.26 While the PAM reactor applies high levels of oxidants 28 with a short reaction time, the PAM-generatedSOA have been found to have similar characteristic with ambient andchamber-generated SOA in terms of mass yield, oxidation state, hygroscopicity,and chemical composition with similar mass spectra measured by anAerodyne ToF AMS.31 The relative humidity in the PAM reactor was 40–50%.A scanning mobility particle sizer  wasused to record the particle size distribution. SOA particles werecollected on 47 mm poly(tetrafluoroethylene) (PTFE) filters  for 60–120 min with averagemass loadings of 0.45 ± 0.04, 1.19 ± 0.26, 0.73 ± 0.20,0.67 ± 0.10, 2.52 ± 0.50, and 0.43 ± 0.12 mg for isoprene,α-terpineol, α-pinene, β-pinene, toluene, and naphthaleneSOA, respectively. The filter samples were extracted into 1 mL of10 mM spin-trap solutions with preadjusted pH  for 7 min. The filters after extraction were dried under nitrogenflow for 10–20 min. The mass difference before and after theextraction was considered as the amount of SOA dissolved in the solution,and an average molar mass of 200 g mol–121 was assumed to calculate the SOA molar concentrationsin filter extracts. SOA concentrations were in the range of 1.9–2.5,4.8–7.8, 2.6–5.2, 2.6–4.0, 9.2–15.7, and1.2–2.5 mM for isoprene, α-terpineol, α-pinene,β-pinene, toluene, and naphthalene SOA, respectively. Two SOAsamples were prepared for each pH for the quantification of radicals.The radical formation under different pH values was also quantifiedusing several model compounds including cumene hydroperoxide , tert-butyl hydroperoxide , 5-hydroxy-1,4-naphthoquinone ,and ascaridole .A potentialaerosol mass (PAM) reactor11 pH was adjusted to 1.0 by adding sulfuric acid in the spin-trap solution. The moderately acidiccondition is in line with aerosols containing a lower amount of sulfate,biomass burning aerosols,32 and the lowerend of cloud/fog water droplets.15 Theoriginal pH of the SOA extracts varied from 2.5 to 3.5, representinga moderately acidic condition. The neutral pH is of interest for clouddroplets as well as the physiological environment upon inhalationand respiratory deposition of SOA.15 Aphosphate-buffered saline  was used to adjustthe pH at 7.4 in the SOA extracts. SOA particles were extracted intoa spin-trap solution with preadjusted pH. The pH of the model compoundswas maintained the same way for the highly acidic (sulfuric acid)and neutral (PBS) conditions, while a smaller amount of sulfuric acidwas used to reach the moderately acidic condition (pH = 3.0). A pHmeter (VWR sympHony) was used to measure the pH of the reagents.TheSOA-extracted solutions were maintainedat highly acidic (pH = 1.0), moderately acidic (pH = 2.5–3.5),or neutral (pH = 7.4) conditions. The highly acidic condition mimicsthe ambient internally mixed particles of sulfate and organics.tert-butoxycarbonyl-5-methyl-1-pyrroline-N-oxide (BMPO) . Afterparticle extraction into 1 mL of 10 mM BMPO solutions, a 50 μLaliquot of the SOA extracts was loaded into a 50 μL capillarytube (VWR) and inserted in the resonator of the EPR spectrometer at10, 20, 60, 90, and 120 min from the start of aqueous reactions. Theparameters for EPR measurements are as follows: a center field of3515.0 G, a sweep width of 100.0 G, a receiver gain of 30 dB, a modulationamplitude of 1.0 G, a scan number of 10–50, attenuation of12 dB, a microwave power of 12.6 mW, a modulation frequency of 100kHz, a microwave frequency of 9.86 GHz, and a conversion time andtime constant of 5.12 ms. After obtaining the EPR spectra, SpinFitand SpinCount methods embedded in the Bruker Xenon software were appliedto quantify BMPO-radical adducts at each time point.21A continuous-wave electron paramagneticresonance (CW–EPR) spectrometer  coupled witha spin-trapping technique was applied to quantify the free-radicalformation in the aqueous phase. The free radicals were captured bya spin-trapping agent 5-in situ UV irradiationsystem  equipped with a 100 W Hg lamp wasused with EPR to characterize the radical formation upon illumination.The lamp was usually warmed up for roughly 10 min before the startof any irradiation experiments. A safety shutter between the lampand the resonator was used to control the start and stop of irradiation.A liquid light guide focused the light to the EPR resonator wheresamples were exposed to UV to visible light with a wavelength rangeof 220–600 nm. To test the pH effect on BMPO trapping efficiencies,1 mM H2O2 was mixed with 10 mM BMPO at pH 7.4,3.0, and 1.0 and then placed into the irradiation system, where H2O2 can be photolyzed to form •OH. The background spectrum was recorded at the starting point, withthe shutter raised after the first EPR measurement was finished (∼1min). Temporal measurements were then conducted every minute for 10min to monitor the change in BMPO-OH concentrations over time.In addition, an 33 was applied for the H2O2 measurement using a fluorometric H2O2 assaykit . Detailed procedures of assay preparationcan be found in our previous study.23 TheH2O2 formation was quantified within 2 h fromthe preparation of working solutions due to the instability of theprobe. A calibration was performed using H2O2 standard solutions with concentrations ranging from 0.05 to 1.5μM in PBS to maintain pH at 7.4 . The reaction vials consisted of 2.94 mL sample (Milli-Qwater + filter extracts + PBS) and 60 μL working solution. Thedilution factors were adjusted for different SOA samples so that thefinal H2O2 concentrations would be below 1.5μM. All H2O2 measurements were conductedwith a filter blank with the same dilution factor as the samples.The addition of working solution to the samples was considered asthe start of the reaction, and the measurement was conducted afterthe reaction vials were incubated at the room temperature of 298 Kfor 15 min. A spectrofluorophotometer  was usedto measure the fluorescence of the reagents at excitation and emissionwavelengths of 540 and 590 nm, respectively.A modified protocol2•– emit flash chemiluminescencesignal proportional to the O2•– production rate.34 A Microplate Reader was used to measure the chemiluminescence in a relativelight unit (RLU). To convert RLU to O2•– production rate, the Diogenes assay was calibrated by the EPR spectrometerusing a standardized cell-free O2•– generation system—the hypoxanthine (HX) and xanthine oxidase(XO) system. The oxidation process of HX catalyzed by XO can passelectrons to dissolved oxygen to form O2•–.35 A spin probe CMH  was used to react with O2•– to form nitroxide radical CM• that can be quantified by EPR.36 Concentrations of O2•– at different time points were obtained by simulating the CM• spectra and then used to calculate the O2•– production rate. The detail of the calibrationis provided in Supporting Information, andthe calibration curve is shown in Figure S2. The O2•– production rates fromsix SOA were measured using the Diogenes method. The SOA samples weredirectly extracted in 1 mL PBS, after which 135 μL of the SOAextracts were added to Diogenes to initiate the reaction. Two sampleswere used for each SOA, while a filter blank was used for blank correction.The chemiluminescence measurement was conducted from 1 to 10 min afterthe extraction was completed. The first data points with the reactiontime up to 2 min were used to calculate initial O2•– production rates. The chemiluminescence signalswere observed to decrease over 10 min, and we integrated O2•– production rates to estimate the cumulativeO2•– production.A Diogenes chemiluminescenceassay was applied to quantify the superoxide formation from SOA atneutral pH of 7.4. The reaction products between the Diogenes probeand OFigure S3, the simulated EPR spectrareproduced the observed spectra very well with small residuals. Thesolid color bars in •OH (45%) and carbon-centeredorganic radicals (44%) with a very minor contribution from oxygen-centeredorganic radicals (8%), while α-terpineol SOA shows dominantcarbon-centered radical formation at neutral pH. In comparison, bothisoprene and α-terpineol SOA produce O2•–/HO2• (50–60%) predominantlyin acidic conditions, while •OH (10–20%)and organic radicals (15–33%) only constitute minor fractions,as consistent with our recent study.26 Itshould be noted that the highly acidic condition (pH = 1.0) does notlead to notable differences in radicals yields and relative abundancecompared to the original SOA extracts with moderately acidic conditions(pH = 3.0–3.5).Figure S4. α-pinene and β-pinene SOA (2•–/HO2• (>60%) is observed at pH 1.0 and 3.0, similar to isoprene andα-terpineolSOA. For aromatic  SOA , the predominant form of superoxide in acidic conditions(pH 3.0 and 1.0) should be HO2•, whereasit is O2•– at neutral pH.37 It has been reported that a nitrone spin trapcan react with HO2• very efficiently,while the trapping of O2•– isa two-step process via the initial addition of O2•– to BMPO to form the BMPO-O2– adductfollowed by protonation by water (or other acidic sources) to formBMPO-OOH.38 As a consequence, the overallrate of O2•– trapping in neutralconditions can be an order of magnitude slower compared to HO2• trapping in acidic conditions.39 Hence, we applied the Diogenes chemiluminescenceassay which is more sensitive in superoxide measurements at neutralpH . 2•–/HO2• production rates by SOA at neutralpH. All biogenic  SOA show positive superoxide production rates, varyingfrom 0.005 to 0.013 μM min–1. In contrast,toluene and naphthalene SOA do not generate O2•–/HO2• above the detection limit, asconsistent with the EPR-spin-trap method , the most common standard system for •OH generation, is unsuitable for the assessment of pH effects onBMPO trapping efficiencies as this reaction is known to be intrinsicallyaffected by pH with higher acidity favoring •OHformation.422O2 andefficient trapping of •OH by BMPO. The sharp declinein [BMPO-OH] after 5 min is likely due to photolytic decay of BMPO-OH.Only marginal differences (<20%) were observed in BMPO-OH concentrationsover the course of reactions for different pH conditions, confirmingthat acidity has minor to negligible effects on the BMPO trappingof •OH. We also speculate that pH effects on BMPOtrapping R• and RO• should betrivial as no H+ is involved in the reactions. This investigationof the potential pH effects on the trapping efficiencies of BMPO shouldelicit precaution for future studies using the EPR-spin-trap methodespecially for detecting superoxide at neutral pH.We also investigated if pH affects the BMPO trappingefficiencyof •OH formation (70–90%)from 10 mM cumene hydroperoxide (CHP) and tert-butylhydroperoxides at neutral pH, with total radical yields up to 0.014and 0.04%, respectively. The unimolecular decomposition of labileorganic hydroperoxides can lead to •OH formationthrough the cleavage of the weaker O–O bond.44 In acidic solutions (pH 3.0 and 1.0), however, both organic hydroperoxidesgenerate much lower •OH .While the first-order decomposition of peroxides should be a thermalprocess depending on temperature instead of pH, it may be suppressedat higher acidity due to the acid-catalyzed rearrangement of alkylhydroperoxides.46 Levin et al.47 also characterized the kinetics of acid-catalyzed cleavageof CHP leading to phenol and acetone formation, which can take placeat or even below room temperature in the presence of sulfuric acid.Further, their study47 provides thermodynamicevidence that the thermal decomposition of CHP forming phenol/acetonefollows a combined linear-exponential function of sulfuric acid concentration at room temperature. This alternative decompositionpathway would lead to alcohol and ketone formation as the end products,involving no radical formation.48 A similarmechanism has also been shown for aliphatic alkyl hydroperoxides including tert-butyl hydroperoxide.49 Therefore,it may partially account for the decreased radical formation by isopreneand α-terpineol SOA at lower pH (tert-butyl hydroperoxides. The majorcontribution from •OH by isoprene SOA is likelydue to its higher fraction of organic hydroperoxides (3–5%)compared to α-terpineol SOA (1–3%) as predicted by kineticmodeling in our previous study.23To better understand the pH effectson the ROS formation mechanism from SOA, we measured radical formationfrom commercially available organic hydroperoxides at different pHvalues. lower pH 2a,b, alt50 recentlyproposed that rapid autoxidation during α-pinene ozonolysismay lead to the formation of endoperoxides through ring opening andhydrogen shift reactions. While common organic peroxides  can be stable under room temperatureand do not decompose into organic radicals,26 the radical formation from endoperoxides has not been investigated.The decay of ROOR may potentially serve as a plausible channel, asthe RO• generated from the O–O cleavage canrapidly undergo isomerization or decomposition to form R•.52 Therefore, we characterized the ROS formationfrom a commercially available endoperoxide, ascaridole. However, weobserved no radicals above the detection limit , indicating that endoperoxides are not responsiblefor the organic radical formation from α-terpineol SOA, or thereactivity of ascaridole is lower compared to endoperoxides containedin α-terpineol SOA. Meanwhile, it may be possible that •OH can abstract H from the tertiary alcohol group inα-terpineol SOA to form RO•, which undergoesβ-scission to form R•.51 While this mechanism has been demonstrated for the tertiaryalcohol group in citric acid,53 furtherstudies are warranted as •OH oxidation of monoterpenealcohol has been rarely studied.Further, the decomposition of organic hydroperoxides is unlikelyto account for the exclusive formation of organic radicals by α-terpineolSOA as measured by EPR. Iyer et al.55 The pH dependenceof the quinone redox cycling has been rarely discussed in the contextof ambient PM, so we measured radical formation in the mixture of0.2 mM 5-hydroxy-1,4-naphthoquinone  and 0.2 mM ascorbate.Note that 5-H-1,4-NQ alone did not generate radicals above the detectionlimit. 40 which follows that increasing pH causes a decreasein the redox potential.56 This providesa thermodynamic explanation on favorable O2•–/HO2• formation through stronger quinoneredox cycling in acidic conditions compared to neutral pH  compared toother SOA (<2.0%) with the H2O2 level (4.2%)in the original extract (pH 3.5) in excellent consistency with ourprevious study (4.3 ± 0.4%).23 NaphthaleneSOA shows the second-highest H2O2 yields (1.4–2.0%),which is comparable with Liu et al.61 (1.9–2.5%).Qiu et al.62 recently proposed that thedecomposition of α-hydroxyalkyl-hydroperoxides (α-HHs)is a proton-catalyzed process associated with H2O2 formation, which is a highly plausible mechanism accounting forthe elevated H2O2 yields from biogenic SOA.They showed that the decay rates of α-HHs derived from α-terpineolincrease drastically from 0.29 × 10–3 to 12× 10–3 s–1 when pH decreasesfrom 5.7 to 3.3.62 For toluene and naphthaleneSOA, the enhanced superoxide formation with higher acidity may subsequentlylead to H2O2 yields because O2•– is known as an important precursor of H2O2.64 Given the lowO2•– formation , α-HHsundergo acid-catalyzeddecomposition forming carbonyls and H2O2 yields from SOA are shown to be 25–100 times and 5–8times higher than the total radical yields in acidic and neutral conditions,respectively, which indicates H2O2 as the mostabundant exogenous ROS in ambient PM especially considering its muchlonger lifetime. Under neutral conditions, organic hydroperoxidescan preferably undergo unimolecular decomposition to generate highlyreactive •OH radicals, which can initiate a cascadeof reactions to propagate further radical formation74 as well as directly attack biological components to inducepathological processes such as lipid peroxidation.77 The formed organic radicals can be persistent even in thepresence of antioxidants,26 although theircapacity to cause oxidative potential still warrants further studies.While this study used the PAM reactor to generate SOA, further experimentsare necessary with SOA generated in an environmental chamber thatapplies lower VOC and oxidant concentrations as well as with organicparticles collected from the ambient atmosphere to consolidate theatmospheric and health relevance of acidity effects on ROS formationby SOA.Inhalation and deposition of organicaerosols can lead to oxidativestress by the formed ROS at physiological pH. H


Redox status assessments are time-consuming, require a large volume of samples and great reagent amounts, and are not adequately described for methodological reproducibility. Here, the objective was to standardize redox balance determination, based on previously described spectrophotometric tests in pregnant rats, to improve precision, time dispensed, and the volume of samples and reagents, while maintaining accuracy and adequate cost benefits. This protocol summarizes oxidative stress markers, which focus on spectrophotometric tests for the assessment of thiobarbituric acid–reactive substances, reduced thiol groups, and hydrogen peroxide, as well as the antioxidant activity of superoxide dismutase, glutathione peroxidase, and catalase in washed erythrocyte and serum samples from full-term pregnant rats. For non-pregnant rats and other species, it is necessary to standardize these determinations, especially the sample volume. All measurements were normalized by the estimated protein concentrations in each sample. To establish optimum conditions for the reproducibility of the proposed methods, we describe all changes made in each assay’s steps based on the reference method reassessed for the new standardizations. Furthermore, the calculations of the concentrations or activities of each marker are presented. Thus, we demonstrate that the analysis of serum samples is easier and faster, but it is impossible to detect catalase activity. Furthermore, the proposed methods can be applied for redox balance determination, especially using smaller reagent amounts and lower sample volumes in lesser time without losing accuracy, as is required in obtaining samples during rat pregnancy. Reactive oxygen species (ROS) are free radicals; their non-radical intermediates, produced in a limited amount in response to physiological stimuli, mediate inter- and intracellular signaling . Hydroxy2-), which is mainly generated by mitochondria and the endoplasmic reticulum (2- to hydrogen peroxide (H2O2). Hydrogen peroxide is not a free radical, making it less reactive than the superoxide anion. However, it comes under the term ROS as it is intimately involved in the generation and detoxification of free radicals. H2O2is, in turn, detoxified to water by the enzymes catalase (CAT) and glutathione peroxidase (GSH-Px)  . An imba(GSH-Px) .The redox status is an important measurement to evaluate pathophysiological mechanisms involved in several disorders in pregnancy  and 2012Other studies must be performed to develop more accurate and low-cost techniques that are relevant for oxidative stress evaluation. Furthermore, investigators need blood samples for the determination of different biomarkers during pregnancy in rats. However, there are many methodologies that use a large volume for each marker, making it difficult to determine the different biomarkers in the same animal. Therefore, our objective was to standardize the redox balance determination based on previously described spectrophotometric tests in pregnant Wistar/Sprague-Dawley rats, to improve precision, time dispensed, and the volume of samples and reagents concentrations, while maintaining accuracy and adequate cost-benefits. Additionally, sample collection and processing steps are described in detail to provide reproducibility.®, catalog number: 6610)Paper towel 50 × 50 cm filter paper Laboratory film 1.5 mL microtube 2.0 mL microtube 0.1–10 μL tip 2–200 μL tip 50–1,000 μL tip 96-well microplate Quartz cuvettes 10 mL volumetric flask 50 mL volumetric flask 100 mL volumetric flask 250 mL volumetric flask 600 mL volumetric flask 1,000 mL volumetric flask 10 mL beaker 100 mL beaker 250 mL beaker 500 mL beaker 1,000 mL beaker Glass sticker 1,000 mL Amber bottle 2-mercaptoethanol (purity ≥99%) 2-thiobarbituric acid  5,5-dithiobis (2-nitro-benzoic acid)  5-sulfosalicylic acid hydrate (purity ≥95%) Bovine serum albumin (BSA)  Calcium chloride Ethylenediaminetetraacetic acid  Glutathione reductase  Horseradish peroxidase (HRP)  L-glutathione reduced   Magnesium chloride Phenol red Phosphate buffered saline  Potassium cyanide (purity ≥97%) Potassium ferricyanide (purity ≥99%) Sodium hydroxide   Sodium phosphate monobasic monohydrate ®, catalog number: N1630)β-nicotinamide adenine dinucleotide 2′-phosphate reduced tetrasodium salt hydrate  Protein assay dye reagent concentrate (Bio-RadTM(purity ≥99.9%) TRIS-Ultrapure®, catalog number: A1028.01.BJ)Hydrochloric acid  Pyrogallic acid (purity ≥99.9%)  Di-potassium hydrogen phosphate trihydrate Ethyl alcohol   Sodium bicarbonate Potassium chloride  Sodium chloride   Hydrogen peroxide  Potassium dihydrogen phosphate  Sodium phosphate dibasic (NaStabilizing solution (2.7 mM EDTA and 0.7 mM 2-mercaptoethanol) (see Recipes)Drabkin’s solution (see Recipes)1 μg/μL stock solution (see Recipes)3% (w/v) 5-sulfosalicylic acid hydrate (for 100 samples) (see Recipes)Thiobarbituric acid solution (TBA) 0.67% (for 200 samples) (see Recipes)1 M HCl (see Recipes)0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0  (see Recipes)10 mM DTNB (for 250 duplicate washed erythrocytes samples and 100 duplicate serum samples) (see Recipes)2HPO4, 0.1 M Na2HPO4(see Recipes)Solution 1: 1.71 M NaCl, 34 mM KCl, 14 mM K2(see Recipes)Solution 2: 90 mM CaCl2(see Recipes)Solution 3: 0.11 M MgClSolution 4: 0.1% phenol red (see Recipes)Phosphate buffer solution A (see Recipes)Peroxidase solution (HRP) (see Recipes)Buffer solution b (see Recipes)1 M NaOH (see Recipes)1 M TRIS/HCl 5 mM EDTA pH 8.0  (see Recipes)10 mM pyrogallol solution  (see Recipes)50 mM TRIS/HCl 5 mM EDTA  (see Recipes)3and 1 mM EDTA (see Recipes)1% NaHCO10 mM HCl solution (see Recipes)0.1 M glutathione reduced (GSH)  (see Recipes)10 U/mL glutathione reductase (GSH-Rd)  (see Recipes)t-butyl hydroxiperoxide  (see Recipes)7 mM4 mM NADPH  (see Recipes)2PO4·H2O  (see Recipes)0.1 M NaH2O2) (see Recipes)1 M hydrogen peroxide 0.1–2.5 μL micropipette 0.5–10 μL micropipette 2–20 μL micropipette 10–100 μL micropipette 20–200 μL micropipette 100–1,000 μL micropipette Centrifuge Water purifier Microplate reader Spectrophotometer Digital microplate mixer Water bath Analytical balance -80 °C ultra-freezer Refrigerator Vortex agitator mixer Magnetic agitator pH meter Autoclave Fume hood (Permution®)Gen5 (Bio Tek®)UVProbe (Shimadzu®)Statistica (StatSoft®)Microsoft Office 365 (MicrosoftSample collectionCollect whole blood samples from pregnant rats at term through a cut at the end of the tail in both dry/free of anticoagulant (approximately 5.0–9.0 mL of blood) and heparinized (approximately 1.5 mL of blood in 0.2 mL of heparin) tubes. These volumes of blood are sufficient to process samples for oxidant and antioxidant biomarker measurements.Store under refrigeration at 4–8 °C for 30 min after collection.Note: The samples cannot be stored on wet ice.Use the heparinized tubes to obtain washed erythrocyte samples :× gfor 10 min at 4 °C.Centrifuge the blood samples in heparinized tubes at 185Discard the plasma (light yellow supernatant) and the buffy coat .Notes:1) If the phases do not separate appropriately, repeat centrifuging. If the phases do not separate again, discard the sample.2) If there is high lipid concentration in the samples, the plasma color will be white.Add 2 mL of PBS 1× (pH 7.4) to the erythrocyte samples for washing.× gfor 1 min at 4 °C.Centrifuge at 1,575Discard the supernatant (translucid supernatant).Repeat procedures A3c–e three times.Identify three microtubes for Hb, TBARS, and SH level measurements :i) Pipette 50 μL of washed erythrocyte samples (pellet obtained from previous steps) directly in each microtube  for later measurements of Hb, TBARS, and SH levels.ii) Add 950 μL of purified water in each microtube containing 50 μL of the washed erythrocyte.iii) Gently homogenize the samples by inversion.iv) Store aliquots at -80 °C until assays (stable for up to six months).2O2, GSH-Px, and CAT level measurements  Pipette 50 μL of the washed erythrocyte samples (pellet obtained from previous steps) directly in each microtube  for activity measurements of Hb, SOD, Hii) Add 950 μL of the stabilizing solution (see Recipe 1) in each microtube.iii) Gently homogenize the samples by inversion.iv) Store the aliquots at -80 °C until assays (stable for up to six months).Serum sample collection in dry/free anticoagulant tubes , SOD (5 μL), and GSH-Px (5 μL) using specific identification in each microtube.Pipette the serum aliquots directly into six microtubes for protein quantification (20 μL), TBARS (550 μL), SH (10 μL), HStore at -80 °C until analysis (stable for up to six months).Hemoglobin (Hb) quantification in the washed erythrocytesTake identified microtubes with Hb [diluted with purified water (step A3g) and stabilizing solution (step A3h)].Note: Hemoglobin concentration measurements are similar in purified water and/or stabilizing solution because they are necessary for the determination of the normalization of oxidant and antioxidant concentrations. Thus, two microplates will be used for Hb determination (one for the purified water and another for the stabilizing solution).Homogenize the samples in the vortex agitator mixer.Pipette 200 μL of Drabkin’s solution (see Recipe 2) in two wells of the microplate as a blank.Pipette 197.5 μL of Drabkin’s solution and 2.5 μL of samples (washed erythrocytes) in each well of the microplate (in duplicates).Cover the microplate and mix using a microplate mixer for 1 min at 750 rpm for homogenization.Rest the microplate for 10 min at 25 °C.Lightly tap the sides of the microplate several times to gently mix.Measure the absorbance at a wavelength of 546 nm using Gen5 software.Calculate Hb concentrations .Quantification of protein concentrations in serum samples by Bradford’s method Dilute one part of the serum samples in 100 parts of purified water in a microtube.Note: Other dilutions can be tested according to the experiment.Identify six microtubes for the BSA solutions: 0, 0.1, 0.2, 0.4, 0.8, and 1.0 μg/μL.Prepare the BSA solutions for a standard curve setting from the 1 μg/μL stock solution (see Recipe 3), pipetting the following volumes of BSA and purified water:Dilute the protein assay dye reagent concentrate in a ratio of one part of the reagent to four parts of purified water to prepare Bradford’s reagent.For example: dilute 10 mL of the protein assay dye reagent concentrate in 40 mL of purified water.Note: Calculate the quantity for Bradford’s reagent according to the number of wells that will be used.Pipette 190 μL of Bradford’s reagent into each well of the microplate (standard curve and sample wells).Pipette 10 μL of each BSA solution into the standard curve wells (in duplicates). Homogenize with the tip.Pipette 10 μL of the serum samples into other wells (in duplicates). Homogenize with the tip.Measure the absorbance at a wavelength of 595 nm using Gen5 software.Calculate total protein concentrations .Thiobarbituric acid reactive substances (TBARS)Prepare 3% (w/v) 5-sulfosalicylic acid hydrate (see Recipe 4) and TBA 0.67% (see Recipe 5) solutions.Add 500 μL of serum or washed erythrocytes to a test microtube containing 500 μL of 3% 5-sulfosalicylic acid hydrate.Vortex the microtube for 10 s.× gfor 3 min at 4 °C.Centrifuge the microtube at 18,000Rest the microtube for 15 min at 25 °C.Pipette the supernatant and store it in new microtubes.Note: Avoid pipetting the brown granules.Add 500 μL of purified water in a tube as a blank.Add 500 μL of the supernatant to 500 μL of the TBA solution in a tube for each sample.Vortex the tubes for 10 s.Heat the mixtures at 95 °C for 30 min in a water bath with the tubes partially sealed.Cool on ice for 10 min to stop further reactions.Do not exceed 30 min for the next steps (a–e):Equilibrate the blanks and samples at 25 °C.Pipette 300 μL of purified water in two wells as a blank.Pipette 300 μL of the samples into each well of the microplate (in duplicates).Lightly tap the sides of the microplate several times to gently mix.Measure the absorbance at a wavelength of 535 nm using Gen5 software.Calculate TBARS concentrations .Reduced thiol groups (-SH)Prepare 0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 solution (see Recipe 7).Prepare the 10 mM DTNB solution (see Recipe 8).For the -SH quantification in the washed erythrocytes:Pipette 200 μL of 0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 in two wells as a blank.Pipette 195 μL of 0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 solution into other wells (in duplicates).Pipette 1.0 μL of the washed erythrocyte (in duplicates except in blank wells).Pipette 4.0 μL of the 10 mM DTNB solution into the sample wells (in duplicates).Cover the microplate and mix using a microplate mixer for 10 min at 750 rpm for homogenization.Measure the absorbance at a wavelength of 412 nm.For the -SH serum quantification:Pipette 200 μL of 0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 in two wells as a blank.Pipette 175 μL of 0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 solution into other wells (in duplicates).Pipette 5.0 μL of serum (in duplicates except in blank wells).Pipette 20 μL of the 10 mM DTNB solution into the sample wells (in duplicates).Mix the microplate using a microplate mixer at 750 rpm for 10 min.Measure the absorbance at a wavelength of 412 nm using Gen5 software.Calculate -SH concentrations .2O2)Hydrogen peroxide (HPrepare the solutions below:Solution 1 (see Recipe 9).Solution 2 (see Recipe 10).Solution 3 (see Recipe 11).Solution 4 (see Recipe 12).Phosphate buffer solution A (see Recipe 13).Peroxidase solution (HRP) (see Recipe 14).Prepare buffer solution B (see Recipe 15).Pipette 110 μL of the buffer solution B into two wells of the microplate as a blank.2O2quantification in the washed erythrocytes:For the HPipette 5 μL of the washed erythrocyte sample into other wells (in duplicates).Pipette 105 μL of the buffer solution B into the sample wells.Mix the microplate using a microplate mixer at 750 rpm for 10 s.Incubate the microplate at 37 °C for 10 min.Pipette 10 μL of 1 M NaOH (see Recipe 16) to stop the reaction.Measure the absorbance at a wavelength of 620 nm.2O2quantification in the serum samples:For the HPipette 10 μL of the serum sample into other wells (in duplicates).Pipette 100 μL of the buffer solution B into the sample wells.Mix the microplate using a microplate mixer at 750 rpm for 10 s.Incubate the microplate at 37 °C for 10 min.Pipette 10 μL of 1 M NaOH to stop the reaction.Measure the absorbance at a wavelength of 620 nm using Gen5 software.2O2concentrations .Calculate HSuperoxide dismutase (SOD)Prepare 1 M TRIS/HCl 5 mM EDTA pH 8.0 solution (see Recipe 17).Prepare 10 mM pyrogallol solution (see Recipe 18).Pipette 147 μL of 1 M TRIS/HCl 5 mM EDTA pH 8.0 solution into two wells of the microplate as a blank.Pipette 144 μL of 1 M TRIS/HCl 5 mM EDTA pH 8.0 into other wells (in duplicates).Pipette 3.0 μL of the washed erythrocyte/serum (in duplicates except in the blank wells).Lightly tap the sides of the microplate several times to gently mix.Add 3.0 μL of 10 mM pyrogallol solution into all wells including the blank .Lightly tap the sides of the microplate several times to gently mix.Measure the absorbance during the time intervals of 0 (T0) and 10 (T10) min at 25 °C at a wavelength of 420 nm.Calculate SOD concentrations using Gen5 software .Glutathione peroxidase (GSH-Px or GPx)Prepare the solutions below:Note: All solutions (except 50 mM TRIS/HCl 5 mM EDTA pH 7.6 and 7 mM t- butyl hydroperoxide) and reagents must be kept on ice until the moment of the assay.0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 solution (see Recipe 7).50 mM TRIS/HCl 5 mM EDTA pH 7.6 solution (see Recipe 19).3and 1 mM EDTA solution (see Recipe 20).1% (w/v) NaHCO10 mM HCl solution (see Recipe 21).0.1 M glutathione reduced (GSH) solution (see Recipe 22).10 U/mL glutathione reductase (GSH-Rd) solution (see Recipe 23).t-butyl hydroperoxide solution (see Recipe 24).7 mM4 mM NADPH solution (see Recipe 25).Prepare a MIX solution containing:Note: The MIX solution must be prepared immediately before the assay. It is recommended to trigger the reaction in up to 16 duplicate samples. If you prepare more than 16 duplicate samples, the results will be different because of the long time between the reactions.240 μL of the 0.1 M GSH solution.1,200 μL of the 10 U/mL GSH-Rd solution.1,200 μL of the 4 mM NADPH.Keep this MIX solution on ice.Pipette 100 μL of the 50 mM TRIS/HCl 5 mM EDTA pH 7.6 solution in all wells of the microplate.Pipette 1.0 μL of the serum or 0.5 μL of the washed erythrocytes into all wells (in duplicates).Lightly tap the sides of the microplate several times to gently mix.Pipette 50 μL of the MIX solution into all wells.Lightly tap the sides of the microplate several times to gently mix.Pipette 20 μL of the 7 mM t-butyl hydroperoxide solution into all wells (to initiate a reaction).Note: It takes 15 s to pipette the 7 mM t-butyl hydroperoxide solution into the sample wells and trigger the reaction in up to 16 duplicate samples.Quickly homogenize the microplate.Measure the absorbance every one-minute interval for 5 min at 25 °C in a microplate at a wavelength of 340 nm using Gen5 software.Calculate GSH-Px concentrations .Catalase (CAT)2PO4·H2O solution (see Recipe 26).Prepare 0.1 M NaH2O2solution (see Recipe 27).Prepare 1 M HVerify hydrogen peroxide molarity:2PO4·H2O solution into a quartz cuvette (cuvette 1) as a blank in the reference position.Pipette 1,000 μL of the 0.1 M NaH2PO4·H2O solution into another quartz cuvette (cuvette 2) in the reading position.Pipette 990 μL of the 0.1 M NaHSelect AUTO ZERO in a UV1800 spectrophotometer for calibration.Wait until the absorbance reaches zero.Keep the cuvette 1 in the reference position.2O2into cuvette 2.Add 10 μL of 1 M HQuickly homogenize.Return cuvette 2 to the reading position.Measure the absorbance at a wavelength of 240 nm.Calculate the hydrogen peroxide real molarity .Wash both cuvettes.2PO4·H2O solution with 1.0 μL of washed erythrocytes into a quartz cuvette (cuvette 1) as a blank (in the reference position).Pipette 999 μL of the 0.1M NaH2PO4·H2O solution with 1.0 μL of the washed erythrocytes into another quartz cuvette (cuvette 2) in the reading position.Pipette 978 μL of the 0.1M NaHSelect AUTO ZERO in a UV1800 spectrophotometer.Wait until the absorbance reaches zero.2O2into the quartz cuvette 2 for a quick homogenization.Add 21 μL of 1 M HPerform the kinetics measurements for 2 min at 25 °C in a UV1800 spectrophotometer at 240 nm using UVProbe software.Calculate CAT concentrations .To ensure sample homogeneity, all measurements were adjusted according to the protein concentration estimated in each sample .AssaysTo determine the minimum sample volume and reagent concentrations required to reproduce each reference method in pregnant rats at term, decreasing amounts of reagents were tested until the results obtained were equal to the reference values of other articles established in the literature. All samples were analyzed in duplicates at 25 °C.Hemoglobin-1cm-1)  of hemoglobin per deciliter (dL), was measured at 546 nm  and calcm-1)  and the Hb =hemoglobin concentration in g/dLA =mean sample absorbance – blank absorbance80 =dilution factor (Drabkin’s solution volume/sample volume)16114.5 =hemoglobin monomer molar mass (complex cyanmethemoglobin–hemoglobin–cyanide monomer at 546 nm)10 =conversion from L to dL1000 =conversion from mg to gTotal protein concentration2, which is the value of the angular coefficient. The closer to 1 the value of R is, the better is the line described by the linear regression of the points aiming to obtain lines ≥ 0.99. Quantification requires knowing the dependence between the measured response and the concentration of the analyte. Linearity is obtained by internal or external standardization and formulated as a mathematical expression used to calculate the concentration of the analyte to be determined in the real sample. The equation of the line that relates the two variables is:Total protein absorbance values were determined by Bradford’s method (1976) for the normalization of oxidative stress biomarkers using Gen5 software. The total protein concentration is determined by the linear equation obtained from the standard curve. Standard curve is an increasing or decreasing succession of points obtained from the relationship between the concentration of the standard species and its signal intensity from the detection system. The most important part of the graph is the quadratic equation, from which the concentration can be calculated. The graph also provides the value of Ry=ax+by= measured response (absorbance)x= concentrationa= slope of the analytical curve = sensitivityb= intersection with the y axis when x = 0®, follow the steps depicted inTo calculate the linear equation from the standard curve in Microsoft Office Excely= measured response (absorbance)x= total protein concentration in μg/μL457.94= slope of the analytical curve = sensitivity127.53= intersection with the y axis when x = 00.9978= angular coefficient2value < 0.98, repeat the standard curve measurements.Note: If RTo calculate total protein concentration in serum samples, follow the example below:Considering an absorbance of 500 of the serum sample, the calculation is:y=457.94x+127.53500=457.94x+127.53x=(500-127.53)/457.94x= 0.81 μg/μL of total protein in the sample. To convert μg/μL to g/dL, divide the concentration value by 10.Note: If the absorbance value of the serum sample is higher than the absorbance of the 1.0 BSA solution concentration (standard curve), a dilution of 1:2 with purified water must be performed in the serum sample and the absorbance must be measured again. Then, the total protein concentration value (g/dL) must be multiplied by the dilution factor .Thiobarbituric acid reactive substances (TBARS)-1cm-1at 25 ºC  in washed erythrocyte or serum samples, respectively. The MDA-TBA complex extinction coefficient of 1.56 × 105 Mt 25 ºC  and 0.9 TBARS =Thiobarbituric acid reactive substance concentration in nM/mg Hb or total proteinA =mean sample absorbance – blank absorbance2 =sample dilution factor9=10conversion from M to nMHb =concentration obtained by Drabkin’s solution expressed as g/dLTotal protein (PTN) =as PTN concentration obtained by Bradford’s method is expressed in μg/μL, it is necessary to divide this by 10 to convert to g/dLFollow the steps below to obtain Hb or PTN mass (mg):To convert g/dL (Hb or PTN concentrations) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the used volume (0.5 mL) must be inserted by multiplication after the last conversion.Finally, the total conversion is: Hb or PTN concentration (g/dL x 10) = Hb or PTN concentration (mg/mL x 0.5 mL) = Hb or PTN mass (mg)Reduced thiol groups (-SH)-1/cm-1at 25°C) . The 5-thio-2-nitrobenzoic acid (TNB) extinction coefficient (14150 Mt 25°C)  and a 0.SH =concentration in mM/mg Hb or total proteinA =mean sample absorbance – blank absorbance3=10conversion from M to mMHb =concentration obtained by Drabkin’s solution expressed as g/dLPTN =as PTN concentration obtained by Bradford’s method is expressed as μg/μL, it is necessary to divide this by 10 to convert to g/dLFollow the steps below to obtain Hb or PTN mass (mg):To convert g/dL (Hb or PTN concentrations) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the sample volume used in the assay (mL) must be inserted by multiplication after the last conversionFinally, the total conversion is: Hb or PTN concentration (g/dL x 10) = Hb or PTN concentration (mg/mL x sample volume used in the assay) = Hb or PTN mass (mg)2O2)Hydrogen peroxide . The extinction coefficient of phenol red oxidation by H2O2was calculated from an adjusted standard curve in our laboratory at 37 °C (0.0296 µM-1cm-1) and 0.33 cm pathlength.Absorbance was measured at a wavelength of 620 nm using Gen5 software. The results were expressed as µM of H2O2=Hconcentration in µM/mg Hb or total proteinA =mean sample absorbance – blank absorbanceHb =concentration obtained by Drabkin’s solution expressed as g/dLPTN =as PTN concentration obtained by Bradford’s method is expressed as μg/μL, it is necessary to divide this by 10 to convert to g/dLFollow the steps below to obtain Hb or PTN mass (mg):To convert g/dL (Hb or PTN concentrations) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the sample volume used in the assay (mL) must be inserted by multiplication after the last conversion.Finally, the total conversion is: Hb or PTN concentration (g/dL x 10) = Hb or PTN concentration (mg/mL x sample volume used in the assay) = Hb or PTN mass (mg)Superoxide dismutase (SOD)Absorbance was immediately measured during the time intervals of 0 (T0) and 10 (T10) min at 25 °C in a microplate at 420 nm using Gen5 software. For the calculations, the absorbances of the blanks and samples at times T0 and T10 were used. The activity of SOD is calculated indirectly by the percentage of inhibition of pyrogallol by the sample. Therefore, the calculations were made following these steps:Calculation of the percentage of inhibition of the pyrogallol autoxidation by the SOD samples:x% =percentage of pyrogallol autoxidation by SOD sampleThus, the inhibition of pyrogallol autoxidation is equal to 100 – x%.Calculation of SOD activity considering that 1 IU is the volume capable of inhibiting 50% of the pyrogallol oxidation :y =sample volume (mL) that inhibits 50% of pyrogallol autoxidation50% =percentage inhibition of pyrogallol that equals 1 U of SOD0.003 =volume (mL) of the sample in the assay100-x% =percentage of pyrogallol inhibition by the sampleSOD enzymatic activity:[SOD] =enzymatic activity in IU/mg HbY =sample volume (mL) that inhibits 50% of pyrogallol autoxidation100 =conversion factor from dL to mL referring to the value of Hb (1 dL = 100 mL)Hb= concentration obtained by Drabkin’s solution expressed as g/dLPTN= as PTN concentration obtained by Bradford’s method is expressed as μg/μL, it is necessary to divide this by 10 to convert to g/dLFollow the steps below to obtain Hb or PTN mass (mg):To convert g/dL (Hb or PTN concentrations) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the sample volume used in the assay (mL) must be inserted by multiplication after the last conversion.Finally, the total conversion is: Hb or PTN concentration (g/dL x 10) = Hb or PTN concentration (mg/mL x sample volume used in the assay) = Hb or PTN mass (mg)Glutathione peroxidase (GSH-Px)340) per min was calculated using Gen5 software and the results were expressed as mM of GSH-Px/min/mg Hb or total protein. The NADPH extinction coefficient (6.220 M-1cm-1at 25 °C) (Absorbance change (ΔA 25 °C)  and 0.5 GSH-Px =activity in mM/min/mg Hb or total protein340/min=ΔAmean delta absorbance3=10conversion from M to mMHb =concentration obtained by Drabkin’s solution expressed as g/dLPTN =as PTN concentration obtained by Bradford’s method is expressed as μg/μL, it is necessary to divide this by 10 to convert to g/dLFollow the steps below to obtain Hb or PTN mass (mg):To convert g/dL (Hb or PTN concentrations) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the sample volume used in the assay (mL) must be inserted by multiplication after the last conversion.Finally, the total conversion is: Hb or PTN concentration (g/dL x 10) = Hb or PTN concentration (mg/mL x sample volume used in the assay) = Hb or PTN mass (mg)Catalase (CAT)Hydrogen peroxide molarity:2O2absorbance, perform the equation to know the solution molarity. For example, the H2O2absorbance was 0.416 and the molar absorptivity of H2O2at 25 °C is 43.6. Therefore:After verifying the HC=A/ε → C=0.416/43.6 → C = 0.0095 M = 9.5 mMC =concentration (mMol/L)A =AbsorbanceΕ =Molar absorptivity (M-1)Thus, although the solution concentration in the cuvette should be 10 mM , it is instead 9.5 mM.To correct the solution concentration, perform the following equation:1× V1= C2× V2C1=CReal concentration1=VVolume to be pipetted2=CConcentration of the prepared H2O2solution2=VVolume to be pipetted in the cuvette 21V= 10 mM × 10 µL9.5 mM ×1V= 10.5 µL – volume that must be pipetted in the cuvette to obtain a final solution at 10 mM.2O2must be 20 mM. Therefore, the volume of the solution that will be pipetted in the cuvette will be 21 µL (V1× 2).In the assay, the final concentration of HCAT activity2O2extinction coefficient (43.6 M-1cm-1at 25 °C) . The H 25 °C)  and a 1.CAT =activity in mM/min/mg of Hb = ∆A240/minmean delta absorbance3=10conversion from mol to mmolHb =concentration obtained by Drabkin’s solution expressed as g/dLFollow the steps below to obtain Hb mass (mg):To convert g/dL (Hb concentration) to mg/mL, it is necessary to multiply by 10 (g to mg and dL to mL). In addition, the sample volume used in the assay (mL) must be inserted by multiplication after the last conversion.Finally, the total conversion is: Hb concentration (g/dL x 10) = Hb concentration (mg/mL x sample volume used in the assay) = Hb mass (mg)All data was obtained and analyzed using the software Statistica (StatSoft). A p-value ≤ 0.05 was considered statistically significant.Resultst-test)  .TBARS and SH measurements were performed in washed erythrocytes samples diluted into purified water solution because the stabilizing solution interfered with the results of the assays . The int2O2) in washed erythrocytes according to the diluent used (purified water or stabilizing solution). Thus, hemoglobin concentration must be twice determined: in purified water and in stabilizing solution.Hemoglobin (Hb) concentration was determined and used to normalize the oxidative stress biomarkers  in all formulas of oxidative stress biomarkers.In TBARS assay, all the samples must be thawed at 25 °C. After TBA solution preparation, the pH should be below 2.0. If the pH is above 2.0, it is possible that the purified water conductivity is impaired.2O2), superoxide dismutase (SOD), GSH-Px, and CAT assays, all the samples must defrost on ice and be kept cooled to not allow degradation to occur.In reduced thiol groups (-SH), hydrogen peroxide (H2O2) is a volatile substance; thus, it is necessary to perform the equation for H2O2real molarity when executing the assay.Hydrogen peroxide , but the sample and reagent volumes were adjusted to the microplate volume .Stabilizing solution (2.7 mM EDTA and 0.7 mM 2-mercaptoethanol)EDTA 10 mL2-mercaptoethanol 0.5 μLNote: Solution stable for up to six months at 2–8 °C.Drabkin’s solutionPotassium ferricyanide 200 mgPotassium cyanide 50 mg31 gNaHCOPurified water 1,000 mLNote: Drabkin’s solution is light sensible. Store at 4 °C in an Amber bottle to protect from light. This solution is stable for up to six months at 2–8°C.1 μg/μL stock solutionDilute 1 g of BSA in 1 mL of purified water.Note: Solution stable for up to six months at 2–8 °C.3% (w/v) 5-sulfosalicylic acid hydrate (for 100 samples)Sulfosalicylic acid 3 gUltrapure water 100 mLDilute sulfosalicylic acid in a volumetric flask adding purified water.Seal it with laboratory film.Keep at 25 °C.Note: Solution must be freshly prepared. After use, discard remaining solution.Thiobarbituric acid solution (TBA) 0.67% (for 200 samples)Dilute 0.67 g of TBA in 95 mL of purified water in an Amber bottle.Keep the bottle partially sealed and heat the solution to 80 °C for 30 min in a water bath.Wait until the solution reaches 25 °C.Adjust pH to 2.0 by adding 1 M NaOH (see Recipe 16).Add the solution to a beaker and complete to 100 mL using purified water.Note: Solution must be freshly prepared at the assay day. This solution is light sensible. Store in Amber bottle. After use, discard remaining solution.1 M HClDilute 8.3 mL of HCl in 100 mL of purified water.Note: Solution stable for undetermined time at 2–8 °C.0.1 M TRIS/HCl 0.5 mM EDTA pH 8.0 (for 250 duplicate samples)TMand 0.0146 g of EDTA in 90 mL of purified water.Dilute 1.2114 g of TRIS-UltrapureAdjust pH to 8.0 by adding 1 M HCl (see Recipe 6).Transfer the solution to a beaker and complete to 100 mL using purified water.Note: Solution stable for undetermined time at 2–8 °C.10 mM DTNB (for 250 duplicate washed erythrocytes samples and 100 duplicate serum samples)Dilute 0.0079 g of DTNB in 2 mL of ethyl alcohol.Aliquot the total volume in two microtubes (1 mL each)Seal and store at -20 °C.Note: Solution must be freshly prepared and stored in a sealed tube at -20°C. After use, discard remaining solution.2HPO4, 0.1 M Na2HPO4Solution 1: 1.71 M NaCl, 34 mM KCl, 14 mM K2HPO4, and 1.15 g of Na2HPO4in a volumetric flask.Mix 8 g of NaCl, 0.2 g of KCl, 0.2 g of KAdd 80 mL of purified water and seal with laboratory film.Keep at 25 °C until buffer solution B preparation (Recipe 15).Store at 2–8 °C after buffer solution B preparation.Note: Solution stable for undetermined time.2Solution 2: 90 mM CaCl2in 30 mL of purified water.Dilute 0.3 g of CaClHomogenize and seal the solution with laboratory film.Keep at 25 °C until buffer solution B preparation (Recipe 15).Store at 2–8 °C after buffer solution B preparation.Note: Solution stable for undetermined time.2Solution 3: 0.11 M MgCl2in 30 mL of purified water.Dilute 0.3 g of MgClHomogenize and seal the solution with laboratory film.Keep at 25 °C until buffer solution B preparation (Recipe 15).Store at 2–8 °C after buffer solution B preparation.Note: Solution stable for undetermined time.Solution 4: 0.1% phenol redDilute 0.3 g of phenol red in 30 mL of purified water in a tube.Homogenize the solution at 54 °C in water bath until complete dissolution.Remove the solution from the water bath.Seal the tube with laboratory film.Keep at 25 °C until buffer solution B preparation (Recipe 15).Store at 25 °C after buffer solution B preparation.Note: Solution stable for undetermined time.Phosphate buffer solution A2HPO4to 150 mL of purified water.Add 2.61 g of KHomogenize and seal the solution with laboratory film.Keep the solution at 25 °C.2PO4to 150 mL of purified water.Add 2.06 g of KHHomogenize and seal the solution with laboratory film.Keep the solution at 25 °C.2HPO4solution in a volumetric flask and add KH2PO4solution until reaching pH 7.0.Add 100 mL of KStore at 2–8 °C.Note: Solution stable for undetermined time.Peroxidase solution (HRP)Dissolve 5 mg of HRP in 1 mL of phosphate buffer solution A.Note: Solution must be freshly prepared. After use, discard remaining solution.Buffer solution B10.56 mL of purified water0.960 mL of Solution 10.120 mL of Solution 20.120 mL of Solution 30.120 mL of Solution 40.120 mL of peroxidase solution (HRP)Note: Solution must be freshly prepared. After use, discard remaining solution.1 M NaOHDilute 0.040 g of NaOH in 0.10 mL of purified water.Note: Solution stable for undetermined time at 2–8 °C.1 M TRIS/HCl 5 mM EDTA pH 8.0 (for 150 duplicate samples)TMand 0.0731 g of EDTA in 45 mL of purified water.Dilute 6.0570 g of TRIS-UltrapureAdjust pH to 8.0 by adding 1 M HCl (see Recipe 6).Transfer the solution to a beaker and complete to 50 mL using purified water.Store in a volumetric flask.Note: Solution stable for undetermined time at 2–8 °C.10 mM pyrogallol solution Dilute 0.0063 g of pyrogallic acid in 5 mL of 10 mM HCl solution (see Recipe 21).Note: The solution must be freshly prepared. Minimum volume of 5 mL due to the pyrogallol quantity to be weighted. After use, discard remaining solution.50 mM TRIS/HCl 5 mM EDTA pH 7.6 TMand 0.1461 g of EDTA in 90 mL of purified water.Dilute 0.6057 g of TRIS-UltrapureAdjust pH to 7.6 by adding 1 M HCl (see Recipe 6).Transfer the solution to a beaker and complete the solution to 100 mL using purified water.Store in a volumetric flask.Note: Solution stable for undetermined time at 2–8 °C.3and 1 mM EDTA1% NaHCO3and 0.0292 g of EDTA to 100 mL of purified water in a volumetric flask.Dilute 1 g of NaHCONote: Solution stable for undetermined time at 2–8 °C.10 mM HCl solutionDilute 50 μL of HCl in 60 mL of purified water.Note: Solution stable for undetermined time at 2–8 °C.0.1 M glutathione reduced (GSH) Dilute 0.0184 g of GSH in 600 μL of 10 mM HCl solution (see Recipe 21).Note: The solution must be freshly prepared and kept on ice. After use, discard remaining solution.10 U/mL glutathione reductase (GSH-Rd) Add 60 μL of GSH-Rd to 2.940 mL of 0.1M TRIS/HCl 0.5 mM EDTA pH 8.0 (see Recipe 7).Note: The solution must be freshly prepared and kept on ice. After use, discard remaining solution.t-butyl hydroxiperoxide 7 mMt-butyl hydroperoxide in 5 mL of purified water.Dilute 5 μL ofNote: The solution must be freshly prepared. After use, discard remaining solution.4 mM NADPH 31 mM EDTA.Dilute 0.010 g of NADPH in 3 mL of 1% NaHCONote: Solution must be freshly prepared and kept on ice during the assay. Solution stable for up to 4 h.2PO4·H2O 0.1 M NaH2PO4·H2O in 190 mL of purified water.Dilute 2.7598 g of NaHAdjust pH to 7.0 by adding 1 M NaOH (see Recipe 16).Transfer the solution to a beaker and complete the solution to 200 mL using purified water.Store in a volumetric flask.Note: Solution stable for undetermined time at 2–8 °C.2O2)1 M hydrogen peroxide (HAdd 970 μL of hydrogen peroxide to 9.03 mL of purified water.Note: Solution must be freshly prepared. After use, discard remaining solution.


The formation of Al2O3 NPs and MAl2O4  nanocomposite was confirmed by using several characteristics such as XRD, UV–Vis, FTIR, FE-SEM, and EDX spectra. The XRD shows that the CoAl2O4 has a smaller crystallite size (12.37 nm) than CuAl2O4, NiAl2O4, and ZnAl2O4. The formation of a single-phase spinel structure of the calcined samples at 1100 °C was confirmed by FTIR. Our studies showed that the pure Al2O3 NPs have a lower energy gap (1.37 eV) than synthesized MAl2O4 under UV–Vis irradiation. Due to the well separation between the light-generated electrons and the formed holes, the cell containing ZnAl2O4 nanocomposite with CdS QDs has the highest efficiency of 8.22% and the current density of 22.86 mA cm−2, while the cell based on NiAl2O4 as a photoelectrode, six cycles of CdS/ZnS QDs, and P-rGO as a counter electrode achieved the best (PCE) power conversion efficiency of 15.14% and the current density of 28.22 mA cm−2. Electrochemical impedance spectroscopy shows that ZnAl2O4 and NiAl2O4 nanocomposites have the highest life times of the photogenerated electrons (τn) of 11*10−2 and 96*10−3 ms, respectively, and the lowest diffusion rates (Keff) of 9.09 and 10.42 ms−1, respectively.The aim of this study is to describe the performance of the aluminum oxide nanoparticle and metal aluminate spinel nanoparticle as photo-anodes in quantum dot photovoltaic. By using a sol–gel auto combustion method, Al They have been widely used as pigments, sensors, photocatalysts, magnetic, refractory, optical, and materials electrodes, involving lubricant additives15. It has been known that the synthesis method can influence the crystallinity, purity, surface area, morphology, and particle size of nanosynthesized materials MAl2O4, which has a respectable impact on their catalytic and optical properties18. MAl2O4 can be used with a variety of techniques including solvothermal method, co-precipitation synthesis, sol–gel method, solid-state reactions, hydrothermal method, microwave-assisted, polymeric precursor synthesis, and hydrothermal method30. Either method requires specialized accommodation and has a respectable cost, while further disadvantages include the shortage of homogeneity and low surface area of product33. The generation of the homogeneous high-purity nanoparticles has been demonstrated by the sol–gel auto combustion technique with swift heating and short reaction time34. The sol–gel auto combustion synthesis method combines chemical sol–gel and combustion processes, representing a swift, attainable technique with low energy costs, and is perfect for the synthesis of materials based on metal oxides. Several organic compounds can be utilized as the fuel, but these have been imitatively fixed to citric acid, urea, glycine, and tartaric acid12. Graphene is an intriguing material with a novel two-dimensional skeleton composed of a single monomolecular layer of sp2-hybridized carbon atoms4. Graphene has excellent properties in many areas of technology and science due to its unique properties6, including superior electronic8, mechanical, and thermodynamic properties10. Graphene has a broad domain of applications such as field-effect transistors (FET), transparent conductive films, energy storage devices, water purification, and sensors due to its elegant physical and chemical properties32. P-rGO is a carbon material that shows optical, chemical, and electrical characteristics similar to those of graphene because it is based on its framework35. In 1958, Hummers and Offman have well-educated a method for the synthesis of P-rGO41. This method uses H2SO4 to peel graphite with NaNO3 and KMnO4 as the oxidizing agents for graphite. The method of Hummer has some features compared to that of Brodie and Staudenmaier. Firstly, KMnO4 is a strong oxidant that aids in hastening the reaction so that the synthesis can be finished in a few hours. Secondly, the chlorate is not available, removing the probability of a ClO2 eruption. Thirdly, the exchange of fumigation with NaNO3 eliminates the acid haze generated by HNO315. To the best of our knowledge, there have been no studies comparing the effects of NiAl2O4, CuAl2O4, and ZnAl2O4 spinel aluminates, which were produced by the same synthetic method, on the performance of QDSSCs. Solar cells sensitized by quantum dots (QDSSCs) have attracted massive awareness in recent years, owing to their easy-fabrication procedure, low-cost, adjustable bandgap, and high theoretically mentioned power conversion efficiency (PCE) of up to 44%42. The photoelectrochemical technicality of QDSSCs has the same behavior as dye-sensitized solar cells (DSSCs), in which the solar cell is sensitized by the QDs instead of the dye molecules as a light absorber layer in QDSSCs43. A QDSSC is typically composed of a quantum dot sensitised photoanode film, an electrolyte containing a redox couple , and a counter electrode (CE) 44. In spite of these qualities, however, the performance of the photovoltaic cells of most QDSSCs is much lower than that acquired from DSSCs. To date, much of the research work has concentrated on enhancing all the elements in QDSSCs. Since one of the essential reasons for low efficiency is the counter electrode interfaces, particular research efforts have settled on the expansion of congruous CEs for achieving simultaneous high efficiency and stable cells.Alpha-alumina and metal aluminate spinel with the formula M-Al2 nanocomposite has a large absorption band of visible light. The graphene-TiO2 nanocomposite photoanode has been deposited onto FTO using the doctor blade method as a photoanode to obtain a negative capacitance. Photovoltaic cells (PV) currently have a low efficiency in converting light into electrical power. Despite the superposition of several PV cells that convert more than 41% of incident light power to electricity and several semiconductor materials having an efficiency lower than 21% as a result of the great use of energy in the industry, these PVs can only convert one third of the light power received. This efficiency is limited by some physical properties of the materials, such as the Joule losses. The challenge of the scientific community is to prevent the leakage of electrons to energy. Most of the PV cells are prepared from semiconductor materials because of their very special properties. It can absorb photons of a well-defined wavelength to generate free electrons, which then gives an electric current. Several studies have shown that the GO can generate many pairs of electron-holes by absorbing one photon and relaxing the primary pair of electron-holes excited by illumination. Instead of losing excess excitation energy by the joule effect or phonon form, graphene is a suitable material to create hot charge carriers by transferring the excess energy to other carriers. The power conversion efficiency (η) and the short current Isc increased with the graphene amount due to the intrinsic properties of this material. The efficiency of the process is correlated to the energy lost by the joule effect or phonon form. The efficiency can be increased if the excited charge carriers use their excess energy to produce electron–hole pairs through an interaction carrier-carrier or a diffusion process, instead of losing energy in the form of heat and phonons. In particular, the best performing solar cells employing rGO exhibited power conversion efficiency (PCE) of up to 18.13%, while the control device without rGO delivered a maximum efficiency of 17.26%. An effective and low-cost approach has been proposed for the fabrication of reduced graphene oxide (rGO) solar cells. The hybrid composite of rGO has been synthesized by using a low-cost chemical method and deposited on a glass substrate by a doctor-blading process for application in heterojunction solar cells. The morphology, crystal phase, surface functional groups, and optoelectronic properties of the heterostructure have been explored. The J-V characteristics of the fabricated heterojunction cell show a power conversion efficiency of 4.35%  = 51.60) due to the enhanced conductivity and charge transfer efficiency of the hybrid semiconductor. This simple approach assures the low-cost mass production of heterojunction solar cells49. Kafle et al. reported that the best performing solar cells employing rGO exhibited power conversion efficiency (PCE) of up to 18.13% in a perovskite cell composed of rGO as a counter electrode, while the control device without rGO delivered a maximum efficiency of 17.26%46. Kadhim et al., used the cell composed of TiO2 as a photoanode, CdS QDs, polysulphide electrolyte, and rGO as a counter electrode to give a PCE of 4.35%47. Mnasri et al., used the quantum dot sensitized solar cell composed of GO-TiO2 nanocomposites as a photoanode, five cycles of CdS QDs, polysulphide electrolyte and Pt as a counter electrode to give a PCE of 0.3%48.In the published literature, graphene-TiO2O4, NiAl2O4, CuAl2O4, and ZnAl2O4 spinel nanostructures by a sol–gel auto combustion technique as the working electrode in hybrid structures of ZnS/CdS quantum dots-based solar cells. The morphology of the surface and optical behavior of the aluminate spinels ) have been inspected by XRD, FT-IR, SEM, and UV–visible spectroscopy (UV–Vis). The effect of hybrid CdS QDSSC/ZnS QDs was also studied. The QDSSCs, which contain ZnAl2O4 as a photoanode, six cycles of CdS QDs as a photosensitized material, polysulphide electrolyte, and rGO as a counter electrode, gave a PCE of 8.22%. The cell contains a NiAl2O4 as a photoanode, twelve cycles of ZnS and CdS QDs, polysulphide electrolyte, and rGO as a counter electrode, given a PCE of 15.14%.In this work, for the first time, we have grown first-step, electrocatalytically active, highly stable CoAlMaterials Aluminum Nitrate (Al2(NO3)3.9H2O , Zinc Acetate Zn(CH3COO)2 , Sodium Sulfide Na2S , Ethanol absolute , Nitric acid HNO3 (Adwic), sulfuric acid H2SO4, , Triton-X100 , Extra pure graphite powder , Sulfur Powder (Adwic), Potasium Chloride (Advent), FTO (Fluorine-dopped SnO2) conductive glass (Aldrich), Nickle Chloride (NiCl2.6H2O), Cobalt Nitrate (Co(NO3)2) and Cupper Chloride (CuCl2.2H2O) from Adwic, Potassium Permanganate , Hydrogen Peroxide .2O3 nanostructured, Al2 (NO3)3.9H2O was used as the precursor. Solution A contains a desired amount of aluminum nitrate nonahydrate and was dissolved in ethanol to prepare a 0.1 M of Al2 (NO3)2 ethanoic solution by stirring for about 2 h using a magnetic stirrer. Solution B contains a 30% ammonia solution. The gel of aluminum oxide was formed by adding the 30% ammonia solution dropwise to the aqueous aluminum nitrate solution under vigorous stirring for one h. This gel was left for 30 h at room temperature for maturation time and dried at 200 °C for 24 h. The resulting gel was annealed in a furnace for 4 h at 1100ºC11.The auto-combustion sol–gel method used for the synthesis of Al2O4 spinel nanostructures were prepared by the sol–gel auto combustion method. Stoichiometric amounts of Al2 (NO3)3.9H2O and metal nitrate were dissolved in absolute ethanol to obtain a 0.2 M solution. Then, a convenient amount of DEA was added as fuel to the solution. The mixture was heated to 200 °C under continuous stirring until all of the ethanol had evaporated and the process took place. The mixed solution mutated into a condensed gel, which self-ignited to produce a pearly yellow powder for ZnAl2O452, a dark powder for CoAl2O4, a brown powder for CuAl2O430 and a grey powder for NiAl2O423. Finally, the as-synthesized powders were annealed in the air at 1100 °C for 4 h16.The M-Al39. In a solution containing 50 ml of concentrated H2SO4 and 50 ml of concentrated HNO3, the extra pure graphite powder (2.0 g) was pre-oxidized by slowly adding it to the mixture and stirring at 80 °C for 4 h. The mixture was cooled down to room temperature and then washed by de-ionized water until the pH value was neutral , followed by drying at 40 °C overnight. The resultant pre-oxidized graphite was dispersed into concentrated H2SO4 in a cold reaction vessel, which was kept in an ice bath and stirred. 10 g of KMnO4 was slowly added to it. During the addition, the temperature was kept below 10 °C. The mixture was stirred at 35 °C for 2 h until the solution became gelled and turned a brownish gray in color. Then 250 ml of de-ionized water was added and the temperature was put forward to 100 °C for 15 min, followed by adding 700 ml of de-ionized water and 30 ml of H2O2 to the mixture, stirred for 1 h. The solid products were composed from the solution after 12 h and washed with 5% HCl until sulphate ions were no longer detectable with BaCl2. Then the solid products were re-dispersed in de-ionized water five times to eliminate any impurities. Finally, the resultant sediment was dried at 60 °C for 4 h in an oven to yield the partially reduced graphene oxide P-r (GO)39. Fig. Partially-reduced graphene oxide (P-rGO) was synthesized from the graphite powder according to the modified Hummers and Offman method2O3 nanoparticle and M-Al2O4 nanocomposite films with a thickness of 0.052 mm were prepared and used for solar cell testing. The samples of Al2O3NP and M-Al2O4 nanocomposites were deposited onto conductive Fluorine-tin-oxide (FTO) glass substrates by the doctor-blading method using a dispersion of 0.2 g of Al2O3 powder in ethanol solution and the same of 0.2 g of CoAl2O4, CuAl2O4, NiAl2O4 and ZnAl2O4. Before spreading the film, the surfaces of the FTO substrates were cleaned for 30–60 min using an ultrasonicator. The substrates were then dried in air. The FTO boundaries of each substrate were covered with scotch tape to control the area of the film. The powder was first strewn in Triton-X100 and ethanol, and then the pending powder was added as drops in the midst of the substrate and prevalence to form a thick film to produce the film. The film was dried in the air for 30 min and then calcined at 300 °C for 5 min53.Both Al3)2 aqueous solution for 2 min as a Cd+2 source, then rinsed in ethanol to remove surplus ions and dried on a hot plate at 60 °C for 1 min. Then, the film was dipped into a 0.2 M Na2S aqueous solution for another 2 min to allow S2 to react with the pre-adsorbed Cd2+ leading to the formation of CdS QDs. Swiping the film in methanol and drying at 60 °C for 1 min removed the loosely bound S2−ions. On the other hand, the hybrid structures of ZnS/CdS QDs are prepared by the same method (SILAR) by the growth of ZnS QDs. The semiconductor film was immersed in a 0.2 M Zn (CH3COO)2 aqueous solution for 2 min before being rinsed with ethanol to remove the excess ions and dried for 1 min on a hot plate at 60 °C. Then, the film was dipped into a 0.2 M Na2S aqueous solution for another 2 min to allow S2 to react with the pre-adsorbed Zn2+, leading to the formation of ZnS QDs. The loosely bound S2−ions were removed by swiping the film in methanol and drying it at 60 °C for 1 min. Second, a semiconductor and ZnS QDs film were dipped into a 2 M Cd (NO3)2 aqueous solution as a Cd+2 source. Then, the film was dipped into a 0.2 M Na2S aqueous solution for another 2 min to allow S2−to react with the pre-adsorbed Cd2+, leading to the formation of hybrid structures of CdS QDs and ZnS QDs. The loosely bound S2−ions were removed by rinsing the film in methanol and then the film was dried at 60 °C for 1 min. To increase the crystallinity of QDs, the films with different quantum dot depositions were calcined at 300 °C for 5 min45.The photo electrode was sensitized with the CdS QDs, which act as light absorbers by the Successive Ionic Layer Adsorption and Reaction (SILAR) method. To grow CdS QDs, the semiconductor film was immersed in a 0.2 M Cadmium nitrate . All the J-V curves of the solar cells were obtained in the dark and under illumination. The J-V characteristics as a function of incident light intensity were used to obtain the open-circuit voltage (Voc), short-circuit current density (Jsc), the maximum voltage point (Vmax), and the maximum current density point (Jmax).The performance of the solar cell was measured using strained solar illumination with a 10.4 mW/cmDifferent techniques were used to examine the surface of prepared materials.The X-ray diffraction patterns were reported using a Pan Analytical Model X 'Pert Pro, which was fitted with CuKα radiation (α = 0.1542 nm), a Ni-filter, and a general area detector. A 40 kV accelerating voltage and a 40 mA emission current were utilized. The diffractograms were measured in the 2θ range from 0.5 to 70 °.The Fourier transform infrared spectroscopy (FT-IR) of the prepared samples was measured using the KBr technique adopted by the Nicolet Is-10 FT-IR spectrophotometer (Thermo Fisher Scientific). The KBr technique was conducted roughly in a quantitative manner for all samples, since the sample weight and that of KBr were both held constant.A Field Emission Scanning Electron Microscope (FE-SEM) is used to examine the material's composition, particle size, and shape. For the samples prepared, a 30 kV acceleration voltage operating on a JSM-7500F electron microscope was reported.Optical absorption spectra of the sample were analyzed using Ultraviolet–Visible absorption spectroscopy .2O3 NPs and M-Al2O4 NCs were executed as thin films on conducting glass (FTO) of dimension (6.25 cm2 surface area and a 0.052 mm thickness) for every sample to serve as electrodes during the measurements by a standard two-probe technique using an impedance analyzer .Dielectric studies of Al2O3 NPs, ZnAl2O4, NiAl2O4, CoAl2O4, and CuAl2O4 spinel is shown in Fig. 2O3 NPs in a pure single hexagonal phase according to JCPDS Card No. 42-1468. Figure 2O4, which exhibited diffraction peaks at 25.73°, 38.006°, 43.87°, 49.593°, 51.626°, 66.306°, 67.59°, 72.146°, 81.966° and 86.046° corresponding to the crystal planes, (200), (220), (311), (400), (410), (422), (511), (440), (531) and (620), which confirmed the formation of a pure single phase of NiAl2O4, in agreement with JCPDS No. 10-033955. The strong and sharp diffraction peaks indicate that NiAl2O4 has a long-range ordered structure. Figure 2O4 cubic spinel (JCPDS No. 33-0448) calcined at 1100 °C40. The Figure also depicts the characteristic CuAl2O4 peaks at 23.66°, 25.406°, 31.305°, 35.103°, 37.73°, 43.43°, 48.65°, 52.50°, 57.62°, 66.46°, 68.138°, 73.29 and 76.83° corresponding to the crystal planes, (511), (110), (111), (220), (311), (422), (311), (400), (422), (511), (440), (620) and (533). The formed peaks are congruent to the formation of ZnAl2O4 (JCPDS No. 05-0669)55 in a single phase structure with characteristic peaks at 35.599°, 38.28°, 40.079°, 51.37°, 60.36°, 66.76°, 69.12°, 71.78°, 72.86° and 80.89° corresponding to the crystal planes, (220), (311), (222), (400), (331), (422), (511), (440), (620) and (533). CoAl2O4, the characteristic peaks appear at 19.034°, 25.46°, 31.28°, 36.85°, 44.84°, 55.67°, 59.31°, 65.71°, 74.15°and 77.33° corresponding to the crystal planes, (220), (311), (222), (400), (331), (422), (511), (440), (620) and (533) (JCPDS No. 44-0160). In addition, the average crystallite size (D) of spinel aluminate nanostructures can be predestined from the full width at half maximum of the strongest diffraction peak by applying the Debye–Scherrer equation, Eq. (The X-ray diffraction pattern (XRD) for Al2O3 NPs, ZnAl2O4, NiAl2O4, CoAl2O4, and CuAl2O4 spinel oxides calcined at 1100 ℃ as a set of transmission peaks in the range of 400–4000 cm−1. The functional groups of aluminum oxide NPs and metal aluminate spinel structures were inspected by FT-IR analysis. The stretching mode of Al–O and the Al–O–Al bending mode in the spectrum of Al2O3 NPs appear at the peaks of 594 and 1598 cm−1, respectively. A band at 3451 cm−1 representing the –OH species was detected for the Al–OH bond formation. A small peak at 1634 cm−1 was attributed to the H–O–H bending vibration characteristic peak. This confirms the existence of water molecules absorbed on the surface of the Al2O3 NPs. Further, a sturdy absorption band at 1398 cm−1 occurs due to the distortion caused by the C–CH3, C–C, and C=O bonds. All the samples implicate concerted absorption bands around 3462, 2922, 2850, 2337, 1633, 1385, 1200, and 1118 cm−1 as shown in Fig. −1 are comforted to absorb water molecules' OH-stretching and bending vibrational modes, respectively27. Different IR studies of Al2O3 NPs and spinel metal aluminate nanocomposites observed the water absorption peaks and declared that the high surface area of these materials could result in swift adsorption of water from the atmosphere during pellet squeeze and IR mensuration56. C–H stretching vibration is represented by small peaks at around 2922 and 2850 cm−11. The absorption band at 2337 cm−1 is supposedly connected to the entity of CO2 on the surface powder, while a small peak at 1385 cm−1 can be attributed to the grout nitrogen groups output from the combustion reaction56. The peaks in the 1200 cm−1 region can be explained as vibrations of the C–C bond, and bands at around 1118 cm−1 can be related foremost to oxygen groups with a single C–O bond57. In the FT-IR spectrum of NiAl2O4 nanocomposites calcined at 1100 °C, the characteristic bands of NiAl2O4 appear around 3390, 1629, 725, and 478 cm−1, which would affirm the consistence of the NiAl2O4 aluminate spinel structure in good agreement with the XRD results58. These bands can be conformed to the uniform stretching, bending, and asymmetric stretching modes of Ni–O, Al–O, and Ni–O–Al bonds at octahedral and tetrahedral sites in M-Al2O4 spinel58. The FT-IR spectrum of CuAl2O4 nanocomposites calcined at 1100 ℃ confirms the formation of CuAl2O4 nanocomposites through the strongest four characteristic bands of CuAl2O4, which are 453, 501, 591, and 640 cm−158. The FT-IR spectrum of ZnAl2O4 nanocomposites calcined at 1100 °C shows sharp bands between 1634 and 3435 cm−1 (C–O and C–C vibration) and 1161 cm−1 (NO3− vibration), including the three OH vibration bands mentioned above. The bands lying at 1111, 668, and 490 cm−1 are comforted by the vibrations of Zn–O, Al–O, and Zn–O–Al bonds in ZnO4 tetrahedral and AlO6 octahedral groups in the spinel structure58. The spectrum of CoAl2O4 nanocomposites calcined at 1100 °C is also shown by FT-IR, with sharp bands between 2893 and 3335 cm−1 (C–O and C–C vibration) and at 1523 cm−1 (NO3− vibration). The bands lying at 1113, 670, and 565 cm−1 are confined to the vibrations of Co–O, Al–O, and Co–O–Al bonds, which confirmed the formation of CoAl2O4 nanocomposites.Figure 2O3 NPs, ZnAl2O4, CuAl2O4, NiAl2O4 and CoAl2O4 nanocomposites calcined at 1100 °C. FE-SEM images of the samples show uniform, spherical, and homogeneous nanoparticles of Al and Zn/Al with mean particle sizes of 29.77 and 44.66 nm, respectively. Figure 2O4 and CoAl2O4 nanocomposites is in the ranges of 18.98–50.07 nm and 40.08–74.53 nm, respectively. In addition, the release of fragile gases such as CO2, N2, O2, and H2O during the combustion process should be one of the prime factors that create different pore structures in the ZnAl2O4, CuAl2O4, NiAl2O4, and CoAl2O4 nanocomposites. This interconnected pore structure is remarkable for catalytic applications. The amount of distribution is dependent on the homogeneity of the size of the particles. The EDX spectrum of Al2O3 NPs, ZnAl2O4, CuAl2O4, NiAl2O4, and CoAl2O4 obtained from FE-SEM is shown in Fig. 2O4, CuAl2O4, CoAl2O4, and NiAl2O4 nanocomposites was found to be 24.12/13.82 = 1.75, 25.17/13.28 = 1.89, 24.66/13.71 = 1.79, 8.31/3.98 = 2.09, respectively, for the best determination with the K-line series. This is in agreement with the results obtained from XRD. Fig. Figure 2O3 NPs, ZnAl2O4, CoAl2O4, NiAl2O4, and CuAl2O4 nanocomposites under UV–Vis irradiation decreases as the wavelength increases. In Fig. 2O3 NPs is 1.59% higher than the optical absorbance of ZnAl2O4, CoAl2O4, NiAl2O4 , and CuAl2O4 nanocomposites, which are , respectively, because the adding of metals like Zn, Co, Ni, and Cu has a high effect on the absorbance and causes a lower shift. Figure 2O3, ZnS/ZnAl2O4, ZnS/CoAl2O4, ZnS/NiAl2O4 and ZnS/CuAl2O4. The ZnS/CoAl2O4 sample has a higher absorbance (2.78%) than ZnS/NiAl2O4, ZnS/CuAl2O4, ZnS/ZnAl2O4, and ZnS/Al2O3, which are  because ZnS QDs are more sensitive to light and rapidly make a shift in absorption edge. Figure 2O3, CdS/ZnAl2O4, CdS/CoAl2O4, CdS/NiAl2O4, and CdS/CuAl2O4, where the CdS/CuAl2O4 sample has a higher absorbance (2.9%) than CdS/Al2O3, CdS/ZnAl2O4, CdS/CoAl2O4, and CdS/NiAl2O4, which are . Figure 2O3, CdS/ZnS/ZnAl2O4, CdS/ZnS/CoAl2O4, CdS/ZnS/NiAl2O4, and CdS/ZnS/CuAl2O4, where the CdS/ZnS/CuAl2O4 sample has a higher absorbance (2.93%) than CdS/ZnS/Al2O3, CdS/ZnS/ZnAl2O4, CdS/ZnS/NiAl2O4 and CdS/ZnS/CoAl2O4, which are . In addition, comparison between the prepared materials and the same method was done by using the same thickness of these materials.The absorbance of pure Al2O3 NPs, ZnAl2O4, CoAl2O4, NiAl2O4, and CuAl2O4 nanocomposites for direct allowed transition is shown in Fig. 2O3 NPs, ZnAl2O4, CoAl2O4, NiAl2O4 and CuAl2O4 nanocomposites are 1.37 eV, 1.5 eV, 1.83 eV, 1.59 eV and 2.07 eV, respectively. The Tauc plots of ZnS/Al2O3 and ZnS/MAl2O4 in direct allowed transition are shown in Fig. 2O3, ZnS/ZnAl2O4, ZnS/CoAl2O4, ZnS/NiAl2O4, and ZnS/CuAl2O4 are 1.61 eV, 1.67 eV, and 1.13. The Tauc plots of CdS/Al2O3 and CdS/MAl2O4 in direct allowed transition are shown in Fig. 2O3, CdS/ZnAl2O4, CdS/CoAl2O4, CdS/NiAl2O4, and CdS/CuAl2O4 are 1.58 eV, 1.38 eV, and 1.33. The Tauc plots of CdS/ZnS/Al2O3 and CdS/ZnS/MAl2O4 in direct allowed transition are shown in Fig. 2O3, CdS/ZnS/ZnAl2O4, CdS/ZnS/CoAl2O4, CdS/ZnS/NiAl2O4, CdS/ZnS/CuAl2O4 are 1.57 eV, 1.42 eV, 1.64 eV, 1.66 eV, and 1.38 eV, respectively. The absorption coefficient of different materials is calculated by this equation, where (A) is the absorbance of material and (t) is the thickness of it. The thickness of thin films is constant for all materials. The presence of metals like Co, Ni, Zn, and Cu has an effect on the energy gap of materials by decreasing it and contributing to narrowing the band gap of the main semiconductor. All the energy gaps for all materials are listed in the below Table To know the optical bandgap, we have made an itemized calculation of the band gap using the Tauc formula; (αhν) = A (hυ − Eg) n, where, α is the absorption coefficient, hυ is the photon energy, Eg is the energy gap, and n is the quality of the transitions. The term n is taken as ½ for direct transition and 2 for indirect transition. The optical bandgap energy is investigated by plotting (αhυ) 1/n versus photon energy and drawing the tangent to the curve that intersects with the energy axis at α = 0. The Tauc plot of pure Al2O3 NPs, ZnAl2O4, CoAl2O4, NiAl2O4, and CuAl2O4 nanocomposites with CdS QDs on FTO conductive glass through the dark field was determined by electrochemical impedance spectroscopy (EIS) and can be illustrated by the kinetic operation of charge relocated in QDSSCs. Figure s. The data for samples gives the values of charge transport resistance, Rct for the first semicircle arc and charge recombination resistance (Rrec) for the second semicircle arc at high frequency and low frequency, respectively. The good adhesion of the bond between ZnAl2O4 nanostructures and the FTO conductive glass substrate pointed to the lower Rct value, which supports more electrons from the external circuit. Charge transfer resistance is of primary significance because it eases the carry of electrons during the catalytic reduction process of an electrolyte. A high electron mobility rate is achieved by a lower value of Rct and causes high electrical outputs, and vice versa. Table 2O4 electrode (7.31 Ω.cm2) is lower than that of the CuAl2O4 electrode (7.56 Ω.cm2), the NiAl2O4 electrode (10.94.cm2), the Al2O3 electrode (7.81 Ω.cm2), and the CoAl2O4 electrode (10.5 Ω.cm2) with CdS QDs60. Due to the good electrocatalytic attitude of ZnAl2O4, it can behave as an efficient WEs catalyst to minimize the oxidized polysulphide electrolyte and high electron mobility rate desired for good photovoltaic performance of QDSSCs. Fig. The impedance of Al2O4, which elevates more electrons from the external circuit. The relocating of electrons during the catalytic reduction process of an electrolyte is facilitated by charge transfer resistance. A lower value of Rct leads to a high electron mobility rate and causes high electrical outputs, and vice versa. Table 2O4 electrode (8.11 Ω cm2) is lower than that of the CuAl2O4 electrode (8.31 Ω.cm2), the Al2O3 electrode (44.75 Ω.cm2), the ZnAl2O4 electrode (49.56 Ω.cm2), and the CoAl2O4 electrode (8.63 Ω.cm2) with hybrid structures of CdS@ZnS QDs60. Due to the good electrocatalytic attitude of the CuAl2O4, it can be behave as an efficient WEs catalyst to minimize the oxidized polysulphide electrolyte and high electron mobility rate desired for good photovoltaic performance of QDSSCs. The kinetic operation of charge transport in QDSSCs, electrochemical impedance spectroscopy (EIS), was executed through the exposition of Al2O3 NPs, ZnAl2O4, CoAl2O4, NiAl2O4, and CuAl2O4 nanocomposites on FTO conductive glass, where by increasing relaxation time, the diffusion rate is decreased, consequently increasing the power activity of the photoanode in the solar cell, which includes the increasing migration of quantum dots on the semiconductor surface in addition to increasing the motion of free electrons in the cell, as indicated by the obtained data. The equivalent circuit is represented in Fig. τn) Eq.  under one sun illumination (10.4 mW*cm−2), respectively. Tables s giving the higher efficiency of ZnAl2O4 nanocomposites (8.22%) surface based on P-rGO CE compared to the efficiency of Al2O3 NPs, NiAl2O4, CoAl2O4, and CuAl2O4 , respectively, shown in Table 2O4 has high chemical, mechanical stability, great separation and transfer of light-generated electrons and holes, which corresponds to the current density of 22.86 mA*cm−2 and fill factor 0.58. (ZnAl2O4) nanocomposite with CdS QDs has the highest efficiency. This is due to several reasons:It has the smallest crystallite size as indicated by XRD Table .It has the lowest band-gap energy Table .4), charge recombination resistance (Rrce = 11.5*105), diffusion rate (Keff = 9.09), and the highest lifetime of the photogenerated electrons (τn = 11*10−2) as indicated by the electrochemical performance , respectively, shown in Table 2O4 photoanode with P-rGO CE. The conversion efficiency of the cell η can be determined by the equation below:SC is the short circuit current density, VOC is the open circuit voltage, FF is the fill factor, and Pin is the power intensity of the incident light. From the J-V curves, FF values can be determined.Zinc oxide has some advantages such that it is a common wide band-gap semiconductor that is now being investigated due to its wide range of applications and adjustable features. The ZnO nanoparticles' wide band-gap semiconductor characteristics are also beneficial in inducing intracellular reactive oxygen species (ROS) production. Conduction electrons (e) and valence holes (h +) in semiconductors have long been employed for photocatalytic oxidation of organic and inorganic contaminants. However, sufficient electrons and holes were routinely generated using UV irradiation and excitation. Even in normal light conditions, great quantities of holes and/or electrons may be accessible in ZnO nanoparticles. P. Sakthivel et al. (2022) show that in the zone wherein light absorption occurs owing to band gap excitation, which reveals that increasing the optical absorption of ZnO by adding defects  to its surface can improve its optical absorption. Aluminum (Al) ions doped on the surface of ZnO nanoparticles boosted the generation of electrons and/or holes in a previous study2O3 NPs and MAl2O4  is greatly affected by the loading of different quantum dots (CdS) and hybrid structures (CdS/ZnS). The hybrid structures of (CdS/ZnS) on metal aluminates showed a high rendering as photoanodes in QDSSCs, whereby by increasing the life time of quantum dots, the diffusion rate is decreased and the (PCE) increased. The highest activity was 15.14% in the cell containing NiAl2O4 spinel with (CdS/ZnS) QDs. The EIS analysis showed that NiAl2O4 has the lowest diffusion rate, Keff, and the highest electron life time, τn.Aluminum Oxide nanoparticles and spinel metal aluminate nanocomposites have been successfully prepared by the auto-combustion sol–gel method. The electrochemical performance of metal aluminates AlSupplementary Information 1.Supplementary Information 2.Supplementary Information 3.Supplementary Information 4.Supplementary Information 5.Supplementary Information 6.Supplementary Information 7.Supplementary Information 8.


Caenorhabditis elegans (CL2006 & GMC101) expressing muscular APP. Model organisms were incubated with either folic acid and magnesium-orotate (ID63) or folic acid, magnesium-orotate and vitamin B6 (ID64) in different concentrations. ID63 and ID64 reduced Aβ, soluble alpha APP (sAPPα), and lactate levels in SH-SY5Y-APP695 cells. The latter might be explained by enhanced expression of lactate dehydrogenase (LDHA). Micronutrient combinations had no effects on mitochondrial parameters in SH-SY5Y-APP695 cells. ID64 showed a significant life-prolonging effect in C. elegans CL2006. Incubation of GMC101 with ID63 significantly lowered Aβ aggregation. Both combinations significantly reduced paralysis and thus improved the phenotype in GMC101. Thus, the combinations of the tested biofactors are effective in pre-clinical models of AD by interfering with Aβ related pathways and glycolysis.Increased amyloid beta (Aβ) levels and mitochondrial dysfunction (MD) in the human brain characterize Alzheimer disease (AD). Folic acid, magnesium and vitamin B6 are essential micro-nutrients that may provide neuroprotection. Bioenergetic parameters and amyloid precursor protein (APP) processing products were investigated in vitro in human neuroblastoma SH-SY5Y-APP695 cells, expressing neuronal APP, and in vivo, in the invertebrate  The strain GMC101 expresses the full-length human Aβ1–42 peptide in body-wall muscle cells that aggregates in vivo. Shifting L4 or young adult animals from 20 °C to 25 °C could induce the expression of Aβ and cause paralysis. The strain CL2006 constitutively expressed Aβ when cultured at 20 °C.E. coli strain OP50. According to standard protocols, the seeded plates were stored at 20 °C [Nematodes were maintained on nematode growth medium (NGM) agar plates seeded with the bacterial at 20 °C . Synchroat 20 °C .worm) and Fol 50 µM, MgOr 0.1 mM and vitamin B6 (Vit B6) 100 µM (together ID64worm). After reaching adulthood (48 h before the experiment), the micronutrients were added to the flasks. Pure M9-buffer was used as a control. Then, 24 h before the experiment amyloid aggregation was proceeded by upshifting young adult GMC101 from 20 °C to 25 °C.Post-bleaching generated larvae were washed twice in M9-buffer and the number of larvae in 10 µL were adjusted to 10 larvae. Afterward, the synchronized larvae were raised in cell culture flasks  in either an amount of 1000 or 5000 nematodes, depending on the experiments. OP50-NGM was added to the flasks as a standardized source of food. The larvae were maintained under shaking at 20 °C until they reached young adulthood within 3 days. The micronutrients were dissolved in advance in M9 buffer. For each micronutrient observed in this study, we generated a series of concentrations as follows. Folic acid (Fol) 50 µM, magnesium orotate (MgOr) 0.1 mM . On a NGM Agar Plate, 25 nematodes were placed and by physically touching with a platinum tip the paralysis status was recorded. Nematodes which normally act after being touched by the wire are recognized as “not paralyzed” whereas uncoordinated movements or just head movements were recorded as “paralyzed”.®20 (1% v/v). In the following step, one nematode in 1 µL M9-buffer was transferred and immersed in the well under a stereomicroscope . SYTOX™ Green  in a final concentration of 1 µM was added to reach a final volume of 15 µL in the well. SYTOX™ Green creates a fluorescent signal after binding to DNA. The plates were sealed with a Rotilab sealing film . The heat-shock was applied and the fluorescence was measured every 30 min for 17 h at 37 °C following the protocol previously described [After 48 h of incubation of CL2006 with the mentioned effectors, the time till death was determined using a microplate thermotolerance assay . In prepescribed . The exc®20 (1% v/v) and separated from larvae. After centrifugation, 200 µL of a thick pellet of nematodes was transferred into a microcentrifuge tube and were frozen in liquid nitrogen. Samples were thawed with 500 µL of PBS including proteinase inhibitor. Afterward, the samples were homogenized with a sonifier 3 × 20 s on ice. Protein contents in the homogenate were assessed according to the Pierce™ BCA Protein Assay Kit . Bovine serum albumin was used as a standard. Finally, fluorescence was measured in a black 96-well plate by adding 1 mM ThT . The volume in each well was 100 µL by adding M9. To determine the fluorescence of ThT, samples were measured by excitation at 440 nm and emission at 482 nm.Detection and quantification of Aβ aggregates (Aβ-aggr) in GMC101 were performed using the fluorescent dye thioflavin T (ThT) according to a previously described method with minor modifications . Synchrot-test . Statistical significance was defined for p values ns = not significant, * p < 0.05, ** p < 0.01, *** p < 0.001 and **** p < 0.0001.Unless otherwise stated, values are presented as mean ± standard error of the mean (SEM). Statistical analyses were performed by applying one-way ANOVA with Tukey’s multiple comparison post-hock test, log-rank (Mantel-Cox) test and student’s unpaired 695 cells and C. elegans. The phenotype of the in vivo model was significantly improved, highlighting the potential of the tested biofactor combinations as candidate therapeutics in AD. However, since the data did not consistently show a benefit of either combination, this study does not allow a clear statement as to whether vitamin B6 is required in addition to the combination of folic acid and magnesium orotate.In the present study, we reported that different combinations of folic acid, magnesium orotate, and vitamin B6 had significant effects on glycolysis, Aβ production, and Aβ aggregation in SH-SY5Y-APP