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1. Fabrication of highly effective hybrid biofuel cell based on integral colloidal platinum and bilirubin oxidase on gold support

   https://doi.org/10.1038/s41598-018-34740-w  

   Hasan, Md Qumrul ; Kuis, Robinson ; Narayanan, J. Shankara ; Slaughter, Gymama ( November 2018 , Scientific Reports)

   A hybrid biofuel cell (HBFC) is explored as a low-cost alternative to abiotic and enzymatic biofuel cells. Here the HBFC provides an enzymeless approach for the fabrication of the anodic electrode while employing an enzymatic approach for the fabrication of the cathodic electrode to develop energy harvesting platform to power bioelectronic devices. The anode employed 250 μm braided gold wire modified with colloidal platinum (Au-co-Pt) and bilirubin oxidase (BODx) modified gold coated Buckypaper (BP-Au-BODx) cathode. The functionalization of the gold coated multi-walled carbon nanotube (MWCNT) structures of the BP electrodes is achieved by 3-mercaptopropionic acid surface modification to possess negatively charged carboxylic groups and subsequently followed by EDC/Sulfo-NHS (1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride and N-Hydroxysulfosuccinimide) crosslinking with BODx. The integration of the BODx and gold coated MWCNTs is evaluated for bioelectrocatalytic activity. The Au-co-Pt and BP-Au-BODx exhibited excellent electrocatalytic activity towards glucose oxidation with a linear dynamic range up to 20 mM glucose and molecular oxygen reduction, respectively. The HBFC demonstrated excellent performance with the largest open circuit voltages of 0.735 V and power density of 46.31 μW/cm²in 3 mM glucose. In addition, the HBFC operating on 3 mM glucose exhibited excellent uninterrupted operational stability while continuously powering a small electronic device. These results provide great opportunities for implementing this simple but efficient HBFC to harvest the biochemical energy of target fuel(s) in diverse medical and environmental applications.

    

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2. Laser-induced atmospheric Cu x O formation on copper surface with enhanced electrochemical performance for non-enzymatic glucose sensing

   https://doi.org/10.1039/d1tc01289d  

   Sedaghat, Sotoudeh ; Nejati, Sina ; Bermejo, Luis Helena ; He, Zihao ; Alcaraz, Alejandro M. ; Roth, Alexander ; Li, Zheng ; Pol, Vilas G. ; Wang, Haiyan ; Rahimi, Rahim ( November 2021 , Journal of Materials Chemistry C)

   Copper oxide nanostructures are widely used for various applications due to their unique optical and electrical properties. In this work, we demonstrate an atmospheric laser-induced oxidation technique for the fabrication of highly electrochemically active copper oxide hierarchical micro/nano structures on copper surfaces to achieve highly sensitive non-enzymatic glucose sensing performance. The effect of laser processing power on the composition, crystallinity, microstructure, wettability, and color of the laser-induced oxide on copper (LIO-Cu) surface was systematically studied using scanning electron microscopy (SEM), grazing incidence X-ray diffraction (GI-XRD), Raman spectroscopy, energy dispersive X-ray spectroscopy (EDX), EDX-mapping, water contact angle measurements, and optical microscopy. Results of these investigations showed a remarkable increase in copper oxide composition by increasing the laser processing power. The pore size distribution and surface area of the pristine and LIO-Cu sample estimated by N 2 adsorption–desorption data showed a developed mesoporous LIO-Cu structure. The size of the generated nano-oxides, crystallinity, and electroactivity of the LIO-Cu were observed to be adjustable by the laser processing power. The electrocatalytic activity of LIO-Cu surfaces was studied by means of cyclic voltammetry (CV) within a potential window of −0.8 to +0.8 V and chronoamperometry in an applied optimized potential of +0.6 V, in 0.1 M NaOH solution and phosphate buffer solution (PBS), respectively. LIO-Cu surfaces with optimized laser processing powers exhibited a sensitivity of 6950 μA mM −1 cm −2 within a wide linear range from 0.01 to 5 mM, with exceptional specificity and response time (<3 seconds). The sensors also showed excellent response stability over a course of 50 days that was originated from the binder-free robust electroactive film fabricated directly onto the copper surface. The demonstrated one-step LIO processing onto commercial metal films, can potentially be applied for tuneable and scalable roll-to-roll fabrication of a wide range of high surface area metal oxide micro/nano structures for non-enzymatic biosensing and electrochemical applications. 

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3. In situ formation of metal–organic framework derived CuO polyhedrons on carbon cloth for highly sensitive non-enzymatic glucose sensing

   https://doi.org/10.1039/c9tb01166h  

   Cheng, Siyi ; Gao, Xiang ; DelaCruz, Steven ; Chen, Chen ; Tang, Zirong ; Shi, Tielin ; Carraro, Carlo ; Maboudian, Roya ( August 2019 , Journal of Materials Chemistry B)

   Metal–organic frameworks (MOFs) are considered promising templates for the fabrication of nanostructured materials with high porosities and high surface areas, which are important parameters for enhanced performance in sensing applications. Here, a facile in situ synthetic strategy to construct MOF-derived porous CuO polyhedrons on carbon cloth (CC) is reported. Uniform Cu(OH) 2 nanorods are first synthesized on carbon cloth, followed by the conversion of Cu(OH) 2 nanorods into porous CuO polyhedrons via a copper-based MOF, Cu–BTC, as the intermediate species. When evaluated as a glucose sensing electrode, the as-fabricated CuO polyhedrons/CC composite exhibits a high sensitivity of 13 575 μA mM −1 cm −2 with a fast response time ( t 90 ) of 2.3 s and a low detection limit of 0.46 μM. This work exemplifies the rational fabrication of porous nanostructures on conductive substrates for enhanced performance in glucose detection. 

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4. Structural control of nickel nanowire arrays for use as non‐enzymatic glucose sensors

   https://doi.org/10.1002/elan.202300200  

   Shey, Nicholas H. ; Margetts, Rebecca M. ; Gillette, Eleanor I. ( November 2023 , Electroanalysis)

   A series of non‐enzymatic electrochemical glucose sensors were prepared by template‐assisted electrodeposition of nickel to produce nickel nanowire arrays (NiNWAs) of varying lengths. These electrodes were analyzed for their performance as non‐enzymatic glucose sensors, in order to explore the relationship between nanowire structure and electrochemical sensor performance. All of the NiNWAs, regardless of length, had linear responses to glucose in the 0.05–10 mM range. However, an inverse relationship between wire length and sensitivity was observed: the shortest of the NiNWAs had a sensitivity of 160 μA mM⁻¹ cm⁻², while the longest NiNWA had a sensitivity of only 100 μA mM⁻¹ cm⁻². This indicates that, with no other change to material parameters, choosing an optimum length of nanowire provides significant improvement in performance. No change in electrochemical mechanism was observed by cyclic voltammetry, and no significant loss of selectivity was observed with varying nanowire length. This study demonstrates that the structural optimization of an electrode can provide significant performance enhancement for nickel‐based electrochemical sensors.

    

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5. Florida mangrove saltmarsh reference surface soils

   https://doi.org/10.6073/pasta/0e08cbe07c84488cb7b9dd16669946d0  

   Lewis, David Bruce ( January 2021 , Environmental Data Initiative)

   Site description. This data package consists of data obtained from sampling surface soil (the 0-7.6 cm depth profile) in black mangrove (Avicennia germinans) dominated forest and black needlerush (Juncus roemerianus) saltmarsh along the Gulf of Mexico coastline in peninsular west-central Florida, USA. This location has a subtropical climate with mean daily temperatures ranging from 15.4 °C in January to 27.8 °C in August, and annual precipitation of 1336 mm. Precipitation falls as rain primarily between June and September. Tides are semi-diurnal, with 0.57 m median amplitudes during the year preceding sampling (U.S. NOAA National Ocean Service, Clearwater Beach, Florida, station 8726724). Sea-level rise is 4.0 ± 0.6 mm per year (1973-2020 trend, mean ± 95 % confidence interval, NOAA NOS Clearwater Beach station). The A. germinans mangrove zone is either adjacent to water or fringed on the seaward side by a narrow band of red mangrove (Rhizophora mangle). A near-monoculture of J. roemerianus is often adjacent to and immediately landward of the A. germinans zone. The transition from the mangrove to the J. roemerianus zone is variable in our study area. An abrupt edge between closed-canopy mangrove and J. roemerianus monoculture may extend for up to several hundred meters in some locations, while other stretches of ecotone present a gradual transition where smaller, widely spaced trees are interspersed into the herbaceous marsh. Juncus roemerianus then extends landward to a high marsh patchwork of succulent halophytes (including Salicornia bigellovi, Sesuvium sp., and Batis maritima), scattered dwarf mangrove, and salt pans, followed in turn by upland vegetation that includes Pinus sp. and Serenoa repens. Field design and sample collection. We established three study sites spaced at approximately 5 km intervals along the western coastline of the central Florida peninsula. The sites consisted of the Salt Springs (28.3298°, -82.7274°), Energy Marine Center (28.2903°, -82.7278°), and Green Key (28.2530°, -82.7496°) sites on the Gulf of Mexico coastline in Pasco County, Florida, USA. At each site, we established three plot pairs, each consisting of one saltmarsh plot and one mangrove plot. Plots were 50 m^2 in size. Plots pairs within a site were separated by 230-1070 m, and the mangrove and saltmarsh plots composing a pair were 70-170 m apart. All plot pairs consisted of directly adjacent patches of mangrove forest and J. roemerianus saltmarsh, with the mangrove forests exhibiting a closed canopy and a tree architecture (height 4-6 m, crown width 1.5-3 m). Mangrove plots were located at approximately the midpoint between the seaward edge (water-mangrove interface) and landward edge (mangrove-marsh interface) of the mangrove zone. Saltmarsh plots were located 20-25 m away from any mangrove trees and into the J. roemerianus zone (i.e., landward from the mangrove-marsh interface). Plot pairs were coarsely similar in geomorphic setting, as all were located on the Gulf of Mexico coastline, rather than within major sheltering formations like Tampa Bay, and all plot pairs fit the tide-dominated domain of the Woodroffe classification (Woodroffe, 2002, "Coasts: Form, Process and Evolution", Cambridge University Press), given their conspicuous semi-diurnal tides. There was nevertheless some geomorphic variation, as some plot pairs were directly open to the Gulf of Mexico while others sat behind keys and spits or along small tidal creeks. Our use of a plot-pair approach is intended to control for this geomorphic variation. Plot center elevations (cm above mean sea level, NAVD 88) were estimated by overlaying the plot locations determined with a global positioning system (Garmin GPS 60, Olathe, KS, USA) on a LiDAR-derived bare-earth digital elevation model (Dewberry, Inc., 2019). The digital elevation model had a vertical accuracy of ± 10 cm (95 % CI) and a horizontal accuracy of ± 116 cm (95 % CI). Soil samples were collected via coring at low tide in June 2011. From each plot, we collected a composite soil sample consisting of three discrete 5.1 cm diameter soil cores taken at equidistant points to 7.6 cm depth. Cores were taken by tapping a sleeve into the soil until its top was flush with the soil surface, sliding a hand under the core, and lifting it up. Cores were then capped and transferred on ice to our laboratory at the University of South Florida (Tampa, Florida, USA), where they were combined in plastic zipper bags, and homogenized by hand into plot-level composite samples on the day they were collected. A damp soil subsample was immediately taken from each composite sample to initiate 1 y incubations for determination of active C and N (see below). The remainder of each composite sample was then placed in a drying oven (60 °C) for 1 week with frequent mixing of the soil to prevent aggregation and liberate water. Organic wetland soils are sometimes dried at 70 °C, however high drying temperatures can volatilize non-water liquids and oxidize and decompose organic matter, so 50 °C is also a common drying temperature for organic soils (Gardner 1986, "Methods of Soil Analysis: Part 1", Soil Science Society of America); we accordingly chose 60 °C as a compromise between sufficient water removal and avoidance of non-water mass loss. Bulk density was determined as soil dry mass per core volume (adding back the dry mass equivalent of the damp subsample removed prior to drying). Dried subsamples were obtained for determination of soil organic matter (SOM), mineral texture composition, and extractable and total carbon (C) and nitrogen (N) within the following week. Sample analyses. A dried subsample was apportioned from each composite sample to determine SOM as mass loss on ignition at 550 °C for 4 h. After organic matter was removed from soil via ignition, mineral particle size composition was determined using a combination of wet sieving and density separation in 49 mM (3 %) sodium hexametaphosphate ((NaPO_3)_6) following procedures in Kettler et al. (2001, Soil Science Society of America Journal 65, 849-852). The percentage of dry soil mass composed of silt and clay particles (hereafter, fines) was calculated as the mass lost from dispersed mineral soil after sieving (0.053 mm mesh sieve). Fines could have been slightly underestimated if any clay particles were burned off during the preceding ignition of soil. An additional subsample was taken from each composite sample to determine extractable N and organic C concentrations via 0.5 M potassium sulfate (K_2SO_4) extractions. We combined soil and extractant (ratio of 1 g dry soil:5 mL extractant) in plastic bottles, reciprocally shook the slurry for 1 h at 120 rpm, and then gravity filtered it through Fisher G6 (1.6 μm pore size) glass fiber filters, followed by colorimetric detection of nitrite (NO_2^-) + nitrate (NO_3^-) and ammonium (NH_4^+) in the filtrate (Hood Nowotny et al., 2010,Soil Science Society of America Journal 74, 1018-1027) using a microplate spectrophotometer (Biotek Epoch, Winooski, VT, USA). Filtrate was also analyzed for dissolved organic C (referred to hereafter as extractable organic C) and total dissolved N via combustion and oxidation followed by detection of the evolved CO_2 and N oxide gases on a Formacs HT TOC/TN analyzer (Skalar, Breda, The Netherlands). Extractable organic N was then computed as total dissolved N in filtrate minus extractable mineral N (itself the sum of extractable NH_4-N and NO_2-N + NO_3-N). We determined soil total C and N from dried, milled subsamples subjected to elemental analysis (ECS 4010, Costech, Inc., Valencia, CA, USA) at the University of South Florida Stable Isotope Laboratory. Median concentration of inorganic C in unvegetated surface soil at our sites is 0.5 % of soil mass (Anderson, 2019, Univ. of South Florida M.S. thesis via methods in Wang et al., 2011, Environmental Monitoring and Assessment 174, 241-257). Inorganic C concentrations are likely even lower in our samples from under vegetation, where organic matter would dilute the contribution of inorganic C to soil mass. Nevertheless, the presence of a small inorganic C pool in our soils may be counted in the total C values we report. Extractable organic C is necessarily of organic C origin given the method (sparging with HCl) used in detection. Active C and N represent the fractions of organic C and N that are mineralizable by soil microorganisms under aerobic conditions in long-term soil incubations. To quantify active C and N, 60 g of field-moist soil were apportioned from each composite sample, placed in a filtration apparatus, and incubated in the dark at 25 °C and field capacity moisture for 365 d (as in Lewis et al., 2014, Ecosphere 5, art59). Moisture levels were maintained by frequently weighing incubated soil and wetting them up to target mass. Daily CO_2 flux was quantified on 29 occasions at 0.5-3 week intervals during the incubation period (with shorter intervals earlier in the incubation), and these per day flux rates were integrated over the 365 d period to compute an estimate of active C. Observations of per day flux were made by sealing samples overnight in airtight chambers fitted with septa and quantifying headspace CO_2 accumulation by injecting headspace samples (obtained through the septa via needle and syringe) into an infrared gas analyzer (PP Systems EGM 4, Amesbury, MA, USA). To estimate active N, each incubated sample was leached with a C and N free, 35 psu solution containing micronutrients (Nadelhoffer, 1990, Soil Science Society of America Journal 54, 411-415) on 19 occasions at increasing 1-6 week intervals during the 365 d incubation, and then extracted in 0.5 M K_2SO_4 at the end of the incubation in order to remove any residual mineral N. Active N was then quantified as the total mass of mineral N leached and extracted. Mineral N in leached and extracted solutions was detected as NH_4-N and NO_2-N + NO_3-N via colorimetry as above. This incubation technique precludes new C and N inputs and persistently leaches mineral N, forcing microorganisms to meet demand by mineralizing existing pools, and thereby directly assays the potential activity of soil organic C and N pools present at the time of soil sampling. Because this analysis commences with disrupting soil physical structure, it is biased toward higher estimates of active fractions. Calculations. Non-mobile C and N fractions were computed as total C and N concentrations minus the extractable and active fractions of each element. This data package reports surface-soil constituents (moisture, fines, SOM, and C and N pools and fractions) in both gravimetric units (mass constituent / mass soil) and areal units (mass constituent / soil surface area integrated through 7.6 cm soil depth, the depth of sampling). Areal concentrations were computed as X × D × 7.6, where X is the gravimetric concentration of a soil constituent, D is soil bulk density (g dry soil / cm^3), and 7.6 is the sampling depth in cm. 

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