More Like this

1. Microplastic in riverine fish is connected to species traits

   https://doi.org/10.1038/s41598-018-29980-9  

   McNeish, R. E. ; Kim, L. H. ; Barrett, H. A. ; Mason, S. A. ; Kelly, J. J. ; Hoellein, T. J. ( August 2018 , Scientific Reports)

   Microplastic is a contaminant of concern worldwide. Rivers are implicated as major pathways of microplastic transport to marine and lake ecosystems, and microplastic ingestion by freshwater biota is a risk associated with microplastic contamination, but there is little research on microplastic ecology within freshwater ecosystems. Microplastic uptake by fish is likely affected by environmental microplastic abundance and aspects of fish ecology, but these relationships have rarely been addressed. We measured the abundance and composition of microplastic in fish and surface waters from 3 major tributaries of Lake Michigan, USA. Microplastic was detected in fish and surface waters from all 3 sites, but there was no correlation between microplastic concentrations in fish and surface waters. Rather, there was a significant effect of functional feeding group on microplastic concentration in fish.Neogobius melanostomus(round goby, a zoobenthivore) had the highest concentration of gut microplastic (19 particles fish⁻¹) compared to 10 other fish taxa measured, and had a positive linear relationship between body size and number of microplastic particles. Surface water microplastic concentrations were lowest in the most northern, forested watershed, and highest in the most southern, agriculturally dominated watershed. Results suggest microplastic pollution is common in river food webs and is connected tomore »species feeding characteristics. Future research should focus on understanding the movement of microplastic from point-source and diffuse sources and into aquatic ecosystems, which will support pollution management efforts on inland waters.

   « less
2. Asynchrony of Gambierdiscus spp. Abundance and Toxicity in the U.S. Virgin Islands: Implications for Monitoring and Management of Ciguatera

   https://doi.org/10.3390/toxins13060413  

   Liefer, Justin D. ; Richlen, Mindy L. ; Smith, Tyler B. ; DeBose, Jennifer L. ; Xu, Yixiao ; Anderson, Donald M. ; Robertson, Alison ( June 2021 , Toxins)

   Ciguatera poisoning (CP) poses a significant threat to ecosystem services and fishery resources in coastal communities. The CP-causative ciguatoxins (CTXs) are produced by benthic dinoflagellates including Gambierdiscus and Fukuyoa spp., and enter reef food webs via grazing on macroalgal substrates. In this study, we report on a 3-year monthly time series in St. Thomas, US Virgin Islands where Gambierdiscus spp. abundance and Caribbean-CTX toxicity in benthic samples were compared to key environmental factors, including temperature, salinity, nutrients, benthic cover, and physical data. We found that peak Gambierdiscus abundance occurred in summer while CTX-specific toxicity peaked in cooler months (February–May) when the mean water temperatures were approximately 26–28 °C. These trends were most evident at deeper offshore sites where macroalgal cover was highest year-round. Other environmental parameters were not correlated with the CTX variability observed over time. The asynchrony between Gambierdiscus spp. abundance and toxicity reflects potential differences in toxin cell quotas among Gambierdiscus species with concomitant variability in their abundances throughout the year. These results have significant implications for monitoring and management of benthic harmful algal blooms and highlights potential seasonal and highly-localized pulses in reef toxin loads that may be transferred to higher trophic levels.
3. First integrative study of the diversity and specificity of metacercariae of Posthodiplostomum Dubois, 1936 from native and introduced fishes in the Caribbean

   https://doi.org/10.1017/S0031182022001214  

   Díaz Pernett, Sandra C. ; Brant, Sara V. ; Locke, Sean A. ( December 2022 , Parasitology)

   Abstract Metacercariae of the genus Posthodiplostomum are often recorded in freshwater fish hosts. While the diversity and taxonomy of this genus are receiving increasing attention in molecular phylogenetic studies, available data remain geographically biased. Most molecular studies of Posthodiplostomum and morphologically similar (neascus) worms originate in North America and Europe and Asia (more than 60% of DNA sequences are from USA and Canada), with few data currently available from the Neotropics, where high host diversity suggests high and under-sampled parasite diversity. In this study, we report molecular and morphological data from metacercariae of Posthodiplostomum in fish in Puerto Rico, where only a single species has been previously reported. Partial sequences of cytochrome c oxidase subunit 1 from metacercariae from Dajaus monticola (native to Puerto Rico) and the introduced fishes Poecilia reticulata , Parachromis managuensis , Lepomis macrochirus and Micropterus salmoides revealed 7 genetically distinct species-level lineages, of which 4 were novel. We report novel molecular life-cycle linkages in Posthodiplostomum macrocotyle (metacercariae in muscle of the cichlid Pa. managuensis ), a species previously known only from adults in birds from South America; and in Posthodiplostomum sp. 23 (metacercariae in poeciliids), which has recently been found in Ardea herodias in Georgia, USA.more »We also report the first molecular data from Posthodiplostomum sp. 8 in M. salmoides in the Caribbean. Metacercariae of most species were morphologically distinguished and all displayed narrow specificity for fish hosts, with no indication of parasite sharing among introduced and native fishes.« less
4. Cryptic Methane-Cycling by Methanogens During Multi-Year Incubation of Estuarine Sediment

   https://doi.org/10.3389/fmicb.2022.847563  

   Kevorkian, Richard T. ; Sipes, Katie ; Winstead, Rachel ; Paul, Raegan ; Lloyd, Karen G. ( March 2022 , Frontiers in Microbiology)

   As marine sediments are buried, microbial communities transition from sulfate-reduction to methane-production after sulfate is depleted. When this biogenic methane diffuses into the overlying sulfate-rich sediments, it forms a sulfate-methane transition zone (SMTZ) because sulfate reducers deplete hydrogen concentrations and make hydrogenotrophic methanogenesis exergonic in the reverse direction, a process called the anaerobic oxidation of methane (AOM). Microbial participation in these processes is often inferred from geochemistry, genes, and gene expression changes with sediment depth, using sedimentation rates to convert depth to time. Less is known about how natural sediments transition through these geochemical states transition in real-time. We examined 16S rRNA gene amplicon libraries and metatranscriptomes in microcosms of anoxic sediment from the White Oak River estuary, NC, with three destructively sampled replicates with methane added (586-day incubations) and three re-sampled un-amended replicates (895-day incubations). Sulfate dropped to a low value (∼0.3 mM) on similar days for both experiments (312 and 320 days, respectively), followed by a peak in hydrogen, intermittent increases in methane-cycling archaea starting on days 375 and 362 (mostly Methanolinea spp. and Methanosaeta spp., and Methanococcoides sp. ANME-3), and a methane peak 1 month later. However, methane δ 13 C values only show net methanogenesis 6more »months after methane-cycling archaea increase and 4 months after the methane peak, when sulfate is consistently below 0.1 mM and hydrogen increases to a stable 0.61 ± 0.13 nM (days 553–586, n = 9). Sulfate-reducing bacteria (mostly Desulfatiglans spp. and Desulfosarcina sp. SEEP-SRB1) increase in relative abundance only during this period of net methane production, suggesting syntrophy with methanogens in the absence of sulfate. The transition from sulfate reduction to methane production in marine sediments occurs through a prolonged period of methane-cycling by methanogens at low sulfate concentrations, and steady growth of sulfate reducers along with methanogens after sulfate is depleted.« less
5. Florida mangrove saltmarsh reference surface soils

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

   Lewis, David Bruce ( January 2021 )

   Abstract

   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.
   More>>