More stories

  • in

    Fungal infections lead to shifts in thermal tolerance and voluntary exposure to extreme temperatures in both prey and predator insects

    Field trialsField trials were conducted in three raised beds (1 × 2 × 0.6 m) on the Penn State University campus from July to August 2020. The raised beds were separated by at least 8 m to avoid treatment cross-contamination. Faba bean (Vicia faba L.) seeds were planted at a density of 20 seeds/ m2 (50 plants per bed), and each bed was caged using a metal-framed tent. “Noseeum” nylon mesh (Outdoor Wilderness Fabric s, Inc., Caldwell, ID) was draped over the frame and the edges buried in the soil of the bed. The sides of the cages were fastened closed with zippers to allow access.InsectsAphid and predator beetle colonies were raised separately on faba bean plants in cages (BugDorm 20 cm × 40 cm × 20 cm, BioQuip Products, Inc., Rancho Dominguez, CA) in the field. Larvae and adults of predator beetles were fed with a combination of A. pisum and Rhopalosiphum padi every other day (Supplementary information Fig. S1). Trials involving plants, insects, and entomopathogenic fungi were conducted according to institutional, national, and international guidelines and legislation.Fungal inoculations (Beavueria bassiana)We released first instar aphid nymphs on each faba bean plant on the raised beds (~ 1100 aphids) by gently shaking plastic containers with groups of 20 nymphs and placing them on the plants using a paintbrush. They were allowed to grow and reproduce for fifteen days. During the night, we sprayed spore suspension of the Beauveria strain GHA (BotaniGard ®, MT, USA) at 1.4 × 106 and 1.4 × 1012 spore ha−1, low and high load respectively. Two days after inoculation, we collected adult aphids (~ 4–5 days old) from the experimental plots and measured physiological parameters (see details below). Next, we released 300 adult beetles inside each aphid–fungal inoculated cage, allowed them to feed for 2–3 days in our experimental cages, and then collected beetles for physiological measurement.Identification of critical thermal limits (CTMax and CTMin) of healthy and infected insectsTo determine critical thermal maximum for locomotion (CTMax) of healthy and infected individuals of each species, we employed a protocol modified from Ribeiro et al.25, using a hotplate with a programmable heating rate controlled by a computer interface (Sable Systems, LV, USA). The temperature was monitored by independent thermocouple channels connected to a Hobo 4-channel data logger. One thermocouple was attached to the surface of the hotplate, and the other sensor was attached inside the glass tube plugged by a cotton ball in which we placed an individual insect. This equipment was located inside an automated thermal chamber (interior dimensions: width 40.5 cm × 35 cm length × 40 cm height). We transferred an adult aphid (4-day-old) into the glass tube and exposed it to increasing temperatures at a rate of 0.3 °C min−1 until its locomotion stopped. CTMax was recorded when the insect turned upside down and could no longer return to the upright position within 5 s. The insect was returned to a faba bean leaf for recovery (n = 10 individuals per treatment).To measure the critical thermal minimum for locomotion (CTMin) of healthy and infected individuals of each species (n = 10 individuals per treatment), we used an insulated incubator where the temperature was monitored by independent thermocouple channels connected to a Hobo 4-channel data logger. The sensors were attached inside three glass tubes, each tube with an adult (3 to 4-day-old), and plugged by a cotton ball. The glass tube was exposed to decreasing temperature at a rate of 0.3 °C min−1 until its locomotion stopped. CTMin was recorded when no movement was recorded within 5 s. The insect was returned to an aphid-infested faba bean leaf for recovery. Data were only considered valid if the insect displayed normal activity 2 h after a CTMax or CTMin test.Impacts of infection on voluntary exposure aphids and predator beetles to extreme thermal zonesTo examine how voluntary exposure to ETZ was affected by fungal infection, we collected aphids and predator beetles (3 to 5 day-old) from our field plots and transferred them to a dark plastic bottle. Next, a bottle containing the insects was attached to a choice test arena following a modified protocol from Navas et al.24. This experimental arena allows insects to freely move across extreme temperatures to access food in containers located at each end of the device. To reach food, individuals had to cross an ETZ, either warm or cold. The location of each insect was recorded after 60 min, and it was classified as: exploration for individuals that left the initial black bottle, warm or cold ETZ crossings. The experiment was replicated ten times for each species and treatment condition [aphid: healthy, infected (low and high spore load); predator beetle: healthy, infected (low and high spore load)].Effects of fungal infection and thermal conditions (critical thermal limits and voluntary exposure to ETZs) on longevity of aphids and predator beetlesTo examine whether fungal infection and thermal conditions alter longevity in aphids and beetles, we isolated three individuals from each factor combination (low, high fungal load, CTMin, CTMax, behavior: crosses to ETZ cold, warm, and no cross) from previous experiments, and counted the number of days the adults survived after the exposure to the thermal condition (n = 3 factor combination).Energetic cost associated with fungal infection of aphid and predator beetles under critical thermal limits and voluntary exposure to ETIntracellular ATP content was determined in neutralized perchloric acid extracts and by a spectrophotometric coupled enzyme assay, based on modified protocol from Churchill and Storey26 content (n = 3 per treatment condition). An insect was ground to powder using a mortar and pestle cooled in liquid nitrogen, and then weighed into 1.5 mL microcentrifuge tubes (Eppendorf). Powder was dissolved with 0.1 mL ice-cold TE buffer (50 mM Tris–HCl, pH 7.5 plus 1 mM EGTA) and homogenized by sonication (15 s, three times), using a Q500 Sonicator system (QSonica, Newtown, CT, USA). An aliquot (10 µL) of the well-mixed homogenate was removed for protein determination. Cells were lysed by adding 6% (v/v) ice-cold perchloric acid, strongly vortexed for 2 min and incubated at 4 °C for 10 min. Next, the cell homogenate was centrifuged at 14,462 rpm and 4 °C for 5 min. The resulting supernatant was neutralized by adding KOH/Tris (3 M/0.1 M) and centrifuged again to discard the perchlorate salts. Extracts were kept at 4 °C for their immediate utilization. ATP content was determined spectrophotometrically by following the production of NADPH at 340 nm (ε = 6.22 mM−1 cm−1) and using CARY WinUV-Vis Spectrophotometer (Agilent, Santa Clara, CA, USA). The following reagents were used for the spectrophotometric coupled enzyme assay: 5 U Hexokinase, 10 U Glucose 6-phosphate dehydrogenase, 1 mM NADP + , 5 mM MgCl2 and 10 mM Glucose in HE buffer (100 mM Hepes-HCl plus 1 mM EGTA, pH 7.0) at 25 °C. Chemicals were purchased from Roche (Manheim, Germany) and Sigma (St Louis, MO, USA).Infection statusWe used two different protocols to confirm fungal infection: (1) placing each individual in wet towel paper inside a Ziploc bag to observe hyphal growth27. (2) For insects used in ATP measurements, we followed a modified protocol from Wraight and Ramos28 and Castrillo et al.29. Insect were washed using a serial dilution technique, vortexed for 10 s, and mounted in a drop of lactophenol blue, diluted with distilled water. We then preserved insect body parts (i.e., legs and abdomen terga) at − 80 °C for 12 months and placed in Petri dishes containing potato dextrose agar (PDA HiMedia-GM096) medium (pH 6.8), and incubated for ten days. To confirm infection by B. bassiana, we observed plates every 3 days, identified fungal growth (dense white mycelia), then randomly chose three samples, collected mycelia, and DNA was extracted using PureLink Genomic DNA Kit (Invitrogen by Thermo Fisher Scientific, Waltham, MA, USA), according to manufacturer’s protocol. Next, we used PCR essays (25 µL) contained 1 × Q5 Hot Start High-Fidelity Master Mix (New England BioLabs), following a protocol modified from Castrillo et al.29 using primers GHTqF1 (5′-TTTTCATCGAAAGGTTGTTTCTCG) and GHTq R1 (5′-CTGTGCTGGGTACTGACGTG) amplified a 96-bp region of the SCAR fragment. The PCR protocol was initial denaturation at 98 °C, followed by 30 cycles at 98 °C for 1 min, annealing at 58 °C for 1 min; and extension at 72 °C for 1 min. PCR products were visualized in a 1.0% (wt/vol) agarose gel stained with ethidium bromide.Data analysisAll data were tested for statistical test assumptions using a qqplot, Levene’s homogeneity test and the Shapiro–Wilk normality test at alpha = 0.05 significance level. For critical thermal limits (CTMax and CTMin) experiments, the data sets were non-normal and transformation did not normalize the residuals, so we used nonparametric ANOVAs (Kruskal–Wallis) followed by post-hoc nonparametric multiple comparisons. For voluntary exposure to ETZs, we used a generalized linear model with treatment (healthy, low and high spore load) with Poisson distribution, followed by comparisons within each treatment group. For healthy insects, we used a t-test to compare crosses between warm or cold ETZs; for infected insects, we conducted ANOVAS for comparisons among 23 °C, warm or cold ETZs.ATP data: Data for CTMax of A. pisum were non-normal, and transformation did not normalize the residuals, nonparametric ANOVAs (Kruskal–Wallis) were then used and followed by post-hoc nonparametric pairwise comparisons with Wilcoxon tests. ATP data sets from voluntary exposure to ETZs were analyzed following the same protocol as described previously for in crosses analysis of ETZ experiment. Longevity was analyzed using a two-way ANOVA with fungal load and thermal condition (critical temperature and behavior) as factors. Analyses were performed in the R programming environment (v. 3.4.3., CRAN project)30 and JMP-Pro version 15 (SAS Institute 2020). More

  • in

    Molecular species delimitation refines the taxonomy of native and nonnative physinine snails in North America

    1.Mayr, E. The species concept: Semantics versus semantics. Evolution 3, 371–372 (1949).Article 

    Google Scholar 
    2.Hortal, J. et al. Seven shortfalls that beset large-scale knowledge of biodiversity. Annu. Rev. Ecol. Evol. Syst. 46, 523–549 (2015).Article 

    Google Scholar 
    3.Mace, G. M. The role of taxonomy in species conservation. Philos. Trans. R. Soc. Lond. B Biol. Sci. 359, 711–719 (2004).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    4.Gustafson, K. D., Kensinger, B. J., Bolek, M. G. & Luttbeg, B. Distinct snail (Physa) morphotypes from different habitats converge in shell shape and size under common garden conditions. Evol. Ecol. Res. 16, 77–89 (2014).
    Google Scholar 
    5.Aksenova, O. V. et al. Species richness, molecular taxonomy and biogeography of the radicine pond snails (Gastropoda: Lymnaeidae) in the Old World. Sci. Rep. 8, 1–7 (2018).ADS 
    CAS 
    Article 

    Google Scholar 
    6.Liu, H. P. & Hershler, R. A new species and range extensions for three other species of pebblesnails (Lithoglyphidae, Fluminicola) from the upper Klamath basin, California-Oregon. ZooKeys 812, 47–67 (2019).Article 

    Google Scholar 
    7.Alda, P. et al. Systematics and geographical distribution of Galba species, a group of cryptic and worldwide freshwater snails. Mol. Phylogenet. Evol. 157, 107035 (2021).PubMed 
    Article 

    Google Scholar 
    8.Taylor, D. W. Introduction to Physidae (Gastropoda: Hygrophila); biogeography, classification, morphology. Rev. Biol. Trop. 51(Supplement 1), 1–287 (2003).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    9.Wethington, A. R. & Lydeard, C. A molecular phylogeny of Physidae (Gastropoda: Basommatophora) based on mitochondrial DNA sequences. J. Molluscan Stud. 73, 241–257 (2007).Article 

    Google Scholar 
    10.Ng, T. H. et al. Molluscs for sale: assessment of freshwater gastropods and bivalves in the ornamental pet trade. PLoS ONE 11, e0161130 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    11.Saito, T., Prozorova, L., Hirano, T., Fukuda, H. & Chiba, S. Endangered freshwater limpets in Japan are actually alien invasive species. Conserv. Genet. 19, 947–958 (2018).Article 

    Google Scholar 
    12.Lydeard, C., Campbell, D. & Golz, M. Physa acuta Draparnaud, 1805 should be treated as a native of North America, not Europe. Malacologia 59, 347–350 (2016).Article 

    Google Scholar 
    13.Albrecht, C., Kroll, O., Terrazas, E. M. & Wilke, T. Invasion of ancient Lake Titicaca by the globally invasive Physa acuta (Gastropoda: Pulmonata: Hygrophila). Biol. Invasions 11, 1821–1826 (2009).Article 

    Google Scholar 
    14.Ng, T. H., Tan, S. K. & Yeo, D. C. Clarifying the identity of the long-established, globally-invasive Physa acuta Draparnaud, 1805 (Gastropoda: Physidae) in Singapore. BioInvasions Rec. 4, 189–194 (2015).Article 

    Google Scholar 
    15.Collado, G. A. Unraveling cryptic invasion of a freshwater snail in Chile based on molecular and morphological data. Biodivers. Conserv. 26, 567–578 (2017).Article 

    Google Scholar 
    16.Johnson, P. D. et al. Conservation status of freshwater gastropods of Canada and the United States. Fisheries 38, 247–282 (2013).Article 

    Google Scholar 
    17.Strong, E. E. & Whelan, N. V. Assessing the diversity of western North American Juga (Semisulcospiridae, Gastropoda). Mol. Phylogenet. Evol. 136, 87–103 (2019).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    18.Hebert, P. D., Ratnasingham, S. & De Waard, J. R. Barcoding animal life: cytochrome c oxidase subunit 1 divergences among closely related species. Proc. R. Soc. Lond. B 270(supplement 1), S96-99 (2003).CAS 

    Google Scholar 
    19.Stöger, I. & Schrödl, M. Mitogenomics does not resolve deep molluscan relationships (yet?). Mol. Phylogenet. Evol. 69, 376–392 (2013).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    20.Cunha, T. J. & Giribet, G. A congruent topology for deep gastropod relationships. Proc. R. Soc. B 286, 20182776 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    21.Varney, R. M. et al. Assessment of mitochondrial genomes for heterobranch gastropod phylogenetics. BMC Ecol. Evol. 21, 6 (2021).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    22.Remigio, E. A. & Hebert, P. D. Testing the utility of partial COI sequences for phylogenetic estimates of gastropod relationships. Mol. Phylogenet. Evol. 29, 641–647 (2003).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    23.Collins, R. A. & Cruickshank, R. H. The seven deadly sins of DNA barcoding. Mol. Ecol. Resour. 13, 969–975 (2013).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    24.Ratnasingham, S. & Hebert, P. D. A DNA-based registry for all animal species: The Barcode Index Number (BIN) system. PLoS ONE 8, e66213 (2013).ADS 
    CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    25.Whelan, N. V. & Strong, E. E. Morphology, molecules and taxonomy: Extreme incongruence in pleurocerids (Gastropoda, Cerithioidea, Pleuroceridae). Zoolog. Scr. 45, 62–87 (2016).Article 

    Google Scholar 
    26.Razkin, O., Gómez-Moliner, B. J., Vardinoyannis, K., Martínez-Ortí, A. & Madeira, M. J. Species delimitation for cryptic species complexes: Case study of Pyramidula (Gastropoda, Pulmonata). Zool. Scr. 46, 55–72 (2017).Article 

    Google Scholar 
    27.Liu, H. P., Hershler, R. & Hovingh, P. Molecular evidence enables further resolution of the western North American Pyrgulopsis kolobensis complex (Caenogastropoda: Hydrobiidae). J. Molluscan Stud. 84, 103–107 (2018).Article 

    Google Scholar 
    28.Ward, R. D. DNA barcode divergence among species and genera of birds and fishes. Mol. Ecol. Resour. 9, 1077–1085 (2009).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    29.Saadi, A. J., Davison, A. & Wade, C. M. Molecular phylogeny of freshwater snails and limpets (Panpulmonata: Hygrophila). Zool. J. Linn. Soc. 190, 518–531 (2020).Article 

    Google Scholar 
    30.Frest, T. J. & Johannes, E. J. An annotated checklist of Idaho land and freshwater mollusks. J. Idaho Acad. Sci. 36(2), 1–51 (2000).
    Google Scholar 
    31.Pip, E. & Franck, J. P. Molecular phylogenetics of central Canadian Physidae (Pulmonata: Basommatophora). Can. J. Zool. 86, 10–16 (2008).CAS 
    Article 

    Google Scholar 
    32.Tariel, J., Plénet, S. & Luquet, É. Transgenerational plasticity of inducible defences: Combined effects of grand-parental, parental and current environments. Ecol. Evol. 10, 2367–2376 (2020).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    33.Perrin, N. The life history parameters of Physa acuta (Gastropoda, Mollusca) in experimental conditions. Revue Suisse Zoologique 93, 725–736 (1986).Article 

    Google Scholar 
    34.Taylor, D. W. New species of Physa (Gastropoda: Hygrophila) from the western United States. Malacol. Rev. 21, 43–79 (1988).
    Google Scholar 
    35.U.S. Fish and Wildlife Service. Determination of endangered or threatened status for five aquatic snails in south central Idaho. Fed. Reg. 57, 59242–59257 (1992).
    Google Scholar 
    36.Rogers, D. C. & Wethington, A. R. Physa natricina Taylor 1988, junior synonym of Physa acuta Draparnaud, 1805 (Pulmonata: Physidae). Zootaxa 1662, 45–51 (2007).
    Google Scholar 
    37.Gates, K. K., Kerans, B. L., Keebaugh, J. L., Kalinowski, S. & Vu, N. Taxonomic identity of the endangered Snake River physa, Physa natricina (Pulmonata: Physidae) combining traditional and molecular techniques. Conserv. Genet. 14, 159–169 (2013).Article 

    Google Scholar 
    38.Moore, A. C., Burch, J. B. & Duda, T. F. Recognition of a highly restricted freshwater snail lineage (Physidae: Physella) in southeastern Oregon: Convergent evolution, historical context, and conservation considerations. Conserv. Genet. 16, 113–123 (2015).Article 

    Google Scholar 
    39.Dillon, R. T., Robinson, J. D. & Wethington, A. R. Empirical estimates of reproductive isolation among the freshwater pulmonate snails Physa acuta, P. pomilia, and P. hendersoni. Malacologia 49, 283–292 (2007).Article 

    Google Scholar 
    40.De Queiroz, K. Species concepts and species delimitation. Syst. Biol. 56, 879–886 (2007).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    41.Dyke, A. S., Moore, A. & Robertson, L. Deglaciation of North America. Geological Survey of Canada Open File 1574 (2003).42.Wethington, A. R., Wise, J. & Dillon, R. T. Jr. Genetic and morphological characterization of the Physidae of South Carolina (Gastropoda: Pulmonata: Basommatophora), with description of a new species. Nautilus 123, 282–292 (2009).
    Google Scholar 
    43.Ebbs, E. T., Loker, E. S. & Brant, S. V. Phylogeography and genetics of the globally invasive snail Physa acuta Draparnaud 1805, and its potential to serve as an intermediate host to larval digenetic trematodes. BMC Evol. Biol. 18, 1–7 (2018).Article 

    Google Scholar 
    44.Duggan, I. C. The freshwater aquarium trade as a vector for incidental invertebrate fauna. Biol. Invasions 12, 3757–3770 (2010).Article 

    Google Scholar 
    45.Van Leeuwen, C. H. et al. How did this snail get here? Several dispersal vectors inferred for an aquatic invasive species. Freshw. Biol. 58, 88–99 (2013).Article 

    Google Scholar 
    46.Coughlan, N. E., Kelly, T. C., Davenport, J. & Jansen, M. A. Up, up and away: Bird-mediated ectozoochorous dispersal between aquatic environments. Freshw. Biol. 62, 631–648 (2017).Article 

    Google Scholar 
    47.Bony, Y. K. et al. Ecological conditions for spread of the invasive snail Physa marmorata (Pulmonata: Physidae) in the Ivory Coast. Afr. Zool. 43, 53–60 (2008).Article 

    Google Scholar 
    48.Pierce, K. L. & Morgan, L. A. Is the track of the Yellowstone hotspot driven by a deep mantle plume?—Review of volcanism, faulting, and uplift in light of new data. J. Volcanol. Geotherm. Res. 188, 1–25 (2009).ADS 
    CAS 
    Article 

    Google Scholar 
    49.Smith, G. R. et al. Biogeography and timing of evolutionary events among Great Basin fishes. In Great Basin Aquatic Systems History. Smithsonian Contributions to the Earth Sciences Vol. 33 (eds Hershler, R. et al.) 175–234 (Smithsonian Institution Press, 2002).
    Google Scholar 
    50.Oviatt, C. G. Chronology of Lake Bonneville, 30,000 to 10,000 yr BP. Quatern. Sci. Rev. 110, 166–171 (2015).Article 

    Google Scholar 
    51.Safran, E. B. et al. Plugs or flood-makers? The unstable landslide dams of eastern Oregon. Geomorphology 248, 237–251 (2015).ADS 
    Article 

    Google Scholar 
    52.Ely, L. L. et al. Owyhee River intracanyon lava flows: Does the river give a dam?. GSA Bull. 124, 1667–1687 (2012).Article 

    Google Scholar 
    53.Matthews, J. et al. Rapid range expansion of the invasive quagga mussel in relation to zebra mussel presence in the Netherlands and western Europe. Biol. Invasions 16, 23–42 (2014).Article 

    Google Scholar 
    54.Folmer, O., Black, M., Hoeh, W., Lutz, R. & Vrijenhoek, R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechol. 3, 294–299 (1994).CAS 

    Google Scholar 
    55.Geller, J., Meyer, C., Parker, M. & Hawk, H. Redesign of PCR primers for mitochondrial cytochrome c oxidase subunit I for marine invertebrates and application in all-taxa biotic surveys. Mol. Ecol. Resour. 13, 851–861 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    56.Uit de Weerd, D. R. & Gittenberger, E. Phylogeny of the land snail family Clausiliidae (Gastropoda: Pulmonata). Mol. Phylogenet. Evol. 67, 201–216 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    57.Katoh, K., Rozewicki, J. & Yamada, K. D. MAFFT online service: multiple sequence alignment, interactive sequence choice and visualization. Brief. Bioinform. 20, 1160–1166 (2019).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    58.Nixon, K. C. & Wheeler, Q. D. An amplification of the phylogenetic species concept. Cladistics 6, 211–223 (1990).Article 

    Google Scholar 
    59.Galtier, N. Delineating species in the speciation continuum: A proposal. Evol. Appl. 12, 657–663 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    60.DeSalle, R., Egan, M. G. & Siddall, M. The unholy trinity: Taxonomy, species delimitation and DNA barcoding. Philos. Trans. R. Soc. B Biol. Sci. 360, 1905–1916 (2005).CAS 
    Article 

    Google Scholar 
    61.Bouchet, P. et al. Revised classification, nomenclator and typification of gastropod and monoplacophoran families. Malacologia 61, 1–526 (2017).Article 

    Google Scholar 
    62.Wethington, A. R. & Guralnick, R. Are populations of physids from different hot springs distinctive lineages?. Am. Malacol. Bull. 19, 135–144 (2004).
    Google Scholar 
    63.Huang, Y., Niu, B., Gao, Y., Fu, L. & Li, W. CD-HIT Suite: A web server for clustering and comparing biological sequences. Bioinformatics 26, 680–682 (2010).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    64.Nguyen, L. T., Schmidt, H. A., Von Haeseler, A. & Minh, B. Q. IQ-TREE: A fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Mol. Biol. Evol. 32, 268–274 (2015).CAS 
    Article 

    Google Scholar 
    65.Puillandre, N., Brouillet, S. & Achaz, G. ASAP: Assemble species by automatic partitioning. Mol. Ecol. Resour. 21, 609–620 (2020).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    66.Puillandre, N., Lambert, A., Brouillet, S. & Achaz, G. ABGD, Automatic Barcode Gap Discovery for primary species delimitation. Mol. Ecol. 21, 1864–1877 (2012).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    67.Delicado, D., Arconada, B., Aguado, A. & Ramos, M. A. Multilocus phylogeny, species delimitation and biogeography of Iberian valvatiform springsnails (Caenogastropoda: Hydrobiidae), with the description of a new genus. Zool. J. Linn. Soc. 186, 892–914 (2019).Article 

    Google Scholar 
    68.Kapli, T. et al. Multi-rate Poisson tree processes for single-locus species delimitation under maximum likelihood and Markov chain Monte Carlo. Bioinformatics 33, 1630–1638 (2016).
    Google Scholar 
    69.Clement, M., Posada, D. C. & Crandall, K. A. TCS: A computer program to estimate gene genealogies. Mol. Ecol. 9, 1657–1659 (2000).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    70.Hart, M. W. & Sunday, J. Things fall apart: Biological species form unconnected parsimony networks. Biol. Lett. 3, 509–512 (2007).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    71.Minh, B. Q., Nguyen, M. A. & von Haeseler, A. Ultrafast approximation for phylogenetic bootstrap. Mol. Biol. Evol. 30, 1188–1195 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    72.Kumar, S., Stecher, G. & Tamura, K. MEGA7: Molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 33, 1870–1874 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    73.Meier, R., Zhang, G. & Ali, F. The use of mean instead of smallest interspecific distances exaggerates the size of the “barcoding gap” and leads to misidentification. Syst. Biol. 57, 809–813 (2008).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    74.Dellicour, S. & Flot, J. F. The hitchhiker’s guide to single-locus species delimitation. Mol. Ecol. Resour. 18, 1234–1246 (2018).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    75.Avise, J. C. Phylogeography: The History and Formation of Species (Harvard University Press, 2000).Book 

    Google Scholar 
    76.Dinapoli, A., Tamer, C., Franssen, S., Naduvilezhath, L. & Klussmann-Kolb, A. Utility of H3-gene sequences for phylogenetic reconstruction—a case study of heterobranch Gastropoda. Bonner Zoologische Beiträge 55(3/4), 191–202 (2006).
    Google Scholar 
    77.Ayyagari, V. S. & Sreerama, K. Molecular phylogenetic analysis of Pulmonata (Mollusca: Gastropoda) on the basis of histone-3 gene. Beni-Suef Univ. J. Basic Appl. Sci. 8, 1–8 (2019).Article 

    Google Scholar  More

  • in

    Co-formulant in a commercial fungicide product causes lethal and sub-lethal effects in bumble bees

    Here we show, for the first time, that the toxicity of a pesticide formulation to bees is caused exclusively by a co-formulant (alcohol ethoxylates), rather than the active ingredient. A 0.8 µL acute oral dose of the agricultural fungicide formulation Amistar® caused a range of damage to bees: both lethal, with 23% mortality, and sublethal, with 45% reduced sucrose consumption, 3.8% drop in body weight (whereas the negative control gained 4.8%), and a 302% increase in gut melanisation. For all metrics tested, the Amistar® and alcohol ethoxylates treatments were not statistically different, demonstrating conclusively that the toxicity of the formulation, Amistar®, to bumble bees is driven by the alcohol ethoxylates. These results demonstrate gaps in the regulatory system and highlight the need for a greater research focus on co-formulants.The mortality in the Amistar® treatment, and treatments containing alcohol ethoxylates reached 32% at its highest, which is substantial given that bees are likely to have a high level of exposure to Amistar® and alcohol ethoxylates. The mechanism by which the alcohol ethoxylates cause mortality has not been explicitly isolated, but our results suggest two potential, possibly related, causes. We recorded a 302% increase in the melanised area of bee midguts in the alcohol ethoxylates treatment. A similar effect was observed in Melipona scutellaris exposed to the pure fungicide active ingredient pyraclostrobin alongside a similar reduction in survival37. We suggest that the alcohol ethoxylates are disrupting the structure of the midgut, which the bee immune system is reacting to with melanisation44 (see Fig. 5). In parallel with this gut damage, alcohol ethoxylate treatment drove a 54% reduction in sugar consumption, which persisted throughout the experiment. Supplementary Fig. S3 shows a plot comparing sugar consumption against gut melanisation, with increasing gut melanisation correlated to reduced sugar consumption in the Amistar®, co-formulant mixture and alcohol ethoxylates treatments. Consequently, we propose that mortality was driven by energy depletion due to reduced consumption, which in turn may have been driven by damage to the gut.Figure 5(Left) Bumble bee midgut in the negative control treatment. (Right) Bumble bee midgut in the co-formulant mixture treatment, which contains alcohol ethoxylates. The dark brown patches are areas of melanisation, indicative of damage to the gut. Both bees survived the full 120 h.Full size imageLikely as a consequence of the reduced consumption of sucrose, bumble bees in the alcohol ethoxylates treatment lost 8.4% of their original weight, in stark contrast to the negative control where bees gained 4.8% over the five-day period. This indicates the alcohol ethoxylate treated bees were expending more energy than they were consuming, and thus exhibiting a negative energy balance. This weight loss, while considerable as a percentage of the bee’s total body mass, is also similar in scale to the weight of the sucrose bees consume in one sitting (EA Straw pers. obs.), for which rigorous data do not exist. As such it is possible that a portion of the weight loss is attributable to the reduced sucrose consumption of the bees, meaning they would have less sucrose in their guts at the time of weighing. Sucrose consumption does not, however, explain the failure of alcohol ethoxylate treated bees to gain weight, which was observed in the control treatment. The weight loss, and lack of weight gain, are concerning because they are likely to indicate a reduction in fat reserves, although this has not been experimentally confirmed. Bee fat reserves are important physiologically, in particular in responding to immune threats45,46. Fat reserves allow bees the energetic resources to buffer against challenges, and thus their depletion could expose bees to greater risk from future threats47.The reduced appetite and negative energy balance in alcohol ethoxylates treated bees could have broader effects in the natural environment. Bees pollinate flowers as they forage for nectar and pollen, so a reduction in their appetite could subsequently have effects on ecosystem services. In our experiment, bumble bee appetite was reduced immediately after ingesting a single dose of alcohol ethoxylates or Amistar®. This effect persisted for five days after exposure, indicating a persistent change in consumption behaviour. While nectar-foraging in bumble bees is driven by the needs of the colony48, a reduction in appetite would reduce overall colony nectar consumption, and thus the number of foraging trips made for nectar. Fewer visits to flowers for nectar may lead to reduced pollination, which would be detrimental to crop yields and farm profits. Further studies of how the impacts we have found map onto foraging and pollination are clearly needed. Importantly, the reduction in appetite recorded in our experiment is a sublethal effect, which standard lower tier testing would not detect. When Amistar® is tested on bumble bees for the 2025 renewal of azoxystrobin, this sublethal effect will be missed by regulatory testing, despite the impact it may have on the pollination services such testing is designed to protect. We suggest that a simple modification to the regulatory protocol OECD 247 would be to weigh the sucrose syringes at the start and end of the trials to calculate sucrose consumption, which would allow measurement of this sublethal effect with minimal additional workload.Our results show a slightly, but not significantly, higher level of mortality in the alcohol ethoxylates treatment (30%) than the Amistar® treatment (23%). If this is a real biological difference, one explanation might be that the concentration of alcohol ethoxylates in the Amistar® formulation was lower than that used in the alcohol ethoxylates treatment solution. This is possible because the Amistar® material safety data sheet lists concentrations as a range (10–20% for alcohol ethoxylates), and here we used the upper end of the range. The co-formulant mixture treatment in all metrics was statistically indistinguishable from the alcohol ethoxylates treatment, showing that the toxicity of alcohol ethoxylates is not a result of synergism with other co-formulants.We believe that the implications of our results are not limited to a laboratory setting and a single species, as other published and unpublished research supports our findings. Semi-field flight cage experiments, where Amistar® was applied to a crop, found effects on full bumble bee colonies (Bombus terrestris). Amistar® caused a reduction in average bee weight and a reduction in foraging activity, as our results predict49. This demonstrates that the effects observed in our laboratory testing scale up to effects at a field realistic level. Additionally, in honeybees (Apis mellifera) Amistar® has been found to cause mortality in laboratory experiments at a range of doses50,51, demonstrating the mortality effect found in our experiment is not species specific. However, no mortality was seen in trials on the red mason bee Osmia bicornis (Hellström and Paxton, unpublished data). Additionally, a similar compound, C11 and lower alcohol ethoxylates, has been found in small scale laboratory testing to cause 100% mortality after contact exposure in honeybees31.To measure the exposure of bees to PPP’s, the EU mandates trials that measure chemical residues in pollen and nectar after crops have been sprayed with either active ingredients or formulations34. However, these residue analysis studies only measure active ingredient concentrations, not the co-formulants. As such, we have no systematic data on the exposure of bees to co-formulants7,8,9. This dearth of data means that the exposure of bees to co-formulants is very poorly characterised. To estimate exposure to alcohol ethoxylates, residue data for Amistar®’s active ingredient azoxystrobin could be used as a proxy18,52. However, the chemical properties of alcohol ethoxylates, specifically their surfactant action, make it unlikely that they have an equivalent environmental fate to azoxystrobin, so this would not be appropriate.While we have very little data to quantify bee exposure to alcohol ethoxylates, we know Amistar® can be applied to crops, such as strawberries, during flowering while bees are foraging on them. The Environmental Information Sheet for Amistar® states “[For bees] no risk management is necessary. Amistar® is of low risk to honey bees”53,54,55. In addition, we would note that exposure of bees to alcohol ethoxylates, and related substances, is not exclusively from Amistar®. For example, a cursory search of the Syngenta website56 immediately identified alcohol ethoxylates in five other Syngenta products. Worryingly, the chemical group alcohol ethoxylates sit in, alkoxylated alcohols, are also widely used in adjuvants, which are products which can be added to tank mixtures to modify the action of the agrichemical6. 89 adjuvant products licenced in the UK contain alkoxylated alcohols as the primary ingredient15. To our knowledge, these adjuvants have never been toxicity tested on bees and have no bee exposure mitigation measures in place whatsoever.To complement measures to promote academic research, moving regulatory research beyond its mortality and active ingredient-centric approach to toxicity testing would better reflect the risks pesticides, as used in the field, pose. For regulatory systems to accurately characterise risk they need to estimate the scale of sublethal effects, regardless of initial mortality results33. The results presented here demonstrate that even substances assessed by regulators as ‘bee safe’ can pose a serious hazard to bee health. To reflect potential sublethal differences caused by co-formulation composition, all formulations could undergo a much more rigorous set of lower tier testing or be automatically entered for higher tier testing.In the face of declining bee populations we advocate that a precautionary approach minimising the exposure of bees to potential stressors, where possible, would be prudent. The current legislation allowing application of PPPs directly onto bees and flowering plants does not align with the emerging evidence that co-formulants, adjuvants, herbicides and fungicides can be hazardous to bees8,57. The wealth of untested and undisclosed co-formulants used abundantly in agriculture is a serious and pressing concern for the health of pollinators worldwide. More

  • in

    The rumen microbiome inhibits methane formation through dietary choline supplementation

    1.Cubasch, U. et al. Climate Change 2013: the physical science basis. In Contribution of Working Group I to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change, Vol 1 (eds Stocker, T. F. et al.) 119–158 (Cambridge University Press, 2013).
    Google Scholar 
    2.Jackson, R. B. et al. Increasing anthropogenic methane emissions arise equally from agricultural and fossil fuel sources. Environ. Res. Lett. https://doi.org/10.1088/1748-9326/ab9ed2 (2020).Article 

    Google Scholar 
    3.Saunois, M. et al. The global methane budget 2000–2017. Earth Syst. Sci. Data 12, 1561–1623. https://doi.org/10.5194/essd-12-1561-2020 (2020).ADS 
    Article 

    Google Scholar 
    4.Hobson, P. N. & Stewart, C. S. The rumen Microbial Ecosystem (Blackie Academic & Professional, 1997).Book 

    Google Scholar 
    5.Henderson, G. et al. Rumen microbial community composition varies with diet and host, but a core microbiome is found across a wide geographical range. Sci. Rep. 5, 14567. https://doi.org/10.1038/srep14567 (2015).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    6.Li, Y. et al. The complete genome sequence of the methanogenic archaeon ISO4-H5 provides insights into the methylotrophic lifestyle of a ruminal representative of the Methanomassiliicoccales. Stand. Genom. Sci. 11, 59. https://doi.org/10.1186/s40793-016-0183-5 (2016).Article 

    Google Scholar 
    7.Lang, K. et al. New mode of energy metabolism in the seventh order of methanogens as revealed by comparative genome analysis of “Candidatus Methanoplasma termitum”. Appl. Environ. Microbiol. 81, 1338–1352. https://doi.org/10.1128/AEM.03389-14 (2015).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    8.Hoehler, T., Losey, N. A., Gunsalus, R. P. & McInerney, M. J. In Biogenesis of Hydrocarbons (eds Stams, A. & Sousa, D.) 1–26 (Springer, 2018).
    Google Scholar 
    9.Neill, A. R., Grime, D. W. & Dawson, R. M. C. Conversion of choline methyl groups through trimethylamine into methane in the rumen. Biochem. J. 170, 529–535. https://doi.org/10.1042/bj1700529 (1978).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    10.Erdman, R. A. & Sharma, B. K. Effect of dietary rumen-protected choline in lactating dairy cows. J. Dairy Sci. 74, 1641–1647. https://doi.org/10.3168/jds.S0022-0302(91)78326-4 (1991).CAS 
    Article 
    PubMed 

    Google Scholar 
    11.Sharma, B. K. & Erdman, R. A. Effects of dietary and abomasally infused choline on milk production responses of lactating dairy cows. J. Nutr. 119, 248–254. https://doi.org/10.1093/jn/119.2.248 (1989).CAS 
    Article 
    PubMed 

    Google Scholar 
    12.Soliva, C. & Hess, H. In Measuring Methane Production from Ruminants: Measuring Methane Emission of Ruminants by In Vitro and In Vivo Techniques (eds Makkar, H. P. & Vercoe, P. E.) 15–31 (Springer, 2007).Chapter 

    Google Scholar 
    13.Craciun, S. & Balskus, E. P. Microbial conversion of choline to trimethylamine requires a glycyl radical enzyme. PNAS 109, 21307–21312. https://doi.org/10.1073/pnas.1215689109 (2012).ADS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    14.Jameson, E. et al. Anaerobic choline metabolism in microcompartments promotes growth and swarming of Proteus mirabilis. Environ. Microbiol. 18, 2886–2898. https://doi.org/10.1111/1462-2920.13059 (2016).CAS 
    Article 
    PubMed 

    Google Scholar 
    15.Herring, T. I., Harris, T. N., Chowdhury, C., Mohanty, S. K. & Bobik, T. A. A bacterial microcompartment is used for choline fermentation by Escherichia coli 536. J. Bacteriol. 200, e00764-e817. https://doi.org/10.1128/JB.00764-17 (2018).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    16.EFSA. Scientific Opinion on safety and efficacy of choline chloride as a feed additive for all animal species. EFSA J. 9, 2353 (2011).
    Google Scholar 
    17.Lewis, D. J. Ammonia toxicity in the ruminant. J. Agric. Sci. 55, 111–117 (1960).CAS 
    Article 

    Google Scholar 
    18.Hogan, J. P. Absorption of ammonia through rumen of sheep. Aust. J. Biol. Sci. 14, 448–450. https://doi.org/10.1071/Bi9610448 (1961).CAS 
    Article 

    Google Scholar 
    19.Sprott, G. D. & Patel, G. B. Ammonia toxicity in pure cultures of methanogenic bacteria. Syst. Appl. Microbiol. 7, 358–363 (1986).CAS 
    Article 

    Google Scholar 
    20.Lewis, D. Ammonia toxicity in the ruminant. J. Agric. Sci. 55(1), 111–117 (1960).CAS 
    Article 

    Google Scholar 
    21.Ungerfeld, E. M., Rust, S. R. & Burnett, R. Increases in microbial nitrogen production and efficiency in vitro with three inhibitors of ruminal methanogenesis. Can. J. Microbiol. 53, 496–503. https://doi.org/10.1139/W07-008 (2007).CAS 
    Article 
    PubMed 

    Google Scholar 
    22.Lundgren, B. R., Sarwar, Z., Pinto, A., Ganley, J. G. & Nomura, C. T. Ethanolamine catabolism in Pseudomonas aeruginosa PAO1 is regulated by the enhancer-binding protein EatR (PA4021) and the alternative sigma factor RpoN. J. Bacteriol. 198, 2318–2329. https://doi.org/10.1128/JB.00357-16 (2016).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    23.Rychlik, J. L., LaVera, R. & Russell, J. B. Amino acid deamination by ruminal Megasphaera elsdenii strains. Curr. Microbiol. 45, 340–345. https://doi.org/10.1007/s00284-002-3743-4 (2002).CAS 
    Article 
    PubMed 

    Google Scholar 
    24.Park, K. & Lee, H. Effects of nitrogen gas flushing in comparison with argon on rumen fermentation characteristics in in vitro studies. J. Anim. Sci. Technol. 62, 52–57. https://doi.org/10.5187/jast.2020.62.1.52 (2020).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    25.Hobson, P. N., Summers, R., Postgate, J. R. & Ware, D. A. Nitrogen fixation in the rumen of a living sheep. J. Gen. Microbiol. 77, 225–226. https://doi.org/10.1099/00221287-77-1-225 (1973).CAS 
    Article 
    PubMed 

    Google Scholar 
    26.Harada, N., Nishiyama, M. & Matsumoto, S. Inhibition of methanogens increases photo-dependent nitrogenase activities in anoxic paddy soil amended with rice straw. FEMS Microbiol. Ecol. 35, 231–238. https://doi.org/10.1111/j.1574-6941.2001.tb00808.x (2001).CAS 
    Article 
    PubMed 

    Google Scholar 
    27.Haaker, H. & Klugkist, J. The bioenergetics of electron transport to nitrogenase. J FEMS Microbiol. Lett. 46, 57–71 (1987).CAS 
    Article 

    Google Scholar 
    28.Edgren, T. & Nordlund, S. The fixABCX genes in Rhodospirillum rubrum encode a putative membrane complex participating in electron transfer to nitrogenase. J. Bacteriol. 186, 2052–2060 (2004).CAS 
    Article 

    Google Scholar 
    29.Igai, K. et al. Nitrogen fixation and nifH diversity in human gut microbiota. Sci. Rep. 6, 31942. https://doi.org/10.1038/srep31942 (2016).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    30.Ungerfeld, E. M. Shifts in metabolic hydrogen sinks in the methanogenesis-inhibited ruminal fermentation: A meta-analysis. Front. Microbiol. 6, 37. https://doi.org/10.3389/fmicb.2015.00037 (2015).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    31.Leahy, S. C. et al. The complete genome sequence of Methanobrevibacter sp. AbM4. Stand. Genom. Sci. 8, 215–227. https://doi.org/10.4056/sigs.3977691 (2013).CAS 
    Article 

    Google Scholar 
    32.Hoedt, E. C. et al. Differences down-under: Alcohol-fueled methanogenesis by archaea present in Australian macropodids. ISME J. 10, 2376–2388. https://doi.org/10.1038/ismej.2016.41 (2016).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    33.Ungerfeld, E. M. & Kohn, R. A. In Ruminant Physiology: Digestion, Metabolism and Impact of Nutrition on Gene Expression, Immunology and Stress (eds Sejrsen, K. et al.) 55–85 (Wageningen Academic Publishers, 2006).
    Google Scholar 
    34.van Zijderveld, S. M. et al. Nitrate and sulfate: Effective alternative hydrogen sinks for mitigation of ruminal methane production in sheep. J. Dairy Sci. 93, 5856–5866. https://doi.org/10.3168/jds.2010-3281 (2010).CAS 
    Article 
    PubMed 

    Google Scholar 
    35.Lan, W. & Yang, C. Ruminal methane production: Associated microorganisms and the potential of applying hydrogen-utilizing bacteria for mitigation. Sci. Total Environ. 654, 1270–1283. https://doi.org/10.1016/j.scitotenv.2018.11.180 (2019).ADS 
    CAS 
    Article 
    PubMed 

    Google Scholar 
    36.Loubinoux, J., Bronowicki, J. P., Pereira, I. A., Mougenel, J. L. & Faou, A. E. Sulfate-reducing bacteria in human feces and their association with inflammatory bowel diseases. FEMS Microbiol. Ecol. 40, 107–112. https://doi.org/10.1111/j.1574-6941.2002.tb00942.x (2002).CAS 
    Article 
    PubMed 

    Google Scholar 
    37.Gould, D. H., Cummings, B. A. & Hamar, D. W. In vivo indicators of pathologic ruminal sulphide production in steers with diet-induced polioencephalomalacia. J. Vet. Diagn. Invest. 9, 72–76. https://doi.org/10.1177/104063879700900113 (1997).CAS 
    Article 
    PubMed 

    Google Scholar 
    38.Anderson, R. C., Rasmussen, M. A., Jensen, N. S. & Allison, M. J. Denitrobacterium detoxificans gen. nov., sp. nov., a ruminal bacterium that respires on nitrocompounds. Int. J. Syst. Evol. Microbiol. 50(Pt 2), 633–638. https://doi.org/10.1099/00207713-50-2-633 (2000).CAS 
    Article 
    PubMed 

    Google Scholar 
    39.Anderson, R. C. et al. Ruminal fermentation of anti-methanogenic nitrate- and nitro-containing forages in vitro. Front. Vet. Sci. 3, 62. https://doi.org/10.3389/fvets.2016.00062 (2016).ADS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    40.Zhang, Z. W. et al. Nitrocompounds as potential methanogenic inhibitors in ruminant animals: A review. Anim. Feed Sci. Tech. 236, 107–114. https://doi.org/10.1016/j.anifeedsci.2017.12.010 (2018).CAS 
    Article 

    Google Scholar 
    41.Marounek, M., Fliegrova, K. & Bartos, S. Metabolism and some characteristics of ruminal strains of Megasphaera elsdenii. Appl. Environ. Microbiol. 55, 1570–1573. https://doi.org/10.1128/AEM.55.6.1570-1573.1989 (1989).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    42.Hackmann, T. J., Ngugi, D. K., Firkins, J. L. & Tao, J. Genomes of rumen bacteria encode atypical pathways for fermenting hexoses to short-chain fatty acids. Environ. Microbiol. 19, 4670–4683. https://doi.org/10.1111/1462-2920.13929 (2017).CAS 
    Article 
    PubMed 

    Google Scholar 
    43.Janssen, P. H. Influence of hydrogen on rumen methane formation and fermentation balances through microbial growth kinetics and fermentation thermodynamics. Anim. Feed Sci. Technol. 160, 1–22. https://doi.org/10.1016/j.anifeedsci.2010.07.002 (2010).CAS 
    Article 

    Google Scholar 
    44.Greening, C. et al. Diverse hydrogen production and consumption pathways influence methane production in ruminants. ISME J. 13, 2617–2632. https://doi.org/10.1038/s41396-019-0464-2 (2019).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    45.Gilmour, M., Flint, H. J. & Mitchell, W. J. Multiple lactate dehydrogenase activities of the rumen bacterium Selenomonas ruminantium. Microbiol. 140(Pt 8), 2077–2084. https://doi.org/10.1099/13500872-140-8-2077 (1994).CAS 
    Article 

    Google Scholar 
    46.Chowdhury, N. P., Kahnt, J. & Buckel, W. Reduction of ferredoxin or oxygen by flavin-based electron bifurcation in Megasphaera elsdenii. FEBS J. 282, 3149–3160. https://doi.org/10.1111/febs.13308 (2015).CAS 
    Article 
    PubMed 

    Google Scholar 
    47.Weghoff, M. C., Bertsch, J. & Muller, V. A novel mode of lactate metabolism in strictly anaerobic bacteria. Environ. Microbiol. 17, 670–677. https://doi.org/10.1111/1462-2920.12493 (2015).CAS 
    Article 
    PubMed 

    Google Scholar 
    48.Hernandez, J., Benedito, J. L., Abuelo, A. & Castillo, C. Ruminal acidosis in feedlot: from aetiology to prevention. Sci. World J. 2014, 702572. https://doi.org/10.1155/2014/702572 (2014).Article 

    Google Scholar 
    49.Vuotto, C., Barbanti, F., Mastrantonio, P. & Donelli, G. Lactobacillus brevis CD2 inhibits Prevotella melaninogenica biofilm. Oral Dis. 20, 668–674. https://doi.org/10.1111/odi.12186 (2014).CAS 
    Article 
    PubMed 

    Google Scholar 
    50.van Lingen, H. J. et al. Thermodynamic driving force of hydrogen on rumen microbial metabolism: A theoretical investigation. PLoS One 11, e0161362. https://doi.org/10.1371/journal.pone.0161362 (2016).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    51.Ungerfeld, E. M., Aedo, M. F., Martinez, E. D. & Saldivia, M. Inhibiting methanogenesis in rumen batch cultures did not increase the recovery of metabolic hydrogen in microbial amino acids. Microorganisms 7, 155. https://doi.org/10.3390/microorganisms7050115 (2019).CAS 
    Article 

    Google Scholar 
    52.Ng, F. et al. An adhesin from hydrogen-utilizing rumen methanogen Methanobrevibacter ruminantium M1 binds a broad range of hydrogen-producing microorganisms. Environ. Microbiol. 18, 3010–3021. https://doi.org/10.1111/1462-2920.13155 (2016).CAS 
    Article 
    PubMed 

    Google Scholar 
    53.Soliva, C. R., Amelchanka, S. L., Duval, S. M. & Kreuzer, M. Ruminal methane inhibition potential of various pure compounds in comparison with garlic oil as determined with a rumen simulation technique (Rusitec). Brit. J. Nutr. 106, 114–122. https://doi.org/10.1017/S0007114510005684 (2011).CAS 
    Article 
    PubMed 

    Google Scholar 
    54.Terranova, M. et al. Increasing the proportion of hazel leaves in the diet of dairy cows reduced methane yield and excretion of nitrogen in volatile form, but not milk yield. Anim. Feed Sci. Technol. 276, 114796. https://doi.org/10.1016/j.anifeedsci.2020.114790 (2021).CAS 
    Article 

    Google Scholar 
    55.Ehrlich, G. G., Goerlitz, D. F., Bourell, J. H., Eisen, G. V. & Godsy, E. M. Liquid chromatographic procedure for fermentation product analysis in the identification of anaerobic bacteria. Appl. Environ. Microbiol. 42, 878–885 (1981).ADS 
    CAS 
    Article 

    Google Scholar 
    56.Bica, R. et al. Nuclear magnetic resonance to detect rumen metabolites associated with enteric methane emissions from beef cattle. Sci. Rep. 10, 5578. https://doi.org/10.1038/s41598-020-62485-y (2020).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    57.Henderson, G. et al. Effect of DNA extraction methods and sampling techniques on the apparent structure of cow and sheep rumen microbial communities. PLoS One 8, e74787. https://doi.org/10.1371/journal.pone.0074787 (2013).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    58.Kittelmann, S. et al. Simultaneous amplicon sequencing to explore co-occurrence patterns of bacterial, archaeal and eukaryotic microorganisms in rumen microbial communities. PLoS One 8, e47879. https://doi.org/10.1371/journal.pone.0047879 (2013).ADS 
    CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    59.Milanese, A. et al. Microbial abundance, activity and population genomic profiling with mOTUs2. Nat. Comm. 10, 1014. https://doi.org/10.1038/s41467-019-08844-4 (2019).ADS 
    CAS 
    Article 

    Google Scholar 
    60.Paoli, L. et al. Uncharted biosynthetic potential of the ocean microbiome. bioRxiv https://doi.org/10.1101/2021.03.24.436479 (2021).Article 

    Google Scholar 
    61.Bushnell, B. BBMap: A fast, accurate, splice-aware aligner. in 9th Annual Genomics of Energy & Environment Meeting. (Lawrence Berkeley National Lab (LBNL), Berkeley, CA, USA). https://www.osti.gov/servlets/purl/1241166 (2014).62.Nurk, S., Meleshko, D., Korobeynikov, A. & Pevzner, P. A. metaSPAdes: A new versatile metagenomic assembler. Genome Res. 27, 824–834. https://doi.org/10.1101/gr.213959.116 (2017).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    63.Li, H. & Durbin, R. Fast and accurate short read alignment with Burrows–Wheeler transform. Bioinformatics 25, 1754–1760. https://doi.org/10.1093/bioinformatics/btp324 (2009).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    64.Kang, D. D. et al. MetaBAT 2: An adaptive binning algorithm for robust and efficient genome reconstruction from metagenome assemblies. PeerJ 7, e7359. https://doi.org/10.7717/peerj.7359 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    65.Parks, D. H., Imelfort, M., Skennerton, C. T., Hugenholtz, P. & Tyson, G. W. CheckM: Assessing the quality of microbial genomes recovered from isolates, single cells, and metagenomes. Genome Res. 25, 1043–1055. https://doi.org/10.1101/gr.186072.114 (2015).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    66.Seshadri, R. et al. Cultivation and sequencing of rumen microbiome members from the Hungate1000 Collection. Nat. Biotechnol. 36, 359–367. https://doi.org/10.1038/nbt.4110 (2018).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    67.Stewart, R. D. et al. Compendium of 4,941 rumen metagenome-assembled genomes for rumen microbiome biology and enzyme discovery. Nat. Biotechnol. 37, 953–961. https://doi.org/10.1038/s41587-019-0202-3 (2019).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    68.Hyatt, D. et al. Prodigal: Prokaryotic gene recognition and translation initiation site identification. BMC Bioinform. 11, 119. https://doi.org/10.1186/1471-2105-11-119 (2010).CAS 
    Article 

    Google Scholar 
    69.Fu, L., Niu, B., Zhu, Z., Wu, S. & Li, W. CD-HIT: Accelerated for clustering the next-generation sequencing data. Bioinformatics 28, 3150–3152. https://doi.org/10.1093/bioinformatics/bts565 (2012).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    70.Kanehisa, M. & Goto, S. KEGG: Kyoto encyclopedia of genes and genomes. Nucleic Acid Res. 28, 27–30. https://doi.org/10.1093/nar/28.1.27 (2000).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    71.Buchfink, B., Xie, C. & Huson, D. H. Fast and sensitive protein alignment using DIAMOND. Nat. Methods 12, 59–60. https://doi.org/10.1038/nmeth.3176 (2015).CAS 
    Article 
    PubMed 

    Google Scholar 
    72.Li, L., Stoeckert, C. J. Jr. & Roos, D. S. OrthoMCL: Identification of ortholog groups for eukaryotic genomes. Genome Res. 13, 2178–2189. https://doi.org/10.1101/gr.1224503 (2003).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    73.Allen, B., Drake, M., Harris, N. & Sullivan, T. Using KBase to assemble and annotate prokaryotic genomes. Curr. Protoc. Microbiol. 46, 1E 13 11-11E 13 18. https://doi.org/10.1002/cpmc.37 (2017).Article 

    Google Scholar 
    74.RStudio Team. R Studio: Integrated development environment for R. Version 1.4.1106 (2021).75.Oksanen, J. et al. The vegan package. Community Ecol. Pack. 10, 631–637 (2007).
    Google Scholar 
    76.Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550. https://doi.org/10.1186/s13059-014-0550-8 (2014).CAS 
    Article 
    PubMed 
    PubMed Central 

    Google Scholar  More

  • in

    Insecticide resistance by a host-symbiont reciprocal detoxification

    Insects and bacteriaBean bugs were reared in petri dishes (90 mm in diameter and 20-mm high) at 25 °C under a long-day regimen (16-h light, 8-h dark) and fed with soybean seeds and distilled water containing 0.05% ascorbic acid (DWA). Burkholderia symbiont strain SFA119, a MEP-degrading strain conferring MEP resistant in the bean bug, and its GFP-(green fluorescent protein) labeled derivative, strain SJ586, were used in this study. The symbiont was cultured at 30 °C on YG medium (0.5% yeast extract, 0.4% glucose, and 0.1% NaCl). The GFP-labeled strain was constructed by the Tn7 mini-transposon system, as previously described31.Genome sequencingDNA was extracted from cultured cells of strain SFA1 by the phenol–chloroform extraction as previously described32. The DNA library for Illumina short reads (the mean insert size: 500 bp) was constructed by using the Covaris S2 ultrasonicator (Covaris) and the KAPA HyperPrep Kit (Kapa Biosystems). For the library construction for Nanopore long reads, Native Barcoding Expansion (EXP-NBD104, Oxford Nanopore Technologies) and the Ligation Sequencing Kit (SQK-LSK109, Oxford Nanopore Technologies) were used. The genome sequencing was performed with NextSeq using the 2 × 151-bp protocol (Illumina) and GridION using an R9.4.1 flow cell (Oxford Nanopore Technologies). The Illumina short reads were processed by using Sickle Ver 1.33 (available at https://github.com/najoshi/sickle) for removing the low-quality and shorter reads. After processing the Nanopore long-reads with Porechop Ver 0.2.3 (available at https://github.com/rrwick/Porechop) and Filtlong Ver 0.2.0 (available at https://github.com/rrwick/Filtlong), error correction was performed by using Canu Ver 1.833. These processed short- and long reads were assembled by using Unicycler Ver 0.4.734, resulting in the eight circular replicons (Supplementary Fig. 1). The assembled genome was annotated by DFAST Ver 1.1.035. After the homology searches of the protein sequences by blastp 2.5.0 + 36 against the COG database (PMID: 25428365), circular replicons were visualized with circos v 0.69-837. The chromosomes and plasmids were assigned according to the genome of Caballeronia (Burkholderia) cordobensis strain YI2338.Phylogenetic analysisNucleotide sequences of 16 S rRNA gene of representative Burkholderia spp. and outgroup species were aligned by using SINA v1.2.1139. Protein sequences of MEP-degrading genes (mpd, pnpB, and mhqA) and a plasmid-transfer gene (traH) on plasmid 2 were subjected to the blastp search against the nr database (downloaded in Jul. 2019) and top ~30 hit sequences were retrieved for each gene. Multiple sequencing alignments of each gene were constructed with L-INS-I of mafft v7.40740. Gap-including and ambiguous sites in the alignments were then removed. Unrooted maximum-likelihood (ML) phylogenetic trees were reconstructed with RAxML v8.2.341 using the GTR + Γ model (for 16 S rRNA gene) or the LG + Γ model42 (for other genes). The bootstrap values of 1000 replicates for all internal branches were calculated with a rapid bootstrapping algorithm43.Preparation of SFA1 cultures for RNA-seqBurkholderia symbiont SFA1 was precultured in minimal medium (20 mM phosphate buffer [pH 7.0], 0.01% yeast, 0.1% (NH4)2SO4, 0.02% NaCl, 0.01% MgSO4⋅7H2O, 0.005% CaCl2⋅2H2O, 0.00025% FeSO4⋅7H2O, and 0.00033% EDTA⋅2Na) containing 1.0 mM of MEP on a gyratory shaker (210 rpm) at 30 °C overnight, and subcultured in newly prepared MEP-containing minimal medium under the same conditions for 5 h. As a control, SFA1 was precultured in minimal medium containing 0.1% citrate overnight, and then the overnighter was subcultured in a newly prepared citrate-containing minimal medium under the same conditions for 10 h. The culture was mixed with an equal amount of RNAprotect Bacteria Regent (Qiagen, Valencia, CA, USA), then centrifuged to harvest the cells for the RNA-seq analysis.Preparation of midgut symbiont cells for RNA-seqThe oral administration of the symbiont strain SFA1 was performed as described19,44. The symbiont was inoculated to 2nd instar nymphs, and three days after molting to the 3rd instar, nymphs were transdermally administered with 1 µl of 0.2 µM or 20 µM of MEP (dissolved in acetone). One- or three days after the treatment, insects were dissected and the crypt-bearing symbiotic gut region was subjected to the RNA extraction and RNA-seq analysis. As a control, untreated insects were analyzed.RNA-seq analysisTotal RNA was extracted from triplicate samples from cultures by the hot-phenol method as previously described45 or from the midgut symbiont cells by using RNAiso Plus (Takara Bi, Kusatsu, Shiga, Japan) and the RNeasy mini kit (Qiagen). The extracted total RNA was purified by phenol–chloroform extraction and digestion by DNase (RQ1 RNase-Free DNase, Promega, Fitchburg, WI, USA) and repurified by using a RNeasy Mini Kit. The mRNA in the samples was further enriched by the RiboMinus Transcriptome Isolation Kit bacteria (Thermo Fisher Scientific, Waltham, MA, USA) and the RiboMinus Eukaryote Kit for RNA-Seq (Thermo Fisher Scientific), and purified by using an AMPure XP kit (Beckman Coulter, Brea, CA, USA). The cDNA libraries were constructed from approximately 100 ng of rRNA-depleted RNA samples by the use of a NextUltraRNA library prep kit (New England Biolabs, Ipswich, MA, USA). Size selection of cDNA (200–300 bp) and determination of the size distribution and concentration of the purified cDNA samples were performed as described previously46. In total, 21 cDNA libraries were constructed and sequenced by MiSeq (Illumina, Inc., San Diego, CA, USA). To ensure high sequence quality, the remaining sequencing adapters and the reads with a cutoff Phred score of 15 (for leading and tailing sequences, Phred score of >20) and a length of less than 80 bp in the obtained RNA-seq data were removed by the program Trimmomatic v0.30 using Illumina TruSeq3 adapter sequences for the clipping47. The remaining paired reads were analyzed by FastQC version 0.11.9 (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/) for quality control, and Bowtie2 ver. 2.2.248 for mapping on the symbiont genome (DDBJ/EMBL/GenBank accession: AP022305–AP022312). After the conversion of the output BAM files to BED files using the bamtobed program in BEDTools ver. 2.14.349, gene expression levels were calculated in TPM (transcripts per kilobase million) values by using in-house scripts46.Gene deletion and complementationMEP-degrading genes (mpd, pnpA1, and pnpA2) were deleted by the homologous-recombination-based deletion method using pK18mobsacB or pUC18, as previously described50,51. Primers used for the mutagenesis are listed in Supplementary Table 1. For mpd gene deletion, pK18mobsacB was used to construct a markerless mutant. For single deletion of pnpA1 and pnpA2 genes, pUC18 was used to substitute each gene locus with a kanamycin-resistance gene cassette. The double deletion of pnpA1 and pnpA2 genes was performed by substituting pnpA2 gene locus with a tetracycline-resistance gene cassette in the pnpA1-deletion mutant. Gene complementation of mpd was also performed by homologous recombination using plasmid pUC18 with primers listed in Supplementary Table 1. To investigate growth profiles of the wild-type SFA1, the gene-deletion mutants (Δmpd, ΔpnpA1, ΔpnpA2, and ΔpnpA1/ΔpnpA2), and the mph-complement mutant (Δmpd/mpd+) in the MEP-containing minimal medium, the strains were precultured in minimal medium containing 1.0 mM MEP on a gyratory shaker (210 rpm) at 30 °C overnight, and then cultured in newly prepared MEP-containing minimal medium under the same condition. The growth of cultures was estimated by OD600 measurements. To confirm the basic growth abilities of the mutants, these bacterial strains were pre- and subcultured in minimal medium containing 0.1% glucose under the same conditions. These symbiont strains and mutants were inoculated to the bean bug as described above.Quantitative PCRSymbiont titers in the midgut crypts were evaluated by quantitative PCR (qPCR) of bacterial dnaA gene copies. The qPCR was performed by using a KAPA SYBR Fast qPCR Master Mix (Kapa Biosystems) and the LightCycler 96 System (Roche Applied Science) with the following primers: BSdnaA–F (5′-AGC GCG AGA TCA GAC GGT CGT CGA T-3′) and BSdnaA–R (5′-TCC GGC AAG TCG CGC ACG CA-3′).MEP treatment of insectsMEP treatment of R. pedestris was performed as previously described19. Soybean seeds were dipped in 0.2 mM MEP for 5 s and dried at room temperature. In each clean plastic container, 15 individuals of 3rd-instar nymphs were reared on three seeds of the MEP-treated soybean and DWA at 25 °C under the long-day regime, and the number of dead insects was counted 24 h after the treatments. The survival rate of the insects was analyzed under Fisher’s exact test by use of the program R ver. 3.6.3 (available at https://www.R-project.org/). Multiple comparisons were corrected by the Bonferroni method.Bactericidal activities of MEP and its degradation product 3M4NTo measure bactericidal activities of MEP and 3M4N on cultured cells of SFA1, 104 cells of log-phase growing bacteria were mixed with a defined concentration of MEP or 3M4N, and spotted on a YG agar plate. To measure the bactericidal activity against midgut crypt-colonizing cells, the symbiotic organs infected with SFA1 were dissected from 3rd-instar insects, homogenized in PBS, and purified by a 5-µm-size pore Syringe filter to harvest colonizing symbiont cells50. MEP or 3M4N was added to approximately 104 cells of the harvested cells and spotted on a YG agar plate. Bactericidal activities of the chemical compounds were then checked in 24 h after incubation at 30 °C.HPLC detection of in vitro and in vivo MEP-degrading activities of the symbiontTo determine in vitro MEP-degradation activity, cultured cells of SFA1 were prepared as above, and 106 cells were incubated at 25 °C in 200 µl of MEP solution (2 mM MEP in Tris-Hcl [pH 8.5] with 0.1% Triton X-100) in a 1.5-ml microtube. To determine in vivo MEP-degradation activity, the midgut of a 5th-instar insect infected with SFA1 was dissected, the posterior and anterior parts of the crypt-bearing symbiotic region were closed with 0.2-mm polyethylene fishline (Supplementary Fig. 6a), and incubated at 25 °C in 200 µl of the MEP solution. For the in vivo determination, 250 mM of trehalose, known as a major sugar source of insects’ hemolymph52, was added to the MEP solution to keep the tissue fresh. After incubation for different times, the reaction was stopped by adding 400 µl of methanol. After centrifugation, supernatants were subjected to high-performance liquid chromatography (HPLC) analyses to detect MEP and 3M4N, as previously reported21, and precipitated cells and tissues were subjected to DNA extraction and qPCR to estimate symbiont-cell numbers of each reaction.LC–ESI–MS detection of 3M4N in feces from 3M4N-fed insectsAn insect-rearing system for feeding 3M4N and collecting feces is shown in Supplementary Fig. 7. Insects were fed with DW or DW containing 10 mM 3M4N in a plastic container, in which the solution supplier was covered by 0.5-mm mesh, so that insects were able to drink the solution by probing with their proboscis, but did not directly touch the solution by their legs or body. Twenty insects were reared per container and their feces were accumulated on the bottom of the container for five days. The collected feces (DW- or 3M4N-treated) were suspended in 1 ml of MilliQ water, and the water-soluble fractions were extracted by thorough vortexing. Solids and insoluble fractions were removed from the suspension by centrifugation and subsequent filtration using a cellulose-acetate membrane (Φ, 0.20 μm, ADVANTEC, Tokyo, Japan). The resultant fraction was diluted 10-fold by MilliQ water and analyzed by liquid chromatography–electrospray-ionization mass spectrometry (LC–ESI–MS) according to a previous report53,54,55. HPLC was performed using the Nexera X2 system (Shimadzu, Kyoto, Japan) composed of LC-30AD pump, SPD-M30A photodiode-array detector, and SIL-30AC autosampler. Develosil HB ODS-UG column (ID 2.0 mm × L 75 mm, Nomura Chemical Co., Ltd, Aichi, Japan) was employed with a flow rate of 0.2 mL/min. The following gradient system was used for analysis of metabolites: MilliQ water (solvent A) and methanol (solvent B), 90% A and 10% B at 0–5 min, linear gradient from 90% A and 10% B to 20% A and 80% B at 5–15 min, 20% A and 80% B at 15–20 min, and 90% A and 10% B at 20–25 min. Retention time of 3M4N standard reagent was 14.2 min. Electrospray-ionization mass spectrometry (ESI–MS) in positive and negative ion modes was simultaneously performed using amaZon SL (Bruker, Billerica, MA, USA). 3M4N (MW = 153.14) standard showed a clear peak in negative mode at m/z of 151.53.Reporting summaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

  • in

    Whale-cams reveal how much they really eat

    Nature Video
    05 November 2021

    Whale-cams reveal how much they really eat

    Baleen whales consume twice as much krill as previously estimated.

    Sara Reardon

    0

    Sara Reardon

    Sara Reardon is a freelance writer in Bozeman, Montana.

    View author publications

    You can also search for this author in PubMed
     Google Scholar

    Share on Twitter
    Share on Twitter

    Share on Facebook
    Share on Facebook

    Share via E-Mail
    Share via E-Mail

    Tagging whales with cameras and sensors has allowed researchers to calculate how much food these huge creatures are consuming. It’s the most accurate estimate yet and reveals an even more significant impact of whales on ocean ecosystems than was previously known.Read the paper here.

    doi: https://doi.org/10.1038/d41586-021-03026-z

    Related Articles

    Vikings were living in North America exactly a thousand years ago

    Strange patterns could be the world’s oldest animal fossil

    Divers are replanting reefs one coral at a time

    How cuttlefish wear their thoughts on their skin

    Swimming shrimp: Causing a stir

    Subjects

    Zoology

    Ecology

    Environmental sciences

    Latest on:

    Zoology

    Sponge cells hint at origins of nervous system
    News 05 NOV 21

    Baby bats try out their ‘sonar’ just after birth
    Research Highlight 27 OCT 21

    Ivory hunting drives evolution of tuskless elephants
    News 21 OCT 21

    Ecology

    Whales’ gigantic appetites, climate fears — the week in infographics
    News 05 NOV 21

    COP26 climate pledges: What scientists think so far
    News 05 NOV 21

    Baleen whale prey consumption based on high-resolution foraging measurements
    Article 03 NOV 21

    Environmental sciences

    Carbon implications of marginal oils from market-derived demand shocks
    Article 03 NOV 21

    For NGOs, article-processing charges sap conservation funds
    Correspondence 02 NOV 21

    Embrace open-source sensors for local climate studies
    Correspondence 02 NOV 21

    Jobs

    Postdoctoral Training Fellow

    Francis Crick Institute
    London, United Kingdom

    Head of GeMS

    Francis Crick Institute
    London, United Kingdom

    Postdoctoral scientist (m/f/div) to work on the comparative genomics of gutless marine oligochaetes

    Max Planck Institute for Marine Microbiology
    Bremen, Germany

    Postdoctoral Positions (m/f/div) in Protist Virology

    Max Planck Institute for Medical Research (MPIMF)
    Heidelberg, Germany More

  • in

    COP26 climate pledges: What scientists think so far

    NEWS
    05 November 2021

    COP26 climate pledges: What scientists think so far

    Nations have promised to end deforestation, curb methane emissions and stop public investment in coal power. Researchers warn that the real work of COP26 is yet to come.

    Ehsan Masood

    &

    Jeff Tollefson

    Ehsan Masood

    View author publications

    You can also search for this author in PubMed
     Google Scholar

    Jeff Tollefson

    View author publications

    You can also search for this author in PubMed
     Google Scholar

    Share on Twitter
    Share on Twitter

    Share on Facebook
    Share on Facebook

    Share via E-Mail
    Share via E-Mail

    Methane burns at an oil pit. Among the key pledges so far at COP26 is an agreement to cut methane emissions by 30% by 2030.Credit: Orjan F. Ellingvag/Corbis via Getty

    The first few days of the 26th United Nations Climate Change Conference of the Parties (COP26) have seen a flurry of announcements from world leaders promising to tackle climate change — from plans to phase out public finance for coal-fired power, to a pledge to end deforestation. This year, many big names — including US President Joe Biden and Indian Prime Minister Narendra Modi — attended the first two days of the conference to make big announcements.
    COP26 climate summit: A scientists’ guide to a momentous meeting
    This is different from what has happened at most previous COP summits, says Beth Martin, a specialist in climate negotiation who is part of RINGO (Research and Independent Non-Governmental Organizations), a network of organizations allowed to observe the COP26 negotiations. Usually, the highest-profile figures aren’t present during the first week, but arrive near the end of the meeting to help bridge differences in time for an agreed statement, and for the obligatory ‘UN family photo’.Nature asked researchers what they think of the pledges that have been made so far, as negotiators from some 200 countries prepare to dive into more detailed talks.Methane emissionsOne of the key developments in the first week was an agreement to curb emissions of methane, a powerful greenhouse gas that is second only to carbon dioxide in terms of its impact on the climate. Led by the United States and the European Union, the global methane pledge seeks to curb methane emissions by 30% by 2030, and has been signed by more than 100 countries.
    Control methane to slow global warming — fast
    “Obviously, as a scientist you’d say, ‘Well, a 50% reduction in the methane emissions by 2030 would be even better,’ but it’s a good start,” says Tim Lenton, who heads the Global Systems Institute at the University of Exeter, UK. “It’s an additional lever that could really help us limit warming.”Research has shown1 that curbing methane emissions using existing technologies could shave up to 0.5 °C off global temperatures by 2100. As with carbon dioxide, however, limiting methane emissions will not happen on its own.With his climate agenda facing challenges in Congress, Biden made methane a centrepiece of his commitments in Glasgow by announcing a new regulation to curb methane emissions from the oil and gas industry. Put forward this week by the US Environmental Protection Agency, the rule would require companies to curb methane emissions from their facilities by 74% over the coming decade, compared with 2005 levels. If implemented as proposed, it could prevent the release of some 37 million tonnes of methane by 2035 — equivalent to more than the annual carbon emissions from the nation’s fleets of passenger vehicles and commercial aircraft.India’s net-zero goalAfter delaying expected updates to India’s climate commitments by more than a year, Modi captured the world’s attention early in the summit by announcing that his country would seek to achieve net-zero emissions by 2070. The deadline is decades after that of many other countries that have made net-zero commitments, and it remains unclear whether India is committing to curbing just carbon dioxide emissions, or the broader category of greenhouse-gas emissions. But scientists say the announcement could mark a significant step forward if India follows through.
    Scientists cheer India’s ambitious carbon-zero climate pledge
    “We are definitely taken by surprise: this is much more than we were expecting to hear,” says Ulka Kelkar, an economist in Bengaluru who heads the Indian climate programme for the World Resources Institute, an environmental think tank based in Washington DC.Many scientists remain sceptical about mid-century net-zero pledges, in part because it’s easy to make long-term promises but hard to make the difficult short-term decisions that are required to meet those pledges. But India’s commitment includes measurable near-term targets, such as a pledge to provide 50% of the nation’s power through renewable resources and to reduce projected carbon emissions by one billion tonnes of carbon dioxide by 2030.Questions remain about how these targets will be defined and measured, but models indicate that there is a 50% chance such net-zero pledges could limit global warming to 2 °C or less, if fully implemented by all countries.

    More than 130 countries have agreed to halt and reverse deforestation by 2030.Credit: Joao Laet/AFP via Getty

    Climate cashAmong a cascade of climate-finance announcements this week is a pledge from more than 450 organizations in the financial sector — including banks, fund managers and insurance companies — in 45 countries to move US$130 trillion of funds under their control into investments where the recipient is committed to net-zero emissions by 2050.The pledging institutions, which are part of the Glasgow Financial Alliance for Net Zero, have not yet specified interim targets or timetables to achieve this goal. On 1 November, UN secretary-general António Guterres announced that a group of independent experts would be convened to propose standards for such commitments to net-zero emissions.
    The broken $100-billion promise of climate finance – and how to fix it
    Governments also announced new investments in clean technologies. And more than 40 countries, including the United Kingdom, Poland, South Korea and Vietnam, have committed to phasing out coal power in the 2030s (for major economies) or 2040s (globally), and to stopping public funding for new coal-fired power plants.“All of this is significant,” says Cristián Samper, an ecologist and president of the Wildlife Conservation Society in New York City. “The involvement of the financial sector and of ministers of finance and energy” in the meeting “is a game-changer”.However, the announcements have been overshadowed by governments’ failure to meet a 2009 pledge to provide $100 billion annually in climate finance for low- and middle-income countries by 2020. Reports suggest that it will take another two years to reach this goal, and that around 70% of the finance will be provided as loans.“We all assumed it would be grant finance. We didn’t pay attention to the fine print or expect that developed countries would hide behind loans,” says climate economist Tariq Banuri, a former director of sustainable development at the UN.Ending deforestationMore than 130 countries have pledged to halt and reverse forest-loss and land degradation by 2030. The signatories, which include Brazil, the Democratic Republic of the Congo and Indonesia, are home to 90% of the world’s forests.It is not the first such commitment: the 2014 New York Declaration on Forests, signed by a broad coalition of nearly 200 countries, regional governments, companies, indigenous groups and others, called for halving deforestation by 2020 and “striving” to end it by 2030.
    The United Nations must get its new biodiversity targets right
    There is also a long-standing UN pledge to slow down and eventually reverse the loss of biodiversity. But this remains unfulfilled and there is no official monitoring. Researchers say the latest target is unlikely to be met without an enforcement mechanism.Separately, a group of high-income countries has pledged $12 billion in public finance for forest protection between 2021 and 2025, but has not specified how the funding will be provided. A statement from the group, which includes Canada, the United States, the United Kingdom and EU countries, says governments will “work closely with the private sector” to “leverage vital funding from private sources to deliver change at scale”. This suggests that the finance is likely to be dominated by loans. Still, Samper says that there are reasons to be optimistic. Few previous climate COPs discussed nature and forests on the scale now seen in Glasgow. In the past, if biodiversity was mentioned at a climate meeting, “it was like the Martians had landed”, he says, because biodiversity and climate are treated as separate challenges by the UN. “We’ve never seen this much attention. It could be a pivot point.”

    doi: https://doi.org/10.1038/d41586-021-03034-z

    References1.Ocko, I. B. et al. Environ. Res. Lett. 16, 054042 (2021).Article 

    Google Scholar 
    Download references

    Related Articles

    COP26 climate summit: A scientists’ guide to a momentous meeting

    Scientists cheer India’s ambitious carbon-zero climate pledge

    The broken $100-billion promise of climate finance – and how to fix it

    The United Nations must get its new biodiversity targets right

    Control methane to slow global warming — fast

    Subjects

    Funding

    Biodiversity

    Politics

    Climate change

    Latest on:

    Funding

    The African Academy of Sciences is in crisis — responsibility must be shared
    Editorial 03 NOV 21

    UK research funding to grow slower than hoped
    News 28 OCT 21

    The high burden of infectious disease
    Nature Index 27 OCT 21

    Biodiversity

    The answer to the biodiversity crisis is not more debt
    Editorial 26 OCT 21

    Illegal mining in the Amazon hits record high amid Indigenous protests
    News 30 SEP 21

    Fine-root traits in the global spectrum of plant form and function
    Article 29 SEP 21

    Politics

    All aboard the climate train! Scientists join activists for COP26 trip
    News 02 NOV 21

    Top climate scientists are sceptical that nations will rein in global warming
    News Feature 01 NOV 21

    UK research funding to grow slower than hoped
    News 28 OCT 21

    Jobs

    Postdoctoral Training Fellow

    Francis Crick Institute
    London, United Kingdom

    Head of GeMS

    Francis Crick Institute
    London, United Kingdom

    Postdoctoral scientist (m/f/div) to work on the comparative genomics of gutless marine oligochaetes

    Max Planck Institute for Marine Microbiology
    Bremen, Germany

    Postdoctoral Positions (m/f/div) in Protist Virology

    Max Planck Institute for Medical Research (MPIMF)
    Heidelberg, Germany More

  • in

    Antibiotic resistance in the environment

    1.D’Costa, V. M. et al. Antibiotic resistance is ancient. Nature 477, 457–461 (2011). This study shows that different ARGs are present in 30,000-year-old permafrost.
    Google Scholar 
    2.Bhullar, K. et al. Antibiotic resistance is prevalent in an isolated cave microbiome. PLoS ONE 7, e34953 (2012).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    3.Lugli, G. A. et al. Ancient bacteria of the Ötzi’s microbiome: a genomic tale from the Copper Age. Microbiome 5, 5 (2017).PubMed 
    PubMed Central 

    Google Scholar 
    4.Perry, J., Waglechner, N. & Wright, G. The prehistory of antibiotic resistance. Cold Spring Harb. Perspect. Med. 6, a025197 (2016).PubMed 
    PubMed Central 

    Google Scholar 
    5.Davies, J. & Davies, D. Origins and evolution of antibiotic resistance. Microbiol. Mol. Biol. Rev. 74, 417–433 (2010). This authoritative and educational review discusses in an insightful way the evolution of resistance, including its origins and future implications.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    6.Allen, H. K. et al. Call of the wild: antibiotic resistance genes in natural environments. Nat. Rev. Microbiol. 8, 251–259 (2010).CAS 
    PubMed 

    Google Scholar 
    7.Martinez, J. L. The role of natural environments in the evolution of resistance traits in pathogenic bacteria. Proc. R. Soc. B Biol. Sci. 276, 2521–2530 (2009).
    Google Scholar 
    8.Alcock, B. P. et al. CARD 2020: antibiotic resistome surveillance with the comprehensive antibiotic resistance database. Nucleic Acids Res. https://doi.org/10.1093/nar/gkz935 (2019).Article 
    PubMed Central 

    Google Scholar 
    9.Mackenzie, J. S. & Jeggo, M. The one health approach — why is it so important? Trop. Med. Infect. Dis. 4, 88 (2019).PubMed Central 

    Google Scholar 
    10.Buschhardt, T. et al. A one health glossary to support communication and information exchange between the human health, animal health and food safety sectors. One Health 13, 100263 (2021).PubMed 
    PubMed Central 

    Google Scholar 
    11.Berendonk, T. U. et al. Tackling antibiotic resistance: the environmental framework. Nat. Rev. Microbiol. 13, 310–317 (2015).CAS 
    PubMed 

    Google Scholar 
    12.Wellington, E. M. et al. The role of the natural environment in the emergence of antibiotic resistance in gram-negative bacteria. Lancet Infect. Dis. 13, 155–165 (2013).CAS 
    PubMed 

    Google Scholar 
    13.Bengtsson-Palme, J., Kristiansson, E. & Larsson, D. G. J. Environmental factors influencing the development and spread of antibiotic resistance. FEMS Microbiol. Rev. https://doi.org/10.1093/femsre/fux053 (2017).Article 
    PubMed Central 

    Google Scholar 
    14.Chow, L. K. M., Ghaly, T. M. & Gillings, M. R. A survey of sub-inhibitory concentrations of antibiotics in the environment. J. Environ. Sci. 99, 21–27 (2021).
    Google Scholar 
    15.Andersson, D. I. et al. Antibiotic resistance: turning evolutionary principles into clinical reality. FEMS Microbiol. Rev. 44, 171–188 (2020).CAS 
    PubMed 

    Google Scholar 
    16.Singer, A. C., Shaw, H., Rhodes, V. & Hart, A. Review of antimicrobial resistance in the environment and its relevance to environmental regulators. Front. Microbiol. https://doi.org/10.3389/fmicb.2016.01728 (2016).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    17.United Nations Environment Programme. Frontiers 2017: emerging issues of environmental concern, https://www.unenvironment.org/resources/frontiers-2017-emerging-issues-environmental-concern (2017).18.Access to Medicines Foundation. 2020 antimicrobial resistance benchmark, https://accesstomedicinefoundation.org/publications/2020-antimicrobial-resistance-benchmark (2020).19.Review on Antimicrobial Resistance. Antimicrobials in agriculture and the environment: reducing unnecessary waste, https://amr-review.org/Publications.html (2015).20.European Parliament. Strategic approach to pharmaceuticals in the environment, https://www.europarl.europa.eu/doceo/document/TA-9-2020-0226_EN.pdf (2020).21.WHO. Technical brief on water, sanitation, hygiene (WASH) and wastewater management to prevent infections and reduce the spread of antimicrobial resistance (AMR)., https://www.who.int/water_sanitation_health/publications/wash-wastewater-management-to-prevent-infections-and-reduce-amr/en/ (2020).22.Graham, D. W. et al. Complexities in understanding antimicrobial resistance across domesticated animal, human, and environmental systems. Ann. N. Y. Acad. Sci. 1441, 17–30 (2019).PubMed 
    PubMed Central 

    Google Scholar 
    23.Smalla, K., Cook, K., Djordjevic, S. P., Klümper, U. & Gillings, M. Environmental dimensions of antibiotic resistance: assessment of basic science gaps. FEMS Microbiol. Ecol. https://doi.org/10.1093/femsec/fiy195 (2018).Article 
    PubMed 

    Google Scholar 
    24.Rinke, C. et al. Insights into the phylogeny and coding potential of microbial dark matter. Nature 499, 431–437 (2013).CAS 
    PubMed 

    Google Scholar 
    25.Schulz, F. et al. Towards a balanced view of the bacterial tree of life. Microbiome https://doi.org/10.1186/s40168-017-0360-9 (2017).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    26.Forsberg, K. J. et al. The shared antibiotic resistome of soil bacteria and human pathogens. Science 337, 1107–1111 (2012). This study demonstrates numerous identical resistance gene loci between multiresistant soil bacteria and diverse human pathogens, providing evidence for recent gene exchange across species and environments.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    27.Berglund, F. et al. Identification of 76 novel B1 metallo-beta-lactamases through large-scale screening of genomic and metagenomic data. Microbiome 5, 134 (2017).PubMed 
    PubMed Central 

    Google Scholar 
    28.Dantas, G., Sommer, M. O. A., Oluwasegun, R. D. & Church, G. M. Bacteria subsisting on antibiotics. Science 320, 100–103 (2008).CAS 
    PubMed 

    Google Scholar 
    29.Berglund, F. et al. Comprehensive screening of genomic and metagenomic data reveals a large diversity of tetracycline resistance genes. Microb. Genomics https://doi.org/10.1099/mgen.0.000455 (2020).Article 

    Google Scholar 
    30.Pawlowski, A. C. et al. A diverse intrinsic antibiotic resistome from a cave bacterium. Nat. Commun. 7, 13803 (2016).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    31.Morar, M. & Wright, G. D. The genomic enzymology of antibiotic resistance. Annu. Rev. Genet. 44, 25–51 (2010).CAS 
    PubMed 

    Google Scholar 
    32.Andersson, D. I., Jerlström-Hultqvist, J. & Näsvall, J. Evolution of new functions de novo and from preexisting genes. Cold Spring Harb. Perspect. Biol. 7, a017996 (2015).PubMed 
    PubMed Central 

    Google Scholar 
    33.Razavi, M., Kristiansson, E., Flach, C.-F. & Larsson, D. G. J. The association between insertion sequences and antibiotic resistance genes. mSphere https://doi.org/10.1128/msphere.00418-20 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    34.Partridge, S. R., Kwong, S. M., Firth, N. & Jensen, S. O. Mobile genetic elements associated with antimicrobial resistance. Clin. Microbiol. Rev. https://doi.org/10.1128/cmr.00088-17 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    35.Gillings, M. et al. The evolution of class 1 integrons and the rise of antibiotic resistance. J. Bacteriol. 190, 5095–5100 (2008).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    36.Razavi, M. et al. Discovery of the fourth mobile sulfonamide resistance gene. Microbiome https://doi.org/10.1186/s40168-017-0379-y (2017).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    37.Flach, C.-F. et al. Does antifouling paint select for antibiotic resistance? Sci. Total Environ. 590–591, 461–468 (2017).PubMed 

    Google Scholar 
    38.Shintani, M. et al. Plant species-dependent increased abundance and diversity of IncP-1 plasmids in the rhizosphere: new insights into their role and ecology. Front. Microbiol. 11, 590776 (2020).PubMed 
    PubMed Central 

    Google Scholar 
    39.Baquero, F., Coque, T. M., Martínez, J.-L., Aracil-Gisbert, S. & Lanza, V. F. Gene transmission in the one health microbiosphere and the channels of antimicrobial resistance. Front. Microbiol. https://doi.org/10.3389/fmicb.2019.02892 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    40.Vandecraen, J., Chandler, M., Aertsen, A. & Van Houdt, R. The impact of insertion sequences on bacterial genome plasticity and adaptability. Crit. Rev. Microbiol. 43, 709–730 (2017).CAS 
    PubMed 

    Google Scholar 
    41.Depardieu, F., Podglajen, I., Leclercq, R., Collatz, E. & Courvalin, P. Modes and modulations of antibiotic resistance gene expression. Clin. Microbiol. Rev. 20, 79–114 (2007).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    42.Jutkina, J., Marathe, N. P., Flach, C. F. & Larsson, D. G. J. Antibiotics and common antibacterial biocides stimulate horizontal transfer of resistance at low concentrations. Sci. Total Environ. 616-617, 172–178 (2018).CAS 
    PubMed 

    Google Scholar 
    43.Scornec, H., Bellanger, X., Guilloteau, H., Groshenry, G. & Merlin, C. Inducibility of Tn916 conjugative transfer in Enterococcus faecalis by subinhibitory concentrations of ribosome-targeting antibiotics. J. Antimicrob. Chemother. 72, 2722–2728 (2017).CAS 
    PubMed 

    Google Scholar 
    44.Aminov, R. I. Horizontal gene exchange in environmental microbiota. Front. Microbiol. https://doi.org/10.3389/fmicb.2011.00158 (2011).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    45.Knöppel, A., Näsvall, J. & Andersson, D. I. Evolution of antibiotic resistance without antibiotic exposure. Antimicrob. Agents Chemother. https://doi.org/10.1128/aac.01495-17 (2017).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    46.Kimura, M. & Ohta, T. The average number of generations until fixation of a mutant gene in a finite population. Genetics 61, 763–771 (1969).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    47.Letten, A. D., Hall, A. R. & Levine, J. M. Using ecological coexistence theory to understand antibiotic resistance and microbial competition. Nat. Ecol. Evol. 5, 431–441 (2021).PubMed 

    Google Scholar 
    48.Waglechner, N. & Wright, G. D. Antibiotic resistance: it’s bad, but why isn’t it worse? BMC Biol. https://doi.org/10.1186/s12915-017-0423-1 (2017).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    49.Ebmeyer, S., Erik, K. & Larsson, D. G. J. A framework for identifying the recent origins of mobile antibiotic resistance genes. Commun. Biol. https://doi.org/10.1038/s42003-020-01545-5 (2021). This study amends, summarizes and scrutinizes current evidence for proposed recent origin species for mobile ARGs.Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    50.Andersson, D. I. & Hughes, D. Persistence of antibiotic resistance in bacterial populations. FEMS Microbiol. Rev. 35, 901–911 (2011).CAS 
    PubMed 

    Google Scholar 
    51.Wang, J., Chu, L., Wojnárovits, L. & Takács, E. Occurrence and fate of antibiotics, antibiotic resistant genes (ARGs) and antibiotic resistant bacteria (ARB) in municipal wastewater treatment plant: an overview. Sci. Total. Environ. 744, 140997 (2020).CAS 
    PubMed 

    Google Scholar 
    52.Tran, N. H., Reinhard, M. & Gin, K. Y.-H. Occurrence and fate of emerging contaminants in municipal wastewater treatment plants from different geographical regions-a review. Water Res. 133, 182–207 (2018).CAS 
    PubMed 

    Google Scholar 
    53.Szymańska, U. et al. Presence of antibiotics in the aquatic environment in Europe and their analytical monitoring: recent trends and perspectives. Microchem. J. 147, 729–740 (2019).
    Google Scholar 
    54.Anwar, M., Iqbal, Q. & Saleem, F. Improper disposal of unused antibiotics: an often overlooked driver of antimicrobial resistance. Expert Rev. Antiinfect Ther. https://doi.org/10.1080/14787210.2020.1754797 (2020).Article 

    Google Scholar 
    55.Cabello, F. C. et al. Antimicrobial use in aquaculture re-examined: its relevance to antimicrobial resistance and to animal and human health. Environ. Microbiol. 15, 1917–1942 (2013).PubMed 

    Google Scholar 
    56.Cabello, F. C., Godfrey, H. P., Buschmann, A. H. & Dölz, H. J. Aquaculture as yet another environmental gateway to the development and globalisation of antimicrobial resistance. Lancet Infect. Dis. 16, e127–e133 (2016).PubMed 

    Google Scholar 
    57.Taylor, P. & Reeder, R. Antibiotic use on crops in low and middle-income countries based on recommendations made by agricultural advisors. CABI Agric. Biosci. https://doi.org/10.1186/s43170-020-00001-y (2020).Article 

    Google Scholar 
    58.Larsson, D. G. J. Pollution from drug manufacturing: review and perspectives. Philos. Trans. R. Soc. B Biol. Sci. 369, 20130571 (2014).
    Google Scholar 
    59.Larsson, D. G. J., De Pedro, C. & Paxeus, N. Effluent from drug manufactures contains extremely high levels of pharmaceuticals. J. Hazard. Mater. 148, 751–755 (2007).CAS 
    PubMed 

    Google Scholar 
    60.Milaković, M. et al. Pollution from azithromycin-manufacturing promotes macrolide-resistance gene propagation and induces spatial and seasonal bacterial community shifts in receiving river sediments. Environ. Int. 123, 501–511 (2019).PubMed 

    Google Scholar 
    61.Bielen, A. et al. Negative environmental impacts of antibiotic-contaminated effluents from pharmaceutical industries. Water Res. 126, 79–87 (2017).CAS 
    PubMed 

    Google Scholar 
    62.Fick, J. et al. Contamination of surface, ground, and drinking water from pharmaceutical production. Environ. Toxicol. Chem. 28, 2522–2527 (2009).CAS 
    PubMed 

    Google Scholar 
    63.Bengtsson-Palme, J. & Larsson, D. G. J. Concentrations of antibiotics predicted to select for resistant bacteria: proposed limits for environmental regulation. Environ. Int. 86, 140–149 (2016). This study uses a simplified approach based on available MIC data for many species to predict concentrations of 111 antibiotics that are not likely to select for resistance.CAS 
    PubMed 

    Google Scholar 
    64.Gullberg, E. et al. Selection of resistant bacteria at very low antibiotic concentrations. PLoS Pathog. 7, e1002158 (2011).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    65.Karkman, A., Pärnänen, K. & Larsson, D. G. J. Fecal pollution can explain antibiotic resistance gene abundances in anthropogenically impacted environments. Nat. Commun. https://doi.org/10.1038/s41467-018-07992-3 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    66.Yang, Y., Li, B., Zou, S., Fang, H. H. P. & Zhang, T. Fate of antibiotic resistance genes in sewage treatment plant revealed by metagenomic approach. Water Res. 62, 97–106 (2014).CAS 
    PubMed 

    Google Scholar 
    67.Bengtsson-Palme, J. et al. Elucidating selection processes for antibiotic resistance in sewage treatment plants using metagenomics. Sci. Total Environ. 572, 697–712 (2016).CAS 
    PubMed 

    Google Scholar 
    68.Manaia, C. M. et al. Antibiotic resistance in wastewater treatment plants: tackling the black box. Environ. Int. 115, 312–324 (2018).CAS 
    PubMed 

    Google Scholar 
    69.Flach, C. F., Genheden, M., Fick, J. & Joakim Larsson, D. G. A comprehensive screening of Escherichia coli isolates from Scandinavia’s largest sewage treatment plant indicates no selection for antibiotic resistance. Environ. Sci. Technol. 52, 11419–11428 (2018).CAS 
    PubMed 

    Google Scholar 
    70.Kraupner, N. et al. Evidence for selection of multi-resistant E. coli by hospital effluent. Environ. Int. 150, 106436 (2021).CAS 
    PubMed 

    Google Scholar 
    71.Flach, C. F. et al. Isolation of novel IncA/C and IncN fluoroquinolone resistance plasmids from an antibiotic-polluted lake. J. Antimicrob. Chemother. 70, 2709–2717 (2015).CAS 
    PubMed 

    Google Scholar 
    72.Bengtsson-Palme, J., Boulund, F., Fick, J., Kristiansson, E. & Larsson, D. G. J. Shotgun metagenomics reveals a wide array of antibiotic resistance genes and mobile elements in a polluted lake in India. Front. Microbiol. https://doi.org/10.3389/fmicb.2014.00648 (2014).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    73.Marathe, N. P. et al. Functional metagenomics reveals a novel carbapenem-hydrolyzing mobile beta-lactamase from Indian river sediments contaminated with antibiotic production waste. Environ. Int. 112, 279–286 (2018).CAS 
    PubMed 

    Google Scholar 
    74.Thiele-Bruhn, S. Pharmaceutical antibiotic compounds in soils–a review. J. Plant Nutr. Soil Sci. 166, 145–167 (2003).CAS 

    Google Scholar 
    75.Li, W., Shi, Y., Gao, L., Liu, J. & Cai, Y. Occurrence, distribution and potential affecting factors of antibiotics in sewage sludge of wastewater treatment plants in China. Sci. Total. Environ. 445–446, 306–313 (2013).PubMed 

    Google Scholar 
    76.Reinthaler, F. F. et al. Resistance patterns of Escherichia coli isolated from sewage sludge in comparison with those isolated from human patients in 2000 and 2009. J. Water Health 11, 13–20 (2013).PubMed 

    Google Scholar 
    77.Rutgersson, C. et al. Long-term application of Swedish sewage sludge on farmland does not cause clear changes in the soil bacterial resistome. Environ. Int. 137, 105339 (2020).CAS 
    PubMed 

    Google Scholar 
    78.Jechalke, S., Heuer, H., Siemens, J., Amelung, W. & Smalla, K. Fate and effects of veterinary antibiotics in soil. Trends Microbiol. 22, 536–545 (2014).CAS 
    PubMed 

    Google Scholar 
    79.Boxall, A. B. et al. Pharmaceuticals and personal care products in the environment: what are the big questions? Environ. Health Perspect. 120, 1221–1229 (2012).PubMed 
    PubMed Central 

    Google Scholar 
    80.Song, J., Rensing, C., Holm, P. E., Virta, M. & Brandt, K. K. Comparison of metals and tetracycline as selective agents for development of tetracycline resistant bacterial communities in agricultural soil. Environ. Sci. Technol. 51, 3040–3047 (2017).CAS 
    PubMed 

    Google Scholar 
    81.Jechalke, S. et al. Plasmid-mediated fitness advantage of Acinetobacter baylyi in sulfadiazine-polluted soil. FEMS Microbiol. Lett. 348, 127–132 (2013). This study shows that a commonly used antibiotic in pig farming has the potential to select for a resistant Acinetobacter strain in manure-amended soils.CAS 
    PubMed 

    Google Scholar 
    82.Pal, C. et al. Metal resistance and its association with antibiotic resistance. Adv. Microb. Physiol. 70, 261–313 (2017).CAS 
    PubMed 

    Google Scholar 
    83.Wales, A. & Davies, R. Co-selection of resistance to antibiotics, biocides and heavy metals, and its relevance to foodborne pathogens. Antibiotics 4, 567–604 (2015).PubMed 
    PubMed Central 

    Google Scholar 
    84.Pal, C., Bengtsson-Palme, J., Kristiansson, E. & Larsson, D. G. J. Co-occurrence of resistance genes to antibiotics, biocides and metals reveals novel insights into their co-selection potential. BMC Genomics https://doi.org/10.1186/s12864-015-2153-5 (2015).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    85.Klümper, U. et al. Metal stressors consistently modulate bacterial conjugal plasmid uptake potential in a phylogenetically conserved manner. ISME J. 11, 152–165 (2017).PubMed 

    Google Scholar 
    86.Jutkina, J., Rutgersson, C., Flach, C. F. & Joakim Larsson, D. G. An assay for determining minimal concentrations of antibiotics that drive horizontal transfer of resistance. Sci. Total. Environ. 548–549, 131–138 (2016).PubMed 

    Google Scholar 
    87.Wang, Y. et al. Non-antibiotic pharmaceuticals enhance the transmission of exogenous antibiotic resistance genes through bacterial transformation. ISME J. 14, 2179–2196 (2020).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    88.Klumper, U. et al. Broad host range plasmids can invade an unexpectedly diverse fraction of a soil bacterial community. ISME J. 9, 934–945 (2015). This study shows that plasmids that are common in pathogens can easily be taken up by diverse environmental bacteria, thereby providing pathways for the exchange of resistance genes.CAS 
    PubMed 

    Google Scholar 
    89.Gillings, M. R., Paulsen, I. T. & Tetu, S. G. Genomics and the evolution of antibiotic resistance. Ann. N. Y. Acad. Sci. 1388, 92–107 (2017).PubMed 

    Google Scholar 
    90.Heuer, H. & Smalla, K. Plasmids foster diversification and adaptation of bacterial populations in soil. FEMS Microbiol. Rev. 36, 1083–1104 (2012).CAS 
    PubMed 

    Google Scholar 
    91.Bengtsson-Palme, J. & Larsson, D. G. Antibiotic resistance genes in the environment: prioritizing risks. Nat. Rev. Microbiol. 13, 396 (2015).CAS 
    PubMed 

    Google Scholar 
    92.Leonard, A. F. C. et al. Exposure to and colonisation by antibiotic-resistant E. coli in UK coastal water users: environmental surveillance, exposure assessment, and epidemiological study (Beach Bum Survey). Environ. Int. 114, 326–333 (2018). This is one of few studies showing that people more likely to ingest surface waters are also more prone to be carriers of resistant bacteria compared with matched controls.PubMed 

    Google Scholar 
    93.Manaia, C. M. Assessing the risk of antibiotic resistance transmission from the environment to humans: non-direct proportionality between abundance and risk. Trends Microbiol. 25, 173–181 (2017).CAS 
    PubMed 

    Google Scholar 
    94.Schijven, J. F., Blaak, H., Schets, F. M. & De Roda Husman, A. M. Fate of extended-spectrum β-lactamase-producing Escherichia coli from faecal sources in surface water and probability of human exposure through swimming. Environ. Sci. Technol. 49, 11825–11833 (2015).CAS 
    PubMed 

    Google Scholar 
    95.Collignon, P., Beggs, J. J., Walsh, T. R., Gandra, S. & Laxminarayan, R. Anthropological and socioeconomic factors contributing to global antimicrobial resistance: a univariate and multivariable analysis. Lancet Planet. Health 2, e398–e405 (2018).PubMed 

    Google Scholar 
    96.Dancer, S. J. Controlling hospital-acquired infection: focus on the role of the environment and new technologies for decontamination. Clin. Microbiol. Rev. 27, 665–690 (2014).PubMed 
    PubMed Central 

    Google Scholar 
    97.Weber, D. J., Anderson, D. & Rutala, W. A. The role of the surface environment in healthcare-associated infections. Curr. Opin. Infect. Dis. 26, 338–344 (2013).PubMed 

    Google Scholar 
    98.Søraas, A., Sundsfjord, A., Sandven, I., Brunborg, C. & Jenum, P. A. Risk factors for community-acquired urinary tract infections caused by ESBL-producing Enterobacteriaceae –a case–control study in a low prevalence country. PLoS ONE 8, e69581 (2013).PubMed 
    PubMed Central 

    Google Scholar 
    99.Zhou, S.-Y.-D. et al. Prevalence of antibiotic resistome in ready-to-eat salad. Front. Public Health https://doi.org/10.3389/fpubh.2020.00092 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    100.Uyttendaele, M. et al. Microbial hazards in irrigation water: standards, norms, and testing to manage use of water in fresh produce primary production. Compr. Rev. Food Sci. Food Saf. 14, 336–356 (2015).
    Google Scholar 
    101.Reid, C. J., Blau, K., Jechalke, S., Smalla, K. & Djordjevic, S. P. Whole genome sequencing of Escherichia coli from store-bought produce. Front. Microbiol. 10, 3050 (2020).PubMed 
    PubMed Central 

    Google Scholar 
    102.Blau, K. et al. The transferable resistome of produce. mBio 9, e01300-18 (2018).PubMed 
    PubMed Central 

    Google Scholar 
    103.Zhu, Y.-G. et al. Soil biota, antimicrobial resistance and planetary health. Environ. Int. 131, 105059 (2019).PubMed 

    Google Scholar 
    104.Pal, C., Bengtsson-Palme, J., Kristiansson, E. & Larsson, D. G. J. The structure and diversity of human, animal and environmental resistomes. Microbiome 4, 54 (2016).PubMed 
    PubMed Central 

    Google Scholar 
    105.Kozajda, A., Jeżak, K. & Kapsa, A. Airborne Staphylococcus aureus in different environments — a review. Environ. Sci. Pollut. Res. 26, 34741–34753 (2019).CAS 

    Google Scholar 
    106.Ashbolt, N. J. et al. Human health risk assessment (HHRA) for environmental development and transfer of antibiotic resistance. Environ. Health Perspect. 121, 993–1001 (2013).PubMed 
    PubMed Central 

    Google Scholar 
    107.Franz, E., Schijven, J., De Roda Husman, A. M. & Blaak, H. Meta-regression analysis of commensal and pathogenic Escherichia coli survival in soil and water. Environ. Sci. Technol. 48, 6763–6771 (2014).CAS 
    PubMed 

    Google Scholar 
    108.Lewis, K. Platforms for antibiotic discovery. Nat. Rev. Drug. Discov. 12, 371–387 (2013).CAS 
    PubMed 

    Google Scholar 
    109.Linton, K. B., Richmond, M. H., Bevan, R. & Gillespie, W. A. Antibiotic resistance and R factors in coliform bacilli isolated from hospital and domestic sewage. J. Med. Microbiol. 7, 91–103 (1974).CAS 
    PubMed 

    Google Scholar 
    110.Huijbers, P., Joakim Larsson, D. G. & Flach, C. F. Surveillance of antibiotic resistant Escherichia coli in human populations through urban wastewater in ten European countries. Environ. Pollut. 261, 114200 (2020).CAS 
    PubMed 

    Google Scholar 
    111.Hutinel, M. et al. Population-level surveillance of antibiotic resistance in Escherichia coli through sewage analysis. Euro Surveill. https://doi.org/10.2807/1560-7917.es.2019.24.37.1800497 (2019).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    112.Aarestrup, F. M. & Woolhouse, M. E. J. Using sewage for surveillance of antimicrobial resistance. Science 367, 630–632 (2020).CAS 
    PubMed 

    Google Scholar 
    113.Kwak, Y. K. et al. Surveillance of antimicrobial resistance among Escherichia coli in wastewater in Stockholm during 1 year: does it reflect the resistance trends in the society? Int. J. Antimicrob. Agents 45, 25–32 (2015).CAS 
    PubMed 

    Google Scholar 
    114.Parnanen, K. M. M. et al. Antibiotic resistance in European wastewater treatment plants mirrors the pattern of clinical antibiotic resistance prevalence. Sci. Adv. 5, eaau9124 (2019).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    115.Hendriksen, R. S. et al. Global monitoring of antimicrobial resistance based on metagenomics analyses of urban sewage. Nat. Commun. 10, 1124 (2019). This is the most comprehensive survey of ARGs in sewage across the world to date, showing distinct differences between regions.PubMed 
    PubMed Central 

    Google Scholar 
    116.Huijbers, P. M. C., Flach, C. F. & Larsson, D. G. J. A conceptual framework for the environmental surveillance of antibiotics and antibiotic resistance. Environ. Int. 130, 104880 (2019).CAS 
    PubMed 

    Google Scholar 
    117.Böhm, M.-E., Razavi, M., Marathe, N. P., Flach, C.-F. & Larsson, D. G. J. Discovery of a novel integron-borne aminoglycoside resistance gene present in clinical pathogens by screening environmental bacterial communities. Microbiome https://doi.org/10.1186/s40168-020-00814-z (2020). Using a functional assay targeting mobile genes, this study explores environment communities and finds a completely novel resistance gene that had escaped discovery in clinics despite its presence in pathogens on different continents.Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    118.Flach, C.-F., Hutinel, M., Razavi, M., Åhrén, C. & Larsson, D. G. J. Monitoring of hospital sewage shows both promise and limitations as an early-warning system for carbapenemase-producing Enterobacterales in a low-prevalence setting. Water Res. 200, 117261 (2021).CAS 
    PubMed 

    Google Scholar 
    119.Karkman, A., Berglund, F., Flach, C.-F., Kristiansson, E. & Larsson, D. G. J. Predicting clinical resistance prevalence using sewage metagenomic data. Commun. Biol. https://doi.org/10.1038/s42003-020-01439-6 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    120.European Centre for Disease Prevention and Control. Surveillance of antimicrobial resistance in Europe 2017 (Stockholm, Sweden, 2018).121.Hovi, T. et al. Role of environmental poliovirus surveillance in global polio eradication and beyond. Epidemiol. Infect. 140, 1–13 (2012).CAS 
    PubMed 

    Google Scholar 
    122.Agrawal, S., Orschler, L. & Lackner, S. Long-term monitoring of SARS-CoV-2 RNA in wastewater of the Frankfurt metropolitan area in southern Germany. Sci. Rep. https://doi.org/10.1038/s41598-021-84914-2 (2021).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    123.Medema, G., Heijnen, L., Elsinga, G., Italiaander, R. & Brouwer, A. Presence of SARS-coronavirus-2 RNA in sewage and correlation with reported COVID-19 prevalence in the early stage of the epidemic in the Netherlands. Environ. Sci. Technol. Lett. 7, 511–516 (2020).CAS 

    Google Scholar 
    124.Lundstrom, S. V. et al. Minimal selective concentrations of tetracycline in complex aquatic bacterial biofilms. Sci. Total Environ. 553, 587–595 (2016).PubMed 

    Google Scholar 
    125.McCann, C. M. et al. Understanding drivers of antibiotic resistance genes in High Arctic soil ecosystems. Environ. Int. 125, 497–504 (2019).CAS 
    PubMed 

    Google Scholar 
    126.Pruden, A., Arabi, M. & Storteboom, H. N. Correlation between upstream human activities and riverine antibiotic resistance genes. Environ. Sci. Technol. 46, 11541–11549 (2012).CAS 
    PubMed 

    Google Scholar 
    127.Zhu, Y.-G. et al. Continental-scale pollution of estuaries with antibiotic resistance genes. Nat. Microbiol. 2, 16270 (2017).CAS 
    PubMed 

    Google Scholar 
    128.Zhu, Y.-G. et al. Diverse and abundant antibiotic resistance genes in Chinese swine farms. Proc. Natl Acad. Sci. USA 110, 3435–3440 (2013).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    129.Knapp, C. W., Dolfing, J., Ehlert, P. A. I. & Graham, D. W. Evidence of increasing antibiotic resistance gene abundances in archived soils since 1940. Environ. Sci. Technol. 44, 580–587 (2010).CAS 
    PubMed 

    Google Scholar 
    130.Nesme, J. & Simonet, P. The soil resistome: a critical review on antibiotic resistance origins, ecology and dissemination potential in telluric bacteria. Environ. Microbiol. 17, 913–930 (2015).PubMed 

    Google Scholar 
    131.Finley, R. L. et al. The scourge of antibiotic resistance: the important role of the environment. Clin. Infect. Dis. 57, 704–710 (2013).PubMed 

    Google Scholar 
    132.Sjölund, M. et al. Dissemination of multidrug-resistant bacteria into the Arctic. Emerg. Infect. Dis. 14, 70–72 (2008).PubMed 
    PubMed Central 

    Google Scholar 
    133.Zhu, G. et al. Air pollution could drive global dissemination of antibiotic resistance genes. ISME J. https://doi.org/10.1038/s41396-020-00780-2 (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    134.Nichols, D. et al. Use of Ichip for high-throughput in situ cultivation of “Uncultivable” microbial species. Appl. Environ. Microbiol. 76, 2445–2450 (2010).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    135.Ashton, P. M. et al. MinION nanopore sequencing identifies the position and structure of a bacterial antibiotic resistance island. Nat. Biotechnol. 33, 296–300 (2015).CAS 
    PubMed 

    Google Scholar 
    136.Spencer, S. J. et al. Massively parallel sequencing of single cells by epicPCR links functional genes with phylogenetic markers. ISME J. 10, 427–436 (2016).CAS 
    PubMed 

    Google Scholar 
    137.Rice, E. W., Wang, P., Smith, A. L. & Stadler, L. B. Determining hosts of antibiotic resistance genes: a review of methodological advances. Environ. Sci. Technol. Lett. 7, 282–291 (2020).CAS 

    Google Scholar 
    138.Sivalingam, P., Poté, J. & Prabakar, K. Extracellular DNA (eDNA): neglected and potential sources of antibiotic resistant genes (ARGs) in the aquatic environments. Pathogens 9, 874 (2020).CAS 
    PubMed Central 

    Google Scholar 
    139.Bengtsson-Palme, J., Larsson, D. G. J. & Kristiansson, E. Using metagenomics to investigate human and environmental resistomes. J. Antimicrob. Chemother. 72, 2690–2703 (2017).CAS 
    PubMed 

    Google Scholar 
    140.Karkman, A. et al. High-throughput quantification of antibiotic resistance genes from an urban wastewater treatment plant. FEMS Microbiol. Ecol. 92, https://doi.org/10.1093/femsec/fiw014 (2016).141.Gillings, M. R. et al. Using the class 1 integron-integrase gene as a proxy for anthropogenic pollution. ISME J. 9, 1269–1279 (2015).CAS 
    PubMed 

    Google Scholar 
    142.Gaze, W. H., Abdouslam, N., Hawkey, P. M. & Wellington, E. M. H. Incidence of Class 1 integrons in a quaternary ammonium compound-polluted environment. Antimicrob. Agents Chemother. 49, 1802–1807 (2005).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    143.Sommer, M. O. A., Munck, C., Toft-Kehler, R. V. & Andersson, D. I. Prediction of antibiotic resistance: time for a new preclinical paradigm? Nat. Rev. Microbiol. 15, 689–696 (2017). This article highlights the needs to consider the environmental gene reservoir and other factors influencing resistance evolution in the development process for new antibiotics.CAS 
    PubMed 

    Google Scholar 
    144.Pehrsson, E. C., Forsberg, K. J., Gibson, M. K., Ahmadi, S. & Dantas, G. Novel resistance functions uncovered using functional metagenomic investigations of resistance reservoirs. Front. Microbiol. https://doi.org/10.3389/fmicb.2013.00145 (2013).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    145.Kim, C., Ryu, H.-D., Chung, E. G., Kim, Y. & Lee, J.-K. A review of analytical procedures for the simultaneous determination of medically important veterinary antibiotics in environmental water: sample preparation, liquid chromatography, and mass spectrometry. J. Environ. Manag. 217, 629–645 (2018).CAS 

    Google Scholar 
    146.Fahrenfeld, N. & Bisceglia, K. J. Emerging investigators series: sewer surveillance for monitoring antibiotic use and prevalence of antibiotic resistance: urban sewer epidemiology. Environ. Sci. Water Res. Technol. 2, 788–799 (2016).CAS 

    Google Scholar 
    147.Anliker, S. et al. Assessing emissions from pharmaceutical manufacturing based on temporal high-resolution mass spectrometry data. Environ. Sci. Technol. 54, 4110–4120 (2020). This recent study elegantly uses the erratic emission profiles of drugs from manufacturing plants to attribute a large portion of the pharmaceutical residues found in a Swiss river to industrial emissions, further showing that curbing such pollution is an ongoing, worldwide challenge.CAS 
    PubMed 

    Google Scholar 
    148.Klümper, U. et al. Selection for antimicrobial resistance is reduced when embedded in a natural microbial community. ISME J. 13, 2927–2937 (2019).PubMed 
    PubMed Central 

    Google Scholar 
    149.Kraupner, N. et al. Selective concentrations for trimethoprim resistance in aquatic environments. Environ. Int. 144, 106083 (2020).CAS 
    PubMed 

    Google Scholar 
    150.Murray, A. K. et al. Novel insights into selection for antibiotic resistance in complex microbial communities. mBio https://doi.org/10.1128/mbio.00969-18 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    151.Government of India. Environment (Protection) Amendment Rules, 2020 – Inviting comments/suggestions on Environmental Standards for Bulk Drug and Formulation (Pharmaceutical) Industry, http://moef.gov.in/g-s-r-44-e-date-23-01-2020-environment-protection-amendment-rules-2020-inviting-commentssuggestions-on-environmental-standards-for-bulk-drug-and-formulation-pharmaceutical-indu/ (2020).152.Tell, J. et al. Science-based targets for antibiotics in receiving waters from pharmaceutical manufacturing operations. Integr. Environ. Assess. Manag. 15, 312–319 (2019).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    153.Greenfield, B. K. et al. Modeling the emergence of antibiotic resistance in the environment: an analytical solution for the minimum selection concentration. Antimicrob. Agents Chemother. https://doi.org/10.1128/aac.01686-17 (2018).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    154.Murray, A. K. et al. The ‘Selection end points in Communities of bacTeria’ (SELECT) method: a novel experimental assay to facilitate risk assessment of selection for antimicrobial resistance in the environment. Environ. Health Perspect. 128, 107007 (2020).PubMed 
    PubMed Central 

    Google Scholar 
    155.Andersson, D. I. & Hughes, D. Antibiotic resistance and its cost: is it possible to reverse resistance? Nat. Rev. Microbiol. 8, 260–271 (2010).CAS 
    PubMed 

    Google Scholar 
    156.Stanton, I. C., Murray, A. K., Zhang, L., Snape, J. & Gaze, W. H. Evolution of antibiotic resistance at low antibiotic concentrations including selection below the minimal selective concentration. Commun. Biol. https://doi.org/10.1038/s42003-020-01176-w (2020).Article 
    PubMed 
    PubMed Central 

    Google Scholar 
    157.Nijsingh, N., Munthe, C. & Larsson, D. G. J. Managing pollution from antibiotics manufacturing: charting actors, incentives and disincentives. Environ. Health 18, 95 (2019).PubMed 
    PubMed Central 

    Google Scholar 
    158.Sundin, G. W. & Wang, N. Antibiotic resistance in plant-pathogenic bacteria. Annu. Rev. Phytopathol. 56, 161–180 (2018).CAS 
    PubMed 

    Google Scholar 
    159.Government of Sweden. Uppdrag angående försöksverksamhet för en miljöpremie i läkemedelsförmånssystemet, https://www.regeringen.se/499677/contentassets/36dcec65be904fd58e5e6b01c2f99709/uppdrag-angaende-forsoksverksamhet-for-en-miljopremie-i-lakemedelsformanssystemet-tlv.pdf (2021).160.Norwegian Hospital Procurement Trust. New environmental criteria for the procurement of pharmaceuticals, https://sykehusinnkjop.no/nyheter/new-environmental-criteria-for-the-procurement-of-pharmaceuticals (2019).161.Swedish Procurement Agency. Pharmaceuticals, https://www.upphandlingsmyndigheten.se/kriterier/sjukvard-och-omsorg/lakemedel/ (2021).162.G7. G7 Health Ministers’ Declaration, Oxford, 4 June 2021, https://assets.publishing.service.gov.uk/government/uploads/system/uploads/attachment_data/file/992268/G7-health_ministers-communique-oxford-4-june-2021_5.pdf (2021).163.Årdal, C. et al. Supply chain transparency and the availability of essential medicines. Bull. World Health Organ. 99, 319–320 (2021).PubMed 
    PubMed Central 

    Google Scholar 
    164.Graham, D., Giesen, M. & Bunce, J. Strategic approach for prioritising local and regional sanitation interventions for reducing global antibiotic resistance. Water 11, 27 (2018).
    Google Scholar 
    165.Margot, J. et al. Treatment of micropollutants in municipal wastewater: ozone or powdered activated carbon? Sci. Total. Environ. 461–462, 480–498 (2013).PubMed 

    Google Scholar 
    166.Larsson, D. G. J. et al. Critical knowledge gaps and research needs related to the environmental dimensions of antibiotic resistance. Environ. Int. 117, 132–138 (2018).PubMed 

    Google Scholar 
    167.Laxminarayan, R. et al. The Lancet Infectious Diseases Commission on antimicrobial resistance: 6 years later. Lancet Infect. Dis. 20, e51–e60 (2020).PubMed 

    Google Scholar 
    168.Ahammad, Z. S., Sreekrishnan, T. R., Hands, C. L., Knapp, C. W. & Graham, D. W. Increased waterborne blaNDM-1 resistance gene abundances associated with seasonal human pilgrimages to the upper Ganges River. Environ. Sci. Technol. 48, 3014–3020 (2014).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    169.Kookana, R. S. et al. Potential ecological footprints of active pharmaceutical ingredients: an examination of risk factors in low-, middle- and high-income countries. Philos. Trans. R. Soc. B Biol. Sci. 369, 20130586 (2014).
    Google Scholar  More