More stories

  • in

    Identifying and characterizing pesticide use on 9,000 fields of organic agriculture

    We first identify the location of organic crop fields in Kern County and then estimate whether status as organic versus conventional fields determines pesticide use (Fig. 5).Fig. 5: Methodology overview.Figure outlines the main method steps from identifying organic fields to creating the analysis data to performing the statistical analyses. All images shown are simplified, visual representations of the datasets. CDFA refers to the California Department of Food and Agriculture, while APN is the Assessor’s Parcel Number and TRS is the Township-Range-Section. Identifying organic fields combines the created CDFA organic APN, CDFA organic TRS, and organic pesticides data layers together to create the final organic versus conventional fields layer used in the analysis data section. All analysis data layers are then inputted into the various statistical analyses.Full size imageIdentifying organic fieldsWe identified organic fields using a combination of California Department of Food and Agriculture (CDFA) records and Kern County Agricultural Commissioner’s Office spatial data (“fields shapefiles”) and pesticide use records. No single source was complete, and as such, we evaluated several different approaches to identifying organic fields.California Department of Food and Agriculture (CDFA) recordsData on the location of organic fields, per the California State Organic Program, for 2013–2019 was obtained by request from the California Department of Food and Agriculture (CDFA). The CDFA, through the State Organic Program, requires annual registration of certified organic producers who have an expected gross sale of over $5000. We were specifically interested in the pesticide aspects of organic production, which is governed in our study region by the USDA’s National List of Allowed and Prohibited Substances. The National List of Allowed and Prohibited Substances delineates which synthetic substances can be used and which natural substances cannot be used for pest control in US organic production. Besides substances specifically (dis)allowed on the National List, allowed substances include non-synthetic biological, botanical, and mineral inputs. Field location data were in the form of either Assessor’s Parcel Number (APN) or PLS System Township-Range-Section (TRS) values, though data were reported without systematic formatting. We harmonized the CDFA APN values to merge with the Kern County Assessor’s parcel shapefile (2017), which we then spatially joined with the Kern fields shapefiles. We followed a similar process with PLSS TRS values, which were then merged with the Kern County PLS Section shapefile, and joined to Kern field shapefiles. We refer to our final organic designation as “CDFA Organic”. Details of the data cleaning process are described in the Ancillary Data Processing Methods section below.Using pesticide use reports to refine organic field identificationAfter spot-checking pesticide use on CDFA Organic fields, it became clear we had not entirely eliminated conventional fields. This was likely due to variation in polygon geometries between PLSS Sections, Kern County Assessor parcels, and Kern agricultural fields data. To further refine our classification, we used field-level pesticide use, again from the Kern County Agricultural Commissioner’s Office. As thousands of pesticide products (active ingredients + adjuvants) are in use in Kern County, we took an iterative approach to eliminate fields using conventional pesticides. We first limited the universe of pesticides to those applied to fields that were CDFA Organic. We then identified the 50 most commonly used pesticide products by a number of applications, and manually classified each as organic or conventional. Having identified these products as described below, we matched them back in, eliminating fields that used conventional products and identifying as “PUR Organic” those that used only organic products. We repeated this process, hand identifying the most commonly used products and eliminating fields using conventional products until we had isolated fields using only organic products.To classify a product as organic or conventional, we first searched for each product’s U.S. EPA-registered product label, using the exact product name and EPA product registration number. If there was any indication on the label that the product was certified as organic by the Organic Materials Review Institute (OMRI), or said “for use in organic production” or “organic”, then the pesticide was identified as organic (n = 132). If there was no organic indication on the product label, we searched the OMRI certification database for products with identical names and manufacturers, and identified products as organic if such certifications existed (n = 39). If all ingredients were defined (i.e., no inert or undefined ingredients) and were known organic active ingredients, then the pesticide was identified as organic (n = 1) (Supplementary Data 1). We failed to find EPA-registered labels for three products and confirmed on the California Department of Pesticide Regulation website that they are either inactive or out of production (EPA registration numbers: 52467-50008-AA-5905, 36208-50020-AA, 2935-48-AA-120). Each of the three was rarely used (n  0) to be the same as the mechanisms determining the amount sprayed when some pesticides are used (pesticides when pesticides  > 0). Double-hurdle models64 are an alternative to the Tobit model that allows for the separation of these two decisions.The mechanisms underlying the two decisions (to spray, how much to spray if spraying) can differ such that different covariates can describe each process, and the same covariates are allowed to influence the two processes in different ways (i.e., sign and magnitude can differ). The first, binary decision is usually modeled with a probit model.$${{{{{rm{P}}}}}}left(y=0|{{{{{bf{x}}}}}}right)=1-Phi left({{{{{bf{x}}}}}}gammaright)$$
    (1)
    Then, the second decision is modeled as a linear model with pesticide use following a lognormal distribution, conditional on positive pesticide use64$$log (y)|{{{{{bf{x}}}}}},y , > , 0 sim {{{{{rm{Normal}}}}}}({{{{{bf{x}}}}}}{{{{{mathbf{upbeta }}}}}},{sigma }^{2})$$
    (2)
    where Φ is the standard normal cdf, x is a vector of explanatory variables including organic status, y is pesticide use, and ({{{{{mathbf{upbeta }}}}}}) is a vector of coefficients. We use a lognormal hurdle model rather than a truncated normal hurdle model since pesticide use is highly non-normal, and Q-Q plots suggested substantial model improvement using a lognormal rather than normal distribution. In contrast to the panel data models described in the Ancillary Statistical Methods below, our estimation equation used natural log-transformed variables for pesticides (and field and farm size) rather than inverse hyperbolic sine (IHS) transformation since only positive observations are included in the second hurdle model. Following insights derived from our panel data models (Supplementary Notes), we build on the basic hurdle model concept using the farm-by-crop family interaction as a random intercept in both the first and second hurdle. We chose the farm-by-crop family interaction rather than a crossed random effect due to computational feasibility with thousands of permits and hundreds of crops, due to similarity of results to the within estimator model (i.e., fixed effects in causal inference terminology; Supplementary Notes, Supplementary Table 2), and due to AIC/BIC (Supplementary Table 3). Further, we find evidence of heteroskedasticity from both visual inspection and Levine’s test, which adds additional complications to computing crossed random effects. Thus, we proceed with the farm-by-crop family interaction in a random intercept model with cluster robust standard errors clustered at the same grouping. In doing so, observations, where the taxonomic family of the crop was unclear, were dropped. Of the 7367 fields that were dropped due to missing crop families, 6684 were uncultivated agriculture.Our data are effectively repeated cross-sections rather than a true panel since fields are defined by the farm-site-year combination and thus generally change year-to-year or when crops rotate. We model it as such. This implies we do not require observations to have no spray in all time periods, as would be the case in a double hurdle panel model. Linking field IDs over time through spatial processing is complicated by crop rotations of different size areas. Since farmers may farm multiple fields under different management systems, as we illustrate here, and have different contractual obligations at a sub-farm level, requiring farms to never use pesticides on all fields is not reflective of on-the-ground decisions.We repeated the lognormal hurdle models individually for carrots, grapes, oranges, potatoes, and onions, which were the only widely-grown crops with more than 100 organic fields. This allowed for a different slope and intercept by crop type.We conduct several robustness checks. First, we do not have data on crop yields. However, to assess the potential implications of a yield gap on our results, we modify our per hectare rates following Ponisio et al.15 as a robustness check. We group commodities into cereals, roots and tubers, oilseeds, legumes/pulses, fruits, and vegetables and assign them the Ponisio et al.15 yield gap estimates for that group. Crops that did not fall into any of the above groups (e.g., cannabis) were provided the all-crop average from Ponisio et al.15. Second, we analyze how conventional and organic differ with respect to soil quality using a within estimator approach to account for crop-specific differences in soil quality. Third, binary toxicity metrics, while valuable given the number of chemicals and endpoints of interest here, nevertheless fail to distinguish gradations of toxicity for chemicals above (or below) the regulatory threshold. As mentioned above, the data needed to calculate many aggregate indices (e.g., Pesticide Load57 and Environmental Impact Quotient58) are not readily available for all of the chemicals in our study. For completeness, we attempted to calculate the Pesticide Toxicity Index for one well-studied endpoint, fish. We supplemented data provided in Nowell et al.41 with data from Standartox42. However, only about 70% of the chemicals used in our study matched, and pesticide products used on organic fields were more likely to lack toxicity information for one or more chemicals. We briefly discuss the highly preliminary investigation, given the non-random missing toxicity data.All spatial analyses were performed in R Statistical Software v 3.5.3 and all statistical analyses were performed Stata 16 MP. For all tests, statistical significance was based on two-tailed tests with (alpha =0.05.)Ancillary data processing methodsCleaning parcel dataTo spatially locate organic fields, we needed to match the Assessor’s parcel numbers (APNs) provided in the CDFA tabular data to APNs in the Kern County Parcel shapefile (from 2017). Over 90% of the APN entries in the CDFA data were in the format [xxx-xxx-xx], though multiple APNs were often provided in the same cell separated by line breaks, semi-colons, commas, and/or spaces. We made initial edits separating values into individual cells in Microsoft Excel since formatting was highly inconsistent. Observations whose APNs were not in the [xxx-xxx-xx] were modified so that their format matched. In the R environment, dashes were inserted after the third, sixth, and eighth characters (1234567895 became 123-456-78-95) for APNs that did not already contain them. Occasionally, APN numbers were provided with dashes, but with segments of incorrect length (e.g., 12-34-567). In these instances, APN segments were either trimmed from the right or padded with a zero on the left so they matched the [xxx-xxx-xx] format. This approach yielded the greatest number of matches and was checked for accuracy as described below. Additional segments (from APNs with more than two dashes and eight numeric characters) were dropped. A handful of APNs with fewer than eight numeric characters and no dashes were dropped entirely.The edited CDFA APNs were then joined with the Kern County Assessor’s parcel shapefile, creating the “CDFA organic shapefile”. In total, 1637 of 1829 individual CDFA records joined successfully. To evaluate the accuracy of joins between CDFA tabular data, Kern County parcel, and Kern County agricultural spatial data, we spot-checked ownership information using “Company” (CDFA) and “PERMITTEE” (Kern County agricultural data) values.To then identify the crop fields within the organic parcels, we performed a spatial join between the CDFA organic shapefile and the Kern County fields shapefiles. Prior to performing the join, the CDFA parcels’ dimensions were reduced with a 50-m buffer to eliminate spatial joins between CDFA parcels and crop fields that were only touching the parcel margins. Of five different buffer widths evaluated, 50 m reduced the number of false positives and negatives, as determined by comparing the “Company” and “PERMITTEE” values. We refer to the fields that match as “APN Organic”.Cleaning PLSS Township-Range-Section valuesEach year several producers reported Township, Section, and Range (TRS) values, consistent with the PLS System (PLSS), rather than APN values. We used these TRS values to identify PLSS Sections that contained organic fields.We separated any cell containing multiple TRS values and removed any prefixes such as “S”, “Section”, “Sec.”, “T”, and “R” that would prevent joining to Kern County PLSS spatial data in Excel. In the R environment, we padded the left side of the “S” value with a 0 if it was a single digit, then concatenated the three columns into a “TRS” column. We joined TRS from the CDFA tabular data to PLSS spatial data, which identified 563 Sections as containing organic fields, from 2013 to 2019, out of a total of 664 unique TRS codes in the CDFA dataset. We then performed a spatial join between PLSS Sections that contain organic fields and Kern County fields shapefiles, to identify all agriculture fields that overlap with those Sections. Additional processing using the Pesticide Use Reports is described above.Ancillary statistical methodsWe began with a pooled ordinary least squares (OLS) model that, as the name suggests, pools observations over farms, years, and crop types. However, there may be attributes of crops or farms that may be systematically different between organic and conventional, and this systematic difference could bias our pooled OLS results. To address this, we first considered propensity score approaches but were unable to find a sufficient balance of our covariate distribution between organic and conventional fields. As an alternative, we limited our sample to fields with overlapping farmers and crop types. In other words, we focused on the subset of fields that are grown by farmers producing both organic and conventional fields and to crops that are produced both conventionally and organically. However, this shrunk our dataset by two-thirds.To leverage more of our data, we next considered panel data models as a means to address unobserved variables. We consider both within-estimator models (also known as “fixed effects” in causal inference terminology, but different from the biostatistical use of the term) and random effects models (with random intercepts), seeking to capture characteristics of the crop, grower, and year. The advantage of a within-estimator approach is that the omitted variables are removed (through differencing) and thus, they can be correlated with covariates without biasing the estimation. In other words, pesticide use and all covariates are differenced from their crop-specific mean (or crop family, farmer, etc. specific mean, depending on the model). In doing so, the propensity for certain crops (crop family, farmer) to be grown organic or to be fast or slow adopters of new technologies is removed. The disadvantage is that characteristics shared by all fields of a crop (e.g., value) are lost in the differencing, and more importantly, that the differencing is not easily translated to nonlinear models that we employ later in the analysis. Random effects are more easily translated to nonlinear models. The disadvantage of random effects is the strong assumption that the unobserved variables are uncorrelated with the covariates18,65, which is required for random effects coefficient estimates to be unbiased. Here, we see the difference in coefficient estimates between the within-estimator and random effects models are quite small (Supplementary Table 2).Random effects particularly crossed random effects with thousands of permits and hundreds of crops, introduce computational challenges due to large, sparse matrices. Further, we find evidence of heteroskedasticity from both visual inspection and Levine’s test, which adds additional complications to computing crossed random effects. We proceed using the farm-by-crop family interaction in a random intercept model with cluster robust standard errors clustered at the same grouping based on AIC/BIC (Supplementary Table 3), computational feasibility, and similarity to the within-estimator results (Supplementary Table 2). Observations, where the taxonomic family of the crop was unclear, were dropped in any models including family in either the random effects or the cluster robust standard errors. Of the 7367 fields that were dropped due to missing crop families, 6684 were uncultivated agriculture.In the panel data models, we used IHS transformations to accommodate highly non-normal pesticide (and field and farm size) data. IHS is very similar to natural log transformation66 but is defined at zero, which is important given a sizable fraction of our observations have zero pesticide use. As with log–log transformations, IHS–IHS transformation can be interpreted as elasticities. We pre-multiply pesticide use by 100 to improve estimation66, though this does not affect interpretation. As described above, we leverage insights on model specification from the panel data models, but rely on the double hurdle models to parse apart the decision to spray from the decision of how much to spray.Reporting summaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

  • in

    Localised labyrinthine patterns in ecosystems

    The absence of the first principles for biological systems in general, and in particular for vegetation populations where phenomena are interconnected makes their mathematical modelling complex. The theory of vegetation pattern formation rests on the self-organisation hypothesis and symmetry-breaking instability that provoke the fragmentation of the uniform cover. The symmetry-breaking instability takes place even if the environment is isotropic31,33,35. This instability may be an advection-induced transition that requires the pre-existence of the environment anisotropy due to the topography of the landscape34,39,40. Generally speaking, this transition requires at least two feedback mechanisms having a short-range activation and a long-range inhibition. In this respect, we consider three different vegetation models that are experimentally relevant systems: (i) the generic interaction redistribution model describing vegetation pattern formation which incorporates explicitly the facilitation, competition and seed dispersion nonlocal interactions (ii) the local nonvariational partial differential model described by a nonvariational Swift–Hohenberg type of model equation, and (iii) the reaction–diffusion system that incorporate explicetely water transport.The interaction-redistribution approachThe integrodifferential modelThis approach consists of considering a well-known logistic equation with nonlocal plant-to-plant interactions. Three types of interactions are considered: the facilitative (M_{f}(mathbf {r},t)), the competitive (M_{c}(mathbf {r},t)), and the seed dispersion (M_{d}(mathbf {r},t)) nonlocal interactions. To simplify further the mathematical modelling, we consider that the seed dispersion obeys a diffusive process (M_{d}(mathbf {r},t)approx nabla ^{2}b(mathbf {r},t)), with D the diffusion coefficient, b the biomass density, and (nabla ^{2}=partial ^2/partial x^2+partial ^2/partial y^2) is the Laplace operator acting in the (x,y) plane. The interaction-redistribution reads$$begin{aligned} M_{i}=expleft{ frac{xi _{i}}{N_{i}}int b(mathbf {r}+mathbf {r}’,t)phi _i(r,t)dmathbf {r}’right} , { text{ with } } phi _i(r,t)= exp(-r/L_{i}) end{aligned}$$
    (1)
    where (i=f,c). (xi _i) represents the strength of the interaction, (N_i) is a normalisation constant. We assume that their Kernels (phi _i(r,t)) are exponential functions with (L_i) the range of their interactions. The facilitative interaction (M_{f}(mathbf {r},t)) favouring vegetation development. They involve the accumulation of nutrients in the neighbourhood of plants, the reciprocal sheltering of neighbouring plants against climatic harshness which improves the water budget in the soil. The range of the facilitative interaction (L_f) operates on the crown size. The competitive interaction operates over a length (L_c) and involves the below-ground structures, i.e., the rhizosphere. In nutrient-poor or/and in water-limited territories, lateral spreading may extend beyond the radius of the crown. This extension of roots relative to their crown size is necessary for the survival and the development of the plant in order to extract enough nutrients and/or water from the soil. When incorporating these nonlocal interactions in the paradigmatic logistic equation, the spatiotemporal evolution of the normalised biomass density (b(mathbf {r}, t)) in isotropic environmental conditions reads14$$begin{aligned} partial _{t} b(mathbf {r},t)=b(mathbf {r},t)[1-b(mathbf {r},t)]M_{f}(mathbf {r},t)- mu b(mathbf {r},t)M_{c}(mathbf {r},t)+Dnabla ^{2}b(mathbf {r},t). end{aligned}$$
    (2)
    The normalisation is performed with respect to the total amount of biomass supported by the system. The first two terms in the logistic equation with nonlocal interaction Eq. (2) describe the biomass gains and losses, respectively. The third term models seed dispersion. The aridity parameter (mu) accounts for the biomass loss and gain ratio, which depends on water availability and nutrients soil distribution, topography, etc. The homogeneous cover solutions of Eq. (2) are: (b_{o}=0) which corresponds to the state totally devoid of vegetation, and the homogeneous cover solutions satisfy the equation$$begin{aligned} mu =(1-b)exp (Delta b), end{aligned}$$
    (3)
    with (Delta =xi _{f}-xi _{c}) measures the community cooperativity if (Delta >0) or anti-cooperativity when (Delta 0). The solution (u_{-}) is always unstable even in the presence of small spatial fluctuations. The linear stability analysis of vegetated cover ((u_{+})) with respect to small spatial fluctuations, yields the dispersion relation$$begin{aligned} sigma (k)=u_{+}(kappa -2u_{+})-(nu -gamma u_{+})k^{2}-alpha u_{+}k^{4}. end{aligned}$$
    (8)
    Imposing (partial sigma /partial k|_{k_{c}}=0) and (sigma (k_{c})=0), the critical mode can be determined$$begin{aligned} k_{c}=sqrt{frac{gamma -nu /u_{c}}{2alpha }}, end{aligned}$$
    (9)
    where (u_{c}) satisfies (4alpha u_{c}^2(2u_{c}-kappa )=(2gamma u_{c}-nu )^2). The corresponding aridity parameter (eta _{c}) can be calculated from Eq. (7).The reaction–diffusion approachThe second approach explicitly adds the water transport by below ground diffusion. The coupling between the water dynamics and the plant biomass involves positive feedbacks that tend to enhance water availability. Negative feedbacks allow for an increase in water consumption caused by vegetation growth, which inhibits further biomass growth.The modelling considers the coupled evolution of biomass density (b(mathbf {r},t)) and groundwater density (w(mathbf {r},t)). In its dimensionless form, this model reads33$$begin{aligned} frac{partial b}{partial t}= & {} frac{gamma w}{1+omega w}b-b^{2}-theta b+nabla ^{2}b, end{aligned}$$
    (10)
    $$begin{aligned} frac{partial w}{partial t}= & {} p-(1-rho b)w-w^{2}b+delta nabla ^{2}(w-beta b). end{aligned}$$
    (11)
    The first term in the first equation describes plant growth at a constant rate ((gamma /omega)) that grows linearly with w for dry soil. The quadratic nonlinearity (-b^{2}) accounts for saturation imposed by poor nutrients soil. The term proportional to (theta) accounts for mortality, grazing or herbivores. The mechanisms of dispersion are modelled by a simple diffusion process. The groundwater evolves due to a precipitation input p. The term ((1-rho b)w) in the second equation accounts for the evaporation and drainage, that decreases with the presence of vegetation. The term (w^{2}b) models the water uptake by the plants due to the transpiration process. The groundwater movement follows the Darcy’s law in unsaturated conditions; that is, the water flux is proportional to the gradient of the water matric potential41. The matric potential is equal to w, under the assumption that the hydraulic diffusivity is constant41. To model the suction of water by the roots, a correction to the matric potential is included; (-beta b), where (beta) is the strength of the suction. More

  • in

    Past environmental changes affected lemur population dynamics prior to human impact in Madagascar

    1.Frankham, R., Briscoe, D. A. & Ballou, J. D. Introduction to conservation genetics (Cambridge university press, 2002).2.Nadachowska-Brzyska, K., Burri, R., Smeds, L. & Ellegren, H. PSMC analysis of effective population sizes in molecular ecology and its application to black-and-white Ficedula flycatchers. Mol. Ecol. 25, 1058–1072 (2016).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    3.Martínez-Freiría, F., Velo-Antón, G. & Brito, J. C. Trapped by climate: Interglacial refuge and recent population expansion in the endemic Iberian adder Vipera seoanei. Divers. Distrib. 21, 331–344 (2015).Article 

    Google Scholar 
    4.Martínez-Freiría, F. et al. Integrative phylogeographical and ecological analysis reveals multiple pleistocene refugia for Mediterranean Daboia vipers in north-west Africa. Biol. J. Linn. Soc. 122, 366–384 (2017).Article 

    Google Scholar 
    5.Veríssimo, J. et al. Pleistocene diversification in Morocco and recent demographic expansion in the Mediterranean pond turtle Mauremys leprosa. Biol. J. Linn. Soc. 119, 943–959 (2016).Article 

    Google Scholar 
    6.Chattopadhyay, B., Garg, K. M., Gwee, C. Y., Edwards, S. V. & Rheindt, F. E. Gene flow during glacial habitat shifts facilitates character displacement in a Neotropical flycatcher radiation. BMC Evol. Biol. 17, 1–15 (2017).Article 

    Google Scholar 
    7.Garg, K. M., Chattopadhyay, B., Koane, B., Sam, K. & Rheindt, F. E. Last Glacial Maximum led to community-wide population expansion in a montane songbird radiation in highland Papua New Guinea. BMC Evol. Biol. 20, 82 (2020).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    8.Vences, M., Wollenberg, K. C., Vieites, D. R. & Lees, D. C. Madagascar as a model region of species diversification. Trends Ecol. Evol. 24, 456–465 (2009).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    9.Goodman, S. M., Raherilalao, M. J. & Wohlhauser, S. The Terrestrial Protected Areas of Madagascar: Their History, Description and Biota (Association Vahatra in Antananarivo, The University of Chicago Press, 2018).10.Douglass, K. The diversity of late holocene shellfish exploitation in Velondriake, Southwest Madagascar. J. Island Coast. Archaeol. 12, 333–359 (2016).11.Yoder, A. D., Campbell, C. R., Blanco, M. B., Ganzhorn, J. U. & Goodman, S. M. Geogenetic patterns in mouse lemurs (genus Microcebus) reveal the ghosts of Madagascar’s forests past. PNAS 113, 8049–8056 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    12.Salmona, J., Heller, R., Quéméré, E. & Chikhi, L., Climate change. and human colonization triggered habitat loss and fragmentation in Madagascar. Mol. Ecol. 26, 5203–5222 (2017).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    13.Townsend, T. M., Vieites, D. R., Glaw, F. & Vences, M. Testing species-level diversification hypotheses in Madagascar: the case of microendemic Brookesia leaf Chameleons. Syst. Biol. 58, 641–656 (2009).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    14.Brown, J. L., Cameron, A., Yoder, A. D. & Vences, M. A necessarily complex model to explain the biogeography of the amphibians and reptiles of Madagascar. Nat. Commun. 5, 5046 (2014).15.Schüßler, D. et al. Ecology and morphology of mouse lemurs (Microcebus spp.) in a hotspot of microendemism in northeastern Madagascar, with the description of a new species. Am. J. Primatol. 82, e23180 (2020).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    16.Chikhi, L. & Bruford, M. Mammalian population genetics and genomics. Mamm. Genome https://doi.org/10.1079/9780851999104.0539 (2005).17.Olivieri, G. L., Sousa, V., Chikhi, L. & Radespiel, U. From genetic diversity and structure to conservation: Genetic signature of recent population declines in three mouse lemur species (Microcebus spp.). Biol. Conserv. 141, 1257–1271 (2008).Article 

    Google Scholar 
    18.Gutenkunst, R. N., Hernandez, R. D., Williamson, S. H. & Bustamante, C. D. Inferring the joint demographic history of multiple populations from multidimensional SNP frequency data. PLoS Genet. 5, e1000695 (2009).19.Li, H. & Durbin, R. Inference of human population history from individual whole-genome sequences. Nature 475, 493–496 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    20.Excoffier, L., Dupanloup, I., Huerta-Sánchez, E., Sousa, V. C. & Foll, M. Robust demographic inference from genomic and SNP data. PLoS Genet. 9, e1003905 (2013).21.Liu, X. & Fu, Y.-X. Exploring population size changes using SNP frequency spectra. Nat Genet. 47, 555–559 (2015).22.Salmona, J., Heller, R., Lascoux, M. & Shafer, A. Inferring demographic history using genomic data. in Population Genomics 511–537 (Springer, 2017).23.Beichman, A. C., Huerta-Sanchez, E. & Lohmueller, K. E. Using genomic data to infer historic population dynamics of nonmodel organisms. Annu. Rev. Ecol. Evol. Syst. 49, 433–456 (2018).Article 

    Google Scholar 
    24.Sgarlata, G. M. et al. Genetic and morphological diversity of mouse lemurs (Microcebus spp.) in northern Madagascar: The discovery of a putative new species? Am. J. Primatol. 81, e23070 (2019).25.Demenocal, P. et al. Abrupt onset and termination of the African humid period:: rapid climate responses to gradual insolation forcing. Quat. Sci. Rev. 19, 347–361 (2000).Article 

    Google Scholar 
    26.Tierney, J. E. & DeMenocal, P. B. Abrupt shifts in Horn of Africa hydroclimate since the last glacial maximum. Science 342, 843–846 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    27.Los, S. O. et al. Sensitivity of a tropical montane cloud forest to climate change, present, past and future: Mt. Marsabit, N. Kenya. Quat. Sci. Rev. 218, 34–48 (2019).Article 

    Google Scholar 
    28.Ivory, S. J. & Russell, J. Climate, herbivory, and fire controls on tropical African forest for the last 60ka. Quat. Sci. Rev. 148, 101–114 (2016).Article 

    Google Scholar 
    29.Conroy, J. L., Overpeck, J. T., Cole, J. E., Shanahan, T. M. & Steinitz-Kannan, M. Holocene changes in eastern tropical Pacific climate inferred from a Galápagos lake sediment record. Quat. Sci. Rev. 27, 1166–1180 (2008).Article 

    Google Scholar 
    30.Martin-Puertas, C., Tjallingii, R., Bloemsma, M. & Brauer, A. Varved sediment responses to early Holocene climate and environmental changes in Lake Meerfelder Maar (Germany) obtained from multivariate analyses of micro X-ray fluorescence core scanning data. J. Quat. Sci. 32, 427–436 (2017).Article 

    Google Scholar 
    31.Flenley, J. R. Tropical forests under the climates of the last 30,000 years. in Potential Impacts of Climate Change on Tropical Forest Ecosystems, 37–57 (Springer, 1998).32.Burrough, S. L. & Thomas, D. S. G. Central southern Africa at the time of the African humid period: a new analysis of Holocene palaeoenvironmental and palaeoclimate data. Quat. Sci. Rev. 80, 29–46 (2013).Article 

    Google Scholar 
    33.Ivory, S. J. & Russell, J. Lowland forest collapse and early human impacts at the end of the African humid period at Lake Edward, equatorial East. Afr. Quat. Res. 89, 7–20 (2018).Article 

    Google Scholar 
    34.Anderson, A. et al. New evidence of megafaunal bone damage indicates late colonization of Madagascar. PLoS ONE 13, 1–14 (2018).
    Google Scholar 
    35.Hansford, J. et al. Early Holocene human presence in Madagascar evidenced by exploitation of avian megafauna. Sci. Adv. 4, eaat6925 (2018).36.Burney, D. A., Robinson, G. S. & Burney, L. P. Sporormiella and the late holocene extinctions in Madagascar. Proc. Natl Acad. Sci. USA 100, 10800–10805 (2003).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    37.Railsback, L. B. et al. Relationships between climate change, human environmental impact, and megafaunal extinction inferred from a 4000-year multi-proxy record from a stalagmite from northwestern Madagascar. Quat. Sci. Rev. 234, 106244 (2020).Article 

    Google Scholar 
    38.Dewar, R. E. et al. Stone tools and foraging in northern Madagascar challenge Holocene extinction models. PNAS 110, 12583–12588 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    39.Radimilahy, C. Mahilaka: an Archaeological Investigation of an Early Town in Northwestern Madagascar. Acta Universitatis Upsaliensis (University of Uppsala, 1998).40.Liu, X. & Fu, Y.-X. Exploring population size changes using SNP frequency spectra. Nat. Genet 47, 555–559 (2015).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    41.Lapierre, M., Lambert, A. & Achaz, G. Accuracy of demographic inferences from the site frequency spectrum: the case of the yoruba population. Genetics 206, 139–449 (2017).Article 

    Google Scholar 
    42.Terhorst, J., Kamm, J. A. & Song, Y. S. Robust and scalable inference of population history from hundreds of unphased whole genomes. Nat. Genet. 49, 303–309 (2017).CAS 
    Article 

    Google Scholar 
    43.Patton, A. H. et al. Contemporary demographic reconstruction methods are robust to genome assembly quality: a case study in Tasmanian devils. Mol. Biol. Evol. 36, 2906–2921 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    44.Mazet, O., Rodríguez, W., Grusea, S., Boitard, S. & Chikhi, L. On the importance of being structured: Instantaneous coalescence rates and human evolution-lessons for ancestral population size inference? Heredity 116, 362–371 (2016).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    45.Orozco-terWengel, P. The devil is in the details: the effect of population structure on demographic inference. Heredity 116, 349–350 (2016).46.Mazet, O., Rodríguez, W. & Chikhi, L. Demographic inference using genetic data from a single individual: separating population size variation from population structure. Theor. Popul. Biol. 104, 46–58 (2015).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    47.Chikhi, L. et al. The IICR (inverse instantaneous coalescence rate) as a summary of genomic diversity: Insights into demographic inference and model choice. Heredity 120, 13–24 (2018).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    48.Simons, E. L., Godfrey, L. R., Vuillaume-Randriamanantena, M., Chatrath, P. S. & Gagnon, M. Discovery of new giant subfossil lemurs in the Ankarana Mountains of Northern Madagascar. J. Hum. Evol. 19, 311–319 (1990).Article 

    Google Scholar 
    49.Jungers, W. L., Godfrey, L. R., Simons, E. L. & Chatrath, P. S. Subfossil Indri indri from the Ankarana Massif of northern Madagascar. Am. J. Phys. Anthropol. 97, 357–366 (1995).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    50.Wilson, J. M., Stewart, P. D. & Fowler, S. V. Ankarana — a rediscovered nature reserve in northern Madagascar. Oryx 22, 163–171 (1988).Article 

    Google Scholar 
    51.Everson, K. M., Jansa, S. A., Goodman, S. M. & Olson, L. E. Montane regions shape patterns of diversification in small mammals and reptiles from Madagascar’s moist evergreen forest. J. Biogeogr. 47, 2059–2072 (2020).Article 

    Google Scholar 
    52.Douglass, K., Hixon, S., Wright, H. T., Godfrey, L. R. & Crowley, B. E. A critical review of radiocarbon dates clarifies the human settlement of Madagascar. Quat. Sci. Rev. 221, 105878 (2019).53.Orozco-Terwengel, P., Andreone, F., Louis, E. & Vences, M. Mitochondrial introgressive hybridization following a demographic expansion in the tomato frogs of Madagascar, genus. Dyscophus. Mol. Ecol. 22, 6074–6090 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    54.Johnson, J. A. et al. Long-term survival despite low genetic diversity in the critically endangered Madagascar fish-eagle. Mol. Ecol. 18, 54–63 (2009).PubMed 
    PubMed Central 

    Google Scholar 
    55.Sommer, S. Effects of habitat fragmentation and changes of dispersal behaviour after a recent population decline on the genetic variability of noncoding and coding DNA of a monogamous Malagasy rodent. Mol. Ecol. 12, 2845–2851 (2003).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    56.Craul, M. et al. Influence of forest fragmentation on an endangered large-bodied lemur in northwestern Madagascar. Biol. Conserv. 142, 2862–2871 (2009).Article 

    Google Scholar 
    57.Parga, J. A., Sauther, M. L., Cuozzo, F. P., Jacky, I. A. Y. & Lawler, R. R. Evaluating ring-tailed lemurs (Lemur catta) from southwestern Madagascar for a genetic population bottleneck. Am. J. Phys. Anthropol. 147, 21–29 (2012).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    58.Dewar, R. E. et al. Stone tools and foraging in northern Madagascar challenge Holocene extinction models. Proc. Natl Acad. Sci. USA 110, 12583–12588 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    59.Schüler, L. & Hemp, A. Atlas of pollen and spores and their parent taxa of Mt. Kilimanjaro and tropical East Africa. Quat. Int. 425, 301–386 (2016).Article 

    Google Scholar 
    60.Du Puy, D. J. & Moat, J. Vegetation mapping and classification in Madagascar (using GIS): implications and recommendations for the conservation of biodiversity. in Chorology, Taxonomy and Ecology of the floras of Africa and Madagascar, 97–117 (1998, in press).61.Guillaumet, J.-L., Betsch, J.-M. & Callmander, M. W. Renaud Paulian et le programme du CNRS sur les hautes montagnes à Madagascar: étage vs domaine. Zoosystema 30, 723 (2008).PubMed 
    PubMed Central 

    Google Scholar 
    62.Weisrock, D. W. et al. Delimiting species without nuclear monophyly in Madagascar’s mouse lemurs. PLoS ONE 5, e9883 (2010).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    63.Croudace, I. W., Rindby, A. & Rothwell, R. G. ITRAX: description and evaluation of a new multi-function X-ray core scanner. Geol. Soc. Lond. Spec. Publ. 267, 51–63 (2006).CAS 
    Article 

    Google Scholar 
    64.Blott, S. J. & Pye, K. GRADISTAT: a grain size distribution and statistics package for the analysis of unconsolidated sediments. Earth Surf. Process. Landforms 26, 1237–1248 (2001).Article 

    Google Scholar 
    65.Hogg, A. G. et al. SHCal13 Southern Hemisphere calibration, 0–50,000 years cal BP. Radiocarbon 55, 1889–1903 (2013).CAS 
    Article 

    Google Scholar 
    66.Rina Evasoa, M. et al. Sources of variation in social tolerance in mouse lemurs (Microcebus spp.). BMC Ecol. 19, 1–16 (2019).CAS 
    Article 

    Google Scholar 
    67.Aleixo-Pais, I. et al. The genetic structure of a mouse lemur living in a fragmented habitat in Northern Madagascar. Conserv. Genet. 20, 229–243 (2019).Article 

    Google Scholar 
    68.Radespiel, U., Jurić, M. & Zimmermann, E. Sociogenetic structures, dispersal and the risk of inbreeding in a small nocturnal lemur, the golden-brown mouse lemur (Microcebus ravelobensis). Behaviour 146, 607–628 (2009).Article 

    Google Scholar 
    69.Radespiel, U., Ehresmann, P. & Zimmermann, E. Species-specific usage of sleeping sites in two sympatric mouse lemur species (Microcebus murinus and M. ravelobensis) in northwestern Madagascar. Am. J. Primatol. 59, 139–151 (2003).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    70.Larsen, P. A. et al. Hybrid de novo genome assembly and centromere characterization of the gray mouse lemur (Microcebus murinus). BMC Biol. 15, 1–17 (2017).Article 
    CAS 

    Google Scholar 
    71.Metzker, M. L. Sequencing technologies — the next generation. Nat. Rev. Genet. 11, 31 (2010).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    72.Skotte, L., Korneliussen, T. S. & Albrechtsen, A. Estimating individual admixture proportions from next generation sequencing. Data 195, 693–702 (2013).CAS 

    Google Scholar 
    73.Korneliussen, T. S., Albrechtsen, A. & Nielsen, R. ANGSD: analysis of next generation sequencing data. BMC Bioinforma. 15, 1–13 (2014).Article 

    Google Scholar 
    74.Korneliussen, T. S. & Moltke, I. Sequence analysis NgsRelate: a software tool for estimating pairwise relatedness from next-generation sequencing data. Bioinformatics 31, 4009–4011 (2015).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    75.Soraggi, S., Wiuf, C. & Albrechtsen, A. Powerful inference with the D-Statistic on low-coverage whole-genome data. G3 8, 551–566 (2017).PubMed Central 
    Article 

    Google Scholar 
    76.Chikhi, L., Sousa, V. C., Luisi, P., Goossens, B. & Beaumont, M. A. The confounding effects of population structure, genetic diversity and the sampling scheme on the detection and quantification of population size changes. Genetics 186, 983–995 (2010).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    77.Kopelman, N. M., Mayzel, J., Jakobsson, M., Rosenberg, N. A. & Mayrose, I. Clumpak: a program for identifying clustering modes and packaging population structure inferences across K. Mol. Ecol. Resour. 15, 1179–1191 (2015).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    78.Evanno, G., Regnaut, S. & Goudet, J. Detecting the number of clusters of individuals using the software STRUCTURE: a simulation study. Mol. Ecol. 14, 2611–2620 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    79.Salmona, J., Heller, R., Quéméré, E. & Chikhi, L. Climate change and human colonization triggered habitat loss and fragmentation in Madagascar. Mol. Ecol. 26, 5203–5222 (2017).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    80.Schneider, N., Chikhi, L., Currat, M. & Radespiel, U. Signals of recent spatial expansions in the grey mouse lemur (Microcebus murinus). BMC Evol. Biol. 10, 105 (2010).81.Radespiel, U., Lutermann, H., Schmelting, B. & Zimmermann, E. An empirical estimate of the generation time of mouse lemurs. Am. J. Primatol. 81, 1–8 (2019).Article 

    Google Scholar 
    82.Hawkins, M. T. R. et al. Genome sequence and population declines in the critically endangered greater bamboo lemur (Prolemur simus) and implications for conservation. BMC Genomics 19, 1–15 (2018).Article 
    CAS 

    Google Scholar 
    83.Poelstra, J. et al. Cryptic patterns of speciation in cryptic primates: microendemic mouse lemurs and the multispecies coalescent. Syst. Biol. https://doi.org/10.1093/sysbio/syaa053 (2020).84.Campbell, C. R. et al. Pedigree-based and phylogenetic methods support surprising patterns of mutation rate and spectrum in the gray mouse lemur. Heredity 127.2, 233–244 (2021).Article 

    Google Scholar 
    85.Hudson, R. R. Generating samples under a Wright–Fisher neutral model of genetic variation. Bioinformatics 18, 337–338 (2002).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    86.Fredsted, T., Pertoldi, C., Schierup, M. H. & Kappeler, P. M. Microsatellite analyses reveal fine-scale genetic structure in grey mouse lemurs (Microcebus murinus). Mol. Ecol. 14, 2363–2372 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    87.Radespiel, U., Schulte, J., Burke, R. J. & Lehman, S. M. Molecular edge effects in the endangered golden-brown mouse lemur Microcebus ravelobensis. Oryx 53, 716–726 (2019).Article 

    Google Scholar 
    88.Radespiel, U., Lutermann, H., Schmelting, B., Bruford, M. W. & Zimmermann, E. Patterns and dynamics of sex-biased dispersal in a nocturnal primate, the grey mouse lemur, Microcebus murinus. Anim. Behav. 65, 709–719 (2003).Article 

    Google Scholar 
    89.Radespiel, U., Rakotondravony, R. & Chikhi, L. Natural and anthropogenic determinants of genetic structure in the largest remaining population of the endangered golden-brown mouse lemur, Microcebus ravelobensis. Am. J. Primatol. 70, 860–870 (2008).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    90.Schliehe-Diecks, S., Eberle, M. & Kappeler, P. M. Walk the line-dispersal movements of gray mouse lemurs (Microcebus murinus). Behav. Ecol. Sociobiol. 66, 1175–1185 (2012).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    91.Excoffier, L., Dupanloup, I., Huerta-Sánchez, E., Sousa, V. C. & Foll, M. Robust demographic inference from genomic and SNP data. PLoS Genet. 9, e1003905 (2013).92.Beerli, P. Effect of unsampled populations on the estimation of population sizes and migration rates between sampled populations. Mol. Ecol. 13, 827–836 (2004).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    93.Akaike, H. A new look at the statistical model identification. IEEE Trans. Automat. Control 19, 716–723 (1974).Article 

    Google Scholar 
    94.Bagley, R. K., Sousa, V. C., Niemiller, M. L. & Linnen, C. R. History, geography and host use shape genomewide patterns of genetic variation in the redheaded pine sawfly (Neodiprion lecontei). Mol. Ecol. 26, 1022–1044 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar  More

  • in

    The use of multi-criteria method in the process of threat assessment to the environment

    The research was carried out on the basis of direct measurements in the surroundings of four selected working coal-fired power plants and four working coking plants. The samples of suspended dust PM10, respirable fraction PM2.5 and submicron particulate matter PM1 were collected in the surroundings of power generation facilities and in the surroundings of coking plants.Location of measurement pointsThe location of the measurement points was selected in southern Poland, around the selected four working coal-fired power plants and four working coking plants. The sampling points in the surroundings of the power plant (P1, P2, P3 and P4) and the coking plant (K1, K2, K3 and K4) were located at the distance of approximately 2 km to the north-east from the respective object (Fig. 1).Figure 1Location of the sampling sites (the map was generated based on data from the BDL18 website).Full size imageThe location of the measurement points was a compromise, taking into account the representativeness of the receptor, the possibility to connect the testing equipment and the consent of the property owners. To eliminate the impact of a heating season, and especially that of low emissions, presented in the studies by19, the measurement sessions were carried out only in the summer season. The samples of particulate matter were collected on a weekly basis, with 4 sessions at one site. The methodology applied in this work is presented in20,21. The location of measurement sites:

    point P1: 50° 08′ 37.87″ N; 18° 32′ 15.76″ (Golejów—a suburban district of Rybnik in the Śląskie Voivodeship, in the vicinity of a working power plant with a capacity of 1775 MW; population:

    2 300);

    point P2: 50° 45′ 35.41″ N; 17° 56′ 20.43″ E (Świerkle—a rural area in the Opolskie Voivodeship (Dobrzeń Wielki commune) near a working power plant with a capacity of 1,492 MW; population: 520);

    point P3: 50° 12′ 33.46″ N; 19° 28′ 28.77″ E (Czyżówka—rural area in the Małopolskie Voivodeship (commune of Trzebinia) near a working power plant with a capacity of 786 MW; population: 700);

    point P4: 50° 13′ 48.90″ N; 19° 13′ 24.45″ E (suburbs of Jaworzno (Śląskie Voivodeship) in the vicinity of a 1,345 MW power plant; number of inhabitants: 95 500);

    K1 point: 50° 10′ 11.36″ N; 18° 40′ 34.35″ E (Czerwionka—Leszczyny in the Śląskie Voivodeship, in the vicinity of a small coking plant; number of inhabitants: 27 300);

    K2 point: 50° 3′ 19.76″ N; 18° 30′ 21.69″ E (Popielów—a suburban district of Rybnik in the Śląskie Voivodeship, surrounded by a small working coking plant; population:3 300);

    K3 point: 50° 21′ 24.08″ N; 19° 21′ 37.46″ E (Łęka—Dąbrowa Górnicza district, in the Śląskie Voivodeship, surrounded by a large coking plant; number of inhabitants: 700);

    K4 point: 50° 21′ 0.47″ N; 18° 53′ 15.44″ E (Bytom—a city in the Śląskie Voivodeship, a small coking plant located on the outskirts of the city; population: 174 700).

    The state of air pollution with particulate matter in the area investigated in the study is affected by various local sources of pollution emissions. At the measurement sites P1, P2, P3 and P4, the emissions are mainly from power plant chimneys, but also from auxiliary processes, i.e. coal storage and its transport. In addition, the recorded emissions are also influenced by other industrial plants operating in the vicinity of the measurement sites, domestic and municipal sector and the impact of automotive industry. The measurement sites K1, K2, K3 and K4 involve primarily the emissions accompanying the processes of coal coking as well as auxiliary processes, i.e. coal deposition, its transmission, management of products and post-production wastes. Additionally, they are affected by the emissions from industrial plants and low emission sources operating in this area, as well as the emission from the combustion of solid fuels for domestic or municipal purposes, as well as by the automotive industry.Sampling processThe samples of suspended dust (PM10), respirable fraction (PM2.5) and submicron particulate matter (PM1) were collected using the Dekati PM10 cascade impactor serial No. 6648 by Dekati (Finland) with the air flow rate of (1.8 {mathrm{m}}^{3}/mathrm{h}). The impactor Dekati PM10 guarantees the collection of dust samples for three cutpoint diameters: 10 μm, 2.5 μm and 1 μm. For the sampling at the first, second and third stages of the impactor, polycarbonate filters were used (Nuclepore 800 203, with the diameter of 25 mm, by Whatman International Ltd., Maidstone, UK). At the fourth stage, the dust was collected on a Teflon filter for particles ≤ 1 μm in diameter (Pall Teflo R2PJ047, 47 mm in diameter, by Pall International Ltd., New York, NY, USA). The average volume of air passing through the filters was approximately 300 m3. The impactor’s capture efficiency was characterized by the uncertainty below 2.8%. The mass of dust collected at the individual stages of the impactor was determined by the gravimetric method, and it was referenced to the volume of passed air (left(mathrm{mu g}/{mathrm{m}}^{3}right)) according to the PN-EN1234122. All impactor samples were analysed by inductively coupled plasma mass spectrometry (ICP-MS).The samples were collected at a height of 1.5 m from the ground, i.e. in the breathing zone for people. The respective dust fractions were collected in 7-day cycles from 28 May to 24 September 2014 (16 weeks) in the surroundings of four working coal-fired power plants and from 4 May to 28 August 2015 (16 weeks) in the surroundings of four working coking plants. The measurement campaign comprised four measurement sessions separately for each sampling site. One session comprised dust sampling at each stage of the Dekati PM10 cascade impactor and filters used for reference. The filters were taken back after study period and labeled during the collection process in the field and stored in the plastic containers for safe transportation and storage in laboratory for further analysis.In each measurement session, blind filters were stored at the sampling site, but they were not subjected to exposure. The sample data were corrected from these blanks. The length of the measurement cycles was conditioned by the need to collect an appropriate amount of research material (with the aerodynamic diameter of the dust grains  10 μm). Analogous (7-day) periods of dust sampling were used in the studies by4,23.Polycarbonate and Teflon filters were conditioned before and after dust collection at a temperature of 20 ± 1 °C (relative humidity 50%(pm ) 5%) for 48 h, and then weighed on a microbalance with an accuracy of 1 (mathrm{mu g}) (MXA5/1, by RADWAG, Poland).Taking into account the measurement sessions at four sites in the surroundings of the power plant (P1 (div) P4) and at four sites in the surroundings of the coking plant (K1 (div) K4), the aggregate number of samples exceeded 450.Chemical analysisThe qualitative and quantitative analysis of the obtained solutions was performed by inductively coupled plasma mass spectrometry using an ICP-MS instrument (NexION 300D, PerkinElmer, Inc., Waltham, MA, USA). For all elements determined simultaneously, the same parameters of the instrument were used, which are presented in the publications20,21,24.As standards for the determination of 75As, 111Cd, 59Co, 53Cr, 200Hg, 55Mn, 60Ni, 206Pb, 121Sb and 82Se, we applied the 1000 (mathrm{mu g}/{mathrm{cm}}^{3}) CertPUR ICP multi-element standard solution VI for ICP-MS by Merck, Germany. Ten repetitions were performed for all samples. The determined limits of detection (LOD) were based on 10 independent measurements for blank test. For the results obtained in that way, the mean value and the value of the standard deviation SD were calculated. The values of LOD for individual elements were determined on the basis of the dependence (1):$$mathrm{LOD}= {mathrm{x}}_{mathrm{sr}}+ 3mathrm{SD}$$
    (1)

    where: xśr—mean concentration value of the element, (mathrm{g}/{mathrm{dm}}^{3}), SD—standard deviation.The determination correctness of the content of the elements was verified with the use of certified reference materials: European Reference Material ERM-CZ120 and Standard Reference Material SRM 1648a (National Institute of Standards and Technology, USA). The recovery with the use of the said certified reference materials was respectively as follows: As (111% for ERM-CZ120 and 96% for SRM 1648a), Cd (97% and 105%), Co (108% and 97%), Cr (103% and 94%), Mn (106% and 100%), Ni (107% and 102%), Pb (107% and 105%) and Sb (99% and 91%). The certified reference materials did not contain Hg or Se. More

  • in

    Towards an integrative view of virus phenotypes

    1.Suttle, C. A. Marine viruses — major players in the global ecosystem. Nat. Rev. Microbiol. 5, 801–812 (2007).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    2.Rohwer, F. & Thurber, R. V. Viruses manipulate the marine environment. Nature 459, 207–212 (2009).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    3.Simmonds, P. et al. Virus taxonomy in the age of metagenomics. Nat. Rev. Microbiol. 15, 161–168 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    4.Fuhrman, J. A. Marine viruses and their biogeochemical and ecological effects. Nature 399, 541–548 (1999).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    5.Suttle, C. A. Viruses in the sea. Nature 437, 356–361 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    6.Jiang, S., Steward, G., Jellison, R., Chu, W. & Choi, S. Abundance, distribution, and diversity of viruses in alkaline, hypersaline Mono Lake, California. Microb. Ecol. 47, 9–17 (2004).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    7.Williamson, K. E., Fuhrmann, J. J., Wommack, K. E. & Radosevich, M. Viruses in soil ecosystems: an unknown quantity within an unexplored territory. Annu. Rev. Virol. 4, 201–219 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    8.Cai, L. et al. Active and diverse viruses persist in the deep sub-seafloor sediments over thousands of years. ISME J. 13, 1857–1864 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    9.Wei, M. & Xu, K. New insights into the virus-to-prokaryote ratio (VPR) in marine sediments. Front. Microbiol. 11, 1102 (2020).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    10.Wilhelm, S. W. & Suttle, C. A. Viruses and nutrient cycles in the sea. BioScience 49, 781–788 (1999).Article 

    Google Scholar 
    11.Brussaard, C. P. D. et al. Global-scale processes with a nanoscale drive: the role of marine viruses. ISME J. 2, 575–578 (2008).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    12.Howard-Varona, C. et al. Phage-specific metabolic reprogramming of virocells. ISME J. 14, 881–895 (2020).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    13.Nee, S. & Maynard Smith, J. The evolutionary biology of molecular parasites. Parasitology 100, S5–S18 (1990).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    14.Hambly, E. & Suttle, C. A. The viriosphere, diversity, and genetic exchange within phage communities. Curr. Opin. Microbiol. 8, 444–450 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    15.Sullivan, M. B. et al. Prevalence and evolution of core photosystem II genes in marine cyanobacterial viruses and their hosts. PLoS Biol. 4, e234 (2006).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    16.Holmes, E. C. What does virus evolution tell us about virus origins? J. Virol. 85, 5247–5251 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Wolf, Y. I. et al. Origins and evolution of the global RNA virome. mBio 9, e02329-18 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    18.Kuhn, J. H. et al. Classify viruses-the gain is worth the pain. Nature 566, 318–320 (2019).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    19.Record, N. R., Talmy, D. & Våge, S. Quantifying tradeoffs for marine viruses. Front. Mar. Sci. https://doi.org/10.3389/fmars.2016.00251 (2016). Investigates trade-offs in phenotypes of marine viruses that may influence virus population dynamics and biogeography.Article 

    Google Scholar 
    20.Domingo, E. et al. Basic concepts in RNA virus evolution. FASEB J. 10, 859–864 (1996).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    21.Solé, R. V., Ferrer, R., González-García, I., Quer, J. & Domingo, E. Red queen dynamics, competition and critical points in a model of RNA virus quasispecies. J. Theor. Biol. 198, 47–59 (1999).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    22.Stern, A. & Sorek, R. The phage-host arms race: shaping the evolution of microbes. Bioessays 33, 43–51 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    23.Daugherty, M. D. & Malik, H. S. Rules of engagement: molecular insights from host-virus arms races. Annu. Rev. Genet. 46, 677–700 (2012).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    24.Tegally, H. et al. Sixteen novel lineages of SARS-CoV-2 in South Africa. Nat. Med. 27, 440–446 (2021).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    25.Lederberg, J. in Emerging Viruses (ed. Morse, S. S.) 3–9 (Oxford University Press, 1993).26.Baltimore, D. Expression of animal virus genomes. Microbiol. Mol. Biol. Rev. 35, 235–241 (1971).CAS 

    Google Scholar 
    27.Coutinho, F. H., Edwards, R. A. & Rodríguez-Valera, F. Charting the diversity of uncultured viruses of archaea and bacteria. BMC Biol. 17, 109 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    28.King, A. M. Q., Adams, M. J., Carstens, E. B. & Lefkowitz, E. J. (eds) Virus Taxonomy. 163–173 (Elsevier, 2012).29.Forterre, P. The virocell concept and environmental microbiology. ISME J. 7, 233–236 (2013). Among the first reports articulating the viewpoint that infected cells undergoing active virus replication should be recognized as the ‘living form’ of a virus known as a virocell.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    30.Lowen, A. C. Constraints, drivers, and implications of influenza A virus reassortment. Annu. Rev. Virol. 4, 105–121 (2017).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    31.Mahner, M. & Kary, M. What exactly are genomes, genotypes and phenotypes? And what about phenomes? J. Theor. Biol. 186, 55–63 (1997).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    32.Edwards, K. F. & Steward, G. F. Host traits drive viral life histories across phytoplankton viruses. Am. Nat. 191, 566–581 (2018). Examines the inter-relationships between virus traits and their consequences for population dynamics and the evolution of burst size.PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    33.Flint, S. J., Racaniello, V. R., Rall, G. F., Skalka, A. M. & Enquist, L. W. Principles of Virology 4th Edn (Wiley, 2015).34.Ghabrial, S. A., Castón, J. R., Jiang, D., Nibert, M. L. & Suzuki, N. 50-plus years of fungal viruses. Virology 479–480, 356–368 (2015).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    35.Dunigan, D. D. et al. Chloroviruses lure hosts through long-distance chemical signaling. J. Virol. 93, e01688-18 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    36.Anantharaman, K. et al. Sulfur oxidation genes in diverse deep-sea viruses. Science 344, 757–760 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    37.Mann, N. H., Cook, A., Millard, A., Bailey, S. & Clokie, M. Bacterial photosynthesis genes in a virus. Nature 424, 741 (2003). Shows how the virus genome interacts with the host to facilitate virus reproduction.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    38.Mavrich, T. N. & Hatfull, G. F. Evolution of superinfection immunity in cluster A mycobacteriophages. mBio 10, e00971-19 (2019).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    39.Marine, R. L., Nasko, D. J., Wray, J., Polson, S. W. & Wommack, K. E. Novel chaperonins are prevalent in the virioplankton and demonstrate links to viral biology and ecology. ISME J. 11, 2479–2491 (2017).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    40.ICTV. Virus Taxonomy: The ICTV Report on Virus Classification and Taxon Nomenclature. https://talk.ictvonline.org/ictv-reports/ictv_9th_report/ (2019).41.Ojosnegros, S. et al. Viral genome segmentation can result from a trade-off between genetic content and particle stability. PLoS Genet 7, e1001344 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    42.Belshaw, R., Pybus, O. G. & Rambaut, A. The evolution of genome compression and genomic novelty in RNA viruses. Genome Res. 17, 1496–1504 (2007).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    43.Van Etten, J. L., Agarkova, I. V. & Dunigan, D. D. Chloroviruses. Viruses 12, 20 (2020).Article 
    CAS 

    Google Scholar 
    44.Iranzo, J. & Manrubia, S. C. Evolutionary dynamics of genome segmentation in multipartite viruses. Proc. Biol. Sci. 279, 3812–3819 (2012).PubMed 
    PubMed Central 

    Google Scholar 
    45.Kellogg, C. A. & Paul, J. H. Degree of ultraviolet radiation damage and repair capabilities are related to G+C content in marine vibriophages. Aquat. Microb. Ecol. 27, 13–20 (2002).Article 

    Google Scholar 
    46.Violle, C. et al. Let the concept of trait be functional! Oikos 116, 882–892 (2007).Article 

    Google Scholar 
    47.Edwards, K. F., Steward, G. F. & Schvarcz, C. R. Making sense of virus size and the tradeoffs shaping viral fitness. Ecol. Lett. 24, 363–373 (2021).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    48.Bonachela, J. A. & Levin, S. A. Evolutionary comparison between viral lysis rate and latent period. J. Theor. Biol. 345, 32–42 (2014).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    49.Yashchenko, V. V., Gavrilova, O. V., Rautian, M. S. & Jakobsen, K. S. Association of Paramecium bursaria Chlorella viruses with Paramecium bursaria cells: ultrastructural studies. Eur. J. Protistol. 48, 149–159 (2012).PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    50.DeLong, J. P., Al-Ameeli, Z., Duncan, G., Van Etten, J. L. & Dunigan, D. D. Predators catalyze an increase in chloroviruses by foraging on the symbiotic hosts of zoochlorellae. Proc. Natl Acad. Sci. USA 113, 13780–13784 (2016).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    51.Wang, I.-N. Lysis timing and bacteriophage fitness. Genetics 172, 17–26 (2006).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    52.Smith, C. & Fretwell, S. The optimal balance between size and number of offspring. Am. Nat. 108, 499–506 (1974).Article 

    Google Scholar 
    53.You, L., Suthers, P. F. & Yin, J. Effects of Escherichia coli physiology on growth of phage T7 In vivo and in silico. J. Bacteriol. 184, 1888–1894 (2002).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    54.Swan, B. K. et al. Prevalent genome streamlining and latitudinal divergence of planktonic bacteria in the surface ocean. Proc. Natl Acad. Sci. USA 110, 11463–11468 (2013).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    55.Hellweger, F. L. Carrying photosynthesis genes increases ecological fitness of cyanophage in silico. Environ. Microbiol. 11, 1386–1394 (2009).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    56.Schenk, H. & Sieber, M. Bacteriophage can promote the emergence of physiologically sub-optimal host phenotypes. bioRxiv https://doi.org/10.1101/621524 (2019).Article 

    Google Scholar 
    57.Howard-Varona, C. et al. Multiple mechanisms drive phage infection efficiency in nearly identical hosts. ISME J. 12, 1605–1618 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    58.Zimmerman, A. E. et al. Metabolic and biogeochemical consequences of viral infection in aquatic ecosystems. Nat. Rev. Microbiol. 18, 21–34 (2020).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    59.Grove, J. & Marsh, M. The cell biology of receptor-mediated virus entry. J. Cell Biol. 195, 1071–1082 (2011).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    60.McFadden, G., Mohamed, M. R., Rahman, M. M. & Bartee, E. Cytokine determinants of viral tropism. Nat. Rev. Immunol. 9, 645–655 (2009).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    61.Bernheim, A. & Sorek, R. The pan-immune system of bacteria: antiviral defence as a community resource. Nat. Rev. Microbiol. 18, 113–119 (2020).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    62.Nussenzweig, P. M. & Marraffini, L. A. Molecular mechanisms of CRISPR-Cas immunity in bacteria. Annu. Rev. Genet. 54, 93–120 (2020).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    63.Hampton, H. G., Watson, B. N. J. & Fineran, P. C. The arms race between bacteria and their phage foes. Nature 577, 327–336 (2020). An overview of the mechanisms and phenotypes related to phage infection and host defence mechanisms.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    64.Samson, J. E., Magadán, A. H., Sabri, M. & Moineau, S. Revenge of the phages: defeating bacterial defences. Nat. Rev. Microbiol. 11, 675–687 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    65.Flores, C. O., Meyer, J. R., Valverde, S., Farr, L. & Weitz, J. S. Statistical structure of host–phage interactions. Proc. Natl Acad. Sci. USA 108, E288–E297 (2011). Demonstrates the role of virus host range in generating community-wide patterns of host–phage interactions.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    66.Regoes, R. R. & Bonhoeffer, S. The HIV coreceptor switch: a population dynamical perspective. Trends Microbiol. 13, 269–277 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    67.Atkinson, D., Ciotti, B. J. & Montagnes, D. J. Protists decrease in size linearly with temperature: ca. 2.5% C-1. Proc. R. Soc. Lond. B 270, 2605–2611 (2003).Article 

    Google Scholar 
    68.Falkowski, P. G. in Primary Productivity in the Sea (ed. Falkowski, P. G.) 99–119 (Springer, 1980).69.Salsbery, M. E. & DeLong, J. P. The benefit of algae endosymbionts in Paramecium bursariais temperature dependent. Evol. Ecol. Res. 19, 669–678 (2018).
    Google Scholar 
    70.Kimmance, S. A., Atkinson, D. & Montagnes, D. J. S. Do temperature–food interactions matter? Responses of production and its components in the model heterotrophic flagellate Oxyrrhis marina. Aquat. Microb. Ecol. 42, 63–73 (2006).Article 

    Google Scholar 
    71.Maat, D. S., van Bleijswijk, J. D. L., Witte, H. J. & Brussaard, C. P. D. Virus production in phosphorus-limited Micromonas pusilla stimulated by a supply of naturally low concentrations of different phosphorus sources, far into the lytic cycle. FEMS Microbiol. Ecol. 92, fiw136 (2016).PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    72.Amla, D. V., Rowell, P. & Stewart, W. D. P. Metabolic changes associated with cyanophage N-1 infection of the cyanobacterium Nostoc muscorum. Arch. Microbiol. 148, 321–327 (1987).CAS 
    Article 

    Google Scholar 
    73.Hadas, H., Einav, M., Fishov, I. & Zaritsky, A. Bacteriophage T4 development depends on the physiology of its host Escherichia coli. Microbiology 143, 179–185 (1997).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    74.Demory, D. et al. Temperature is a key factor in Micromonas–virus interactions. ISME J. 11, 601–612 (2017). Shows the effect of temperature on the kinetics, phenotypes and life history strategies of prasinoviruses.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    75.Schachtele, C. F., Oman, R. W. & Anderson, D. L. Effect of elevated temperature on deoxyribonucleic acid synthesis in bacteriophage φ29-infected Bacillus amyloliquefaciens. J. Virol. 6, 430–437 (1970).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    76.Choua, M., Heath, M. R., Speirs, D. C. & Bonachela, J. A. The effect of viral plasticity on the persistence of host-virus systems. J. Theor. Biol. 498, 110263 (2020).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    77.Ni, T. & Zeng, Q. Diel infection of cyanobacteria by cyanophages. Front. Mar. Sci. https://doi.org/10.3389/fmars.2015.00123 (2016).Article 

    Google Scholar 
    78.Sakowski, E. G. et al. Ribonucleotide reductases reveal novel viral diversity and predict biological and ecological features of unknown marine viruses. Proc. Natl Acad. Sci. USA 111, 15786–15791 (2014). Demonstrates that genomic features in the viral replicon (that is, module of genes responsible for viral genome replication) may predict the biogeographical distribution of viruses.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    79.Reeson, A. F. et al. Effects of phenotypic plasticity on pathogen transmission in the field in a Lepidoptera-NPV system. Oecologia 124, 373–380 (2000).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    80.Stearns, S. C. The evolutionary significance of phenotypic plasticity. BioScience 39, 436–445 (1989).Article 

    Google Scholar 
    81.Leggett, H. C., Benmayor, R., Hodgson, D. J. & Buckling, A. Experimental evolution of adaptive phenotypic plasticity in a parasite. Curr. Biol. 23, 139–142 (2013).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    82.Oppenheim, A. B., Kobiler, O., Stavans, J., Court, D. L. & Adhya, S. Switches in bacteriophage lambda development. Annu. Rev. Genet. 39, 409–429 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    83.Erez, Z. et al. Communication between viruses guides lysis–lysogeny decisions. Nature 541, 488–493 (2017). Demonstrates the use of communication peptides that determine lysogeny in temperate phages.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    84.Weitz, J. S., Li, G., Gulbudak, H., Cortez, M. H. & Whitaker, R. J. Viral invasion fitness across a continuum from lysis to latency. Virus Evol. 5, vez006 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    85.Labonté, J. M. et al. Single cell genomics indicates horizontal gene transfer and viral infections in a deep subsurface Firmicutes population. Front. Microbiol. 6, 349 (2015).PubMed 
    PubMed Central 

    Google Scholar 
    86.Koskella, B. & Brockhurst, M. A. Bacteria–phage coevolution as a driver of ecological and evolutionary processes in microbial communities. FEMS Microbiol. Rev. 38, 916–931 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    87.Meyer, J. R. et al. Repeatability and contingency in the evolution of a key innovation in phage lambda. Science 335, 428–432 (2012).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    88.Marston, M. F. et al. Rapid diversification of coevolving marine Synechococcus and a virus. Proc. Natl Acad. Sci. USA 109, 4544–4549 (2012). Demonstrates the rapid co-evolution of virus and host but highlights the challenge of identifying the critical phenotypes mediating the interaction.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    89.Frickel, J., Feulner, P. G. D., Karakoc, E. & Becks, L. Population size changes and selection drive patterns of parallel evolution in a host–virus system. Nat. Commun. 9, 1706 (2018).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    90.Knowles, B. et al. Temperate infection in a virus–host system previously known for virulent dynamics. Nat. Commun. 11, 4626 (2020).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    91.Wang, I.-N., Dykhuizen, D. E. & Slobodkin, L. B. The evolution of phage lysis timing. Evol. Ecol. 10, 545–558 (1996).Article 

    Google Scholar 
    92.Abedon, S. T., Hyman, P. & Thomas, C. Experimental examination of bacteriophage latent-period evolution as a response to bacterial availability. Appl. Environ. Microbiol. 69, 7499–7506 (2003).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    93.Palkovacs, E. P. & Hendry, A. P. Eco-evolutionary dynamics: intertwining ecological and evolutionary processes in contemporary time. F1000 Biol. Rep. 2, 1 (2010).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    94.Brown, C. M., Lawrence, J. E. & Campbell, D. A. Are phytoplankton population density maxima predictable through analysis of host and viral genomic DNA content? J. Mar. Biol. Assoc. UK 86, 491–498 (2006).CAS 
    Article 

    Google Scholar 
    95.Wommack, K. E. & Colwell, R. R. Virioplankton: viruses in aquatic ecosystems. Microbiol. Mol. Biol. Rev. 64, 69–114 (2000).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    96.Weitz, J. S. et al. A multitrophic model to quantify the effects of marine viruses on microbial food webs and ecosystem processes. ISME J. 9, 1352–1364 (2015).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    97.Poorvin, L., Rinta-Kanto, J. M., Hutchins, D. A. & Wilhelm, S. W. Viral release of iron and its bioavailability to marine plankton. Limnol. Oceanogr. 49, 1734–1741 (2004).CAS 
    Article 

    Google Scholar 
    98.Shelford, E. J., Middelboe, M., Møller, E. F. & Suttle, C. A. Virus-driven nitrogen cycling enhances phytoplankton growth. Aquat. Microb. Ecol. 66, 41–46 (2012).Article 

    Google Scholar 
    99.Ankrah, N. Y. D. et al. Phage infection of an environmentally relevant marine bacterium alters host metabolism and lysate composition. ISME J. 8, 1089–1100 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    100.Jover, L. F., Effler, T. C., Buchan, A., Wilhelm, S. W. & Weitz, J. S. The elemental composition of virus particles: implications for marine biogeochemical cycles. Nat. Rev. Microbiol. 12, 519–528 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    101.Dawkins, R. The Extended Phenotype: The Long Reach of the Gene (Oxford University Press, 1999).102.Dawkins, R. Extended phenotype–but not too extended. A reply to Laland, Turner and Jablonka. Biol. Philosophy 19, 377–396 (2004).Article 

    Google Scholar 
    103.Ogata, H. Habitat alterations by viruses: strategies by Tupanviruses and others. Microbes Environ. 33, 117–119 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    104.Abrahão, J. et al. Tailed giant Tupanvirus possesses the most complete translational apparatus of the known virosphere. Nat. Commun. 9, 749 (2018).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    105.Clark, H. F. & Wiktor, T. J. Plasticity of phenotypic characters of rabies-related viroses: spontaneous variation in the plaque morphology, virulence, and temperature-sensitivity characters of serially propagated Lagos bat and Mokola viruses. J. Infect. Dis. 130, 608–618 (1974).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    106.Abedon, S. T. & Culler, R. R. Optimizing bacteriophage plaque fecundity. J. Theor. Biol. 249, 582–592 (2007).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    107.Luo, E., Eppley, J. M., Romano, A. E., Mende, D. R. & DeLong, E. F. Double-stranded DNA virioplankton dynamics and reproductive strategies in the oligotrophic open ocean water column. ISME J. 14, 1304–1315 (2020).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    108.Bidle, K. D. Elucidating marine virus ecology through a unified heartbeat. Proc. Natl Acad. Sci. USA 111, 15606–15607 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    109.Schmidt, H. F., Sakowski, E. G., Williamson, S. J., Polson, S. W. & Wommack, K. E. Shotgun metagenomics indicates novel family A DNA polymerases predominate within marine virioplankton. ISME J. 8, 103–114 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    110.Nasko, D. J. et al. Family A DNA polymerase phylogeny uncovers diversity and replication gene organization in the virioplankton. Front. Microbiol. 9, 3053 (2018).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    111.Harrison, A. O., Moore, R. M., Polson, S. W. & Wommack, K. E. Reannotation of the ribonucleotide reductase in a cyanophage reveals life history strategies within the virioplankton. Front. Microbiol. 10, 134 (2019).PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    112.Breitbart, M. Marine viruses: truth or dare. Annu. Rev. Mar. Sci. 4, 425–448 (2012).Article 

    Google Scholar 
    113.Hurwitz, B. L. & U’Ren, J. M. Viral metabolic reprogramming in marine ecosystems. Curr. Opin. Microbiol. 31, 161–168 (2016).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    114.Lindell, D., Jaffe, J. D., Johnson, Z. I., Church, G. M. & Chisholm, S. W. Photosynthesis genes in marine viruses yield proteins during host infection. Nature 438, 86–89 (2005).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    115.Rusconi, R., Garren, M. & Stocker, R. Microfluidics expanding the frontiers of microbial ecology. Annu. Rev. Biophys. 43, 65–91 (2014).CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    116.Walker, G. M., Ozers, M. S. & Beebe, D. J. Cell infection within a microfluidic device using virus gradients. Sens. Actuators B Chem. 98, 347–355 (2004).CAS 
    Article 

    Google Scholar 
    117.Cimetta, E. et al. Microfluidic-driven viral infection on cell cultures: theoretical and experimental study. Biomicrofluidics 6, 024127 (2012).PubMed Central 
    Article 
    CAS 

    Google Scholar 
    118.Xu, N. et al. A microfluidic platform for real-time and in situ monitoring of virus infection process. Biomicrofluidics 6, 034122 (2012).PubMed Central 
    Article 
    CAS 

    Google Scholar 
    119.Akin, D., Li, H. & Bashir, R. Real-time virus trapping and fluorescent imaging in microfluidic devices. Nano Lett. 4, 257–259 (2004).CAS 
    Article 

    Google Scholar 
    120.Yu, J. Q. et al. Droplet optofluidic imaging for λ-bacteriophage detection via co-culture with host cell Escherichia coli. Lab. Chip 14, 3519–3524 (2014).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    121.Mashaghi, S. & van Oijen, A. M. Droplet microfluidics for kinetic studies of viral fusion. Biomicrofluidics 10, 024102 (2016).PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    122.Fischer, A. E. et al. A high-throughput drop microfluidic system for virus culture and analysis. J. Virol. Methods 213, 111–117 (2015).CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar  More

  • in

    Characteristics of pulmonary microvascular structure in postnatal yaks

    AnimalsThe experimental yaks were divided into four groups: 1-day old, 30-days-old, 180-days-old and adult. Three yaks were selected for each group, regardless of sex, and purchased from a local herdsmen in Haiyan County of Qinghai Province. All of the yaks showed a good nutritional status, and appeared healthy with no apparent diseases or conditions. The yaks were sacrificed by exsanguination in a slaughterhouse. The lungs were obtained immediately after the yak had died, and tissue samples were immediately collected from the diaphragmatic lobe of right lungs (to ensure that obvious blood vessels and the trachea were not gathered). The tissue samples were divided into three parts. One part was cut into 1 cm3 sections and fixed with 4% paraformaldehyde (PFA). The other two parts were cut into 1 mm3 pieces; one part was fixed with 2.5% glutaraldehyde, and the other was put into a freezing tube and placed into liquid nitrogen.Ethics statementThis study was approved by the Institutional Animal Care and Use Committee of Qinghai University (Xining, China). All methods were carried out in accordance with the ARRIVE guidelines and the Animal Ethics Procedures and Guidelines of the People’s Republic of China. No local regulations or laws were overlooked. All yaks used in this study were purchased from local farmers.Haematoxylin and eosin stainingLung tissue samples (1 cm3) were fixed in 4% PFA, dehydrated in 30%, 50%, 75%, 95% and 100% ethanol and then treated with xylene before embedding in paraffin. Paraffin-embedded lung tissues were cut into 4 µm sections. The sections were deparaffinized in xylene, and sections were stained either with haematoxylin and eosin (HE) (Y&K Bio, Xi’an, China) or Masson’s trichrome stain, to examine general morphology.ImmunohistochemistryThe unstained, deparaffinized sections were rinsed with Phosphate Buffered Saline with Twen-20 (PBST) 3 times for 5 min each time. Then, endogenous peroxidase was quenched using 3% peroxide-methanol at room temperature in the dark for 25 min, and then the samples were placed on a decolorizing shaking table 3 times, for 5 min each. The slides were then incubated with 3% foetal bovine serum (Sangon Biotech, Shanghai, China) at room temperature for 25 min. The serum was discarded, and rabbit anti-cattle CD34 and rabbit anti-CD34 polyclonal antibodies (Proteintech group, Wuhan, China) diluted in phosphate buffer saline (PBS) were added. CD34 is a transmembrane glycoprotein known as an angiogenesis marker. The sections were incubated in the primary antibodies overnight at 4 °C. Then, the sections were rinsed in Phosphate Buffered Saline with Twen-20 (PBST) (3 × 5 min), goat anti-rabbit IgG was added, and the sections were incubated for 30 min at 37 °C. 3,3-Diaminobenzidin (DAB) was added to the sections to visualise antibody binding, and the sections were washed 3 times in PBST. Haematoxylin was used to counterstain the nucleus prior to the samples being dehydrated and mounted.An Olympus BX51 microscope was used to take photomicrographs of the microstructures, images depict 1000× magnification. Transmission electron microscopy.The TEM lung tissue samples were processed using previously published methods16. Fresh lung samples (1 mm3) were fixed with glutaraldehyde (2.5%, 24 h) and postfixed with osmium tetroxide (1%, 2 h). The samples were dehydrated in a series of increasing concentrations of ethanol and embedded in Epon812. After preparing semithin sections, ultrathin sections were double stained with uranyl acetate and lead citrate. A 10,000× magnification was used to observe and photograph the sections with a JEM 1230 electron microscope (JEOL, Tokyo, Japan) set at 120 kV.Quantitative real-time PCR (qPCR)The gene expression levels in lung tissues from the yaks in the four age groups were analysed using qPCR. Total RNA was isolated with TRIzol® reagent (Invitrogen, CA, USA). cDNA was obtained by reverse transcription of total RNA using the SYBR PrimeScript RT reagent Kit with gDNA Eraser (Perfect Real Time; Takara, Dalian, China). The forward and reverse primers sequences for the qPCR are shown in Table 1. The genes expression levels were detected using TB Green™ Premix Ex Taq™ II (TIi RNaseH Plus; Takara, Dalian, China) according to the manufacturer’s instructions. The 2−ΔΔCT method was used to analyse the relative expression of target genes, and the housekeeping gene β-actin was used for normalization.Table 1 Primer sequences.Full size tableWestern blot analysisEqual amounts of proteins of yak lung tissue in different development stages were harvested. These proteins were separated on 10% polyacrylamide gels and transferred onto polyvinylidene difluoride (PVDF) membranes (Sangon Biotech, Shanghai, China). PVDF membranes were blocked in 10% non-fat (skimmed) milk for 3 h and then incubated in rabbit anti-VEGFA polyclonal antibody (OriGene, Maryland, USA) at 4 °C overnight. The membranes were then incubated with a goat anti-rabbit IgG antibody (Abcam, Cambridge, UK) for 2 h being washed 3 times (10 min / time) with Tris-buffered saline with Twen-20 (TBST; containing 0.1% Twen-20). All antibodies were diluted according to the manufacturer’s instructions. Immunoblots were analysed by autograph using a Gel Doc™ XR + Gel documentation system (BIO-RAD, California, USA).Statistical analysisThe experimental data are showed as the mean ± standard deviation (SD). The differences between the four groups were compared using one-way ANOVA. P values at less than 0.05 were considered significantly different. More

  • in

    Plant-microbe interactions in the phyllosphere: facing challenges of the anthropocene

    1.Kalnay E, Cai M. Impact of urbanization and land-use change on climate. Nature. 2003;423:528–31.CAS 
    PubMed 
    Article 

    Google Scholar 
    2.Archer SDJ, Pointing SB. Anthropogenic impact on the atmospheric microbiome. Nat Microbiol. 2020;5:229–31.CAS 
    PubMed 
    Article 

    Google Scholar 
    3.Powers RP, Jetz W. Global habitat loss and extinction risk of terrestrial vertebrates under future land-use-change scenarios. Nat Clim Change. 2019;9:323–9.Article 

    Google Scholar 
    4.Sandifer PA, Sutton-Grier AE, Ward BP. Exploring connections among nature, biodiversity, ecosystem services, and human health and well-being: Opportunities to enhance health and biodiversity conservation. Ecosyst Serv. 2015;12:1–15.Article 

    Google Scholar 
    5.Jansson JK, Hofmockel KS. Soil microbiomes and climate change. Nat Rev Microbiol. 2020;18:35–46.CAS 
    PubMed 
    Article 

    Google Scholar 
    6.Huttenhower C, Gevers D, Knight R, Abubucker S, Badger JH, Chinwalla AT, et al. Structure, function and diversity of the healthy human microbiome. Nature. 2012;486:207–14.CAS 
    Article 

    Google Scholar 
    7.Banerjee S, Schlaeppi K, van der Heijden MGA. Keystone taxa as drivers of microbiome structure and functioning. Nat Rev Microbiol. 2018;16:567–76.CAS 
    PubMed 
    Article 

    Google Scholar 
    8.Sapp M, Ploch S, Fiore-Donno AM, Bonkowski M, Rose LE. Protists are an integral part of the Arabidopsis thaliana microbiome. Environ Microbiol. 2018;20:30–43.CAS 
    PubMed 
    Article 

    Google Scholar 
    9.Vorholt JA. Microbial life in the phyllosphere. Nat Rev Microbiol. 2012;10:828–40.CAS 
    PubMed 
    Article 

    Google Scholar 
    10.Laforest-Lapointe I, Messier C, Kembel SW. Host species identity, site and time drive temperate tree phyllosphere bacterial community structure. Microbiome. 2016;4:27.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    11.Andrews JH, Harris RF. The ecology and biogeography of microorganisms on plant surfaces. Annu Rev Phytopathol. 2000;38:145–80.Article 

    Google Scholar 
    12.Lugtenberg B, Kamilova F. Plant-growth-promoting Rhizobacteria. Annu Rev Microbiol. 2009;63:541–56.CAS 
    PubMed 
    Article 

    Google Scholar 
    13.Philippot L, Raaijmakers JM, Lemanceau P, van der Putten WH. Going back to the roots: the microbial ecology of the rhizosphere. Nat Rev Microbiol. 2013;11:789–99.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    14.Davison J. Plant beneficial bacteria. Bio/Technol. 1988;6:282–6.CAS 

    Google Scholar 
    15.Schauer S, Kutschera U. A novel growth-promoting microbe, Methylobacterium funariae sp. nov., isolated from the leaf surface of a common moss. Plant Signal Behav. 2011;6:510–5.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    16.Innerebner G, Knief C, Vorholt JA. Protection of arabidopsis thaliana against leaf-pathogenic pseudomonas syringae by sphingomonas strains in a controlled model system. Appl Environ Microbiol. 2011;77:3202–10.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    17.Laforest-Lapointe I, Paquette A, Messier C, Kembel SW. Leaf bacterial diversity mediates plant diversity and ecosystem function relationships. Nature. 2017;546:145–7.CAS 
    PubMed 
    Article 

    Google Scholar 
    18.Koskella B, Meaden S, Crowther WJ, Leimu R, Metcalf CJE. A signature of tree health? Shifts in the microbiome and the ecological drivers of horse chestnut bleeding canker disease. N Phytol. 2017;215:737–46.CAS 
    Article 

    Google Scholar 
    19.Isbell F, Tilman D, Polasky S, Loreau M. The biodiversity-dependent ecosystem service debt. Ecol Lett. 2015;18:119–34.PubMed 
    Article 

    Google Scholar 
    20.Barnosky A, Matzke N, Tomiya S, Wogan G, Swartz B, Quental T, et al. Has the earth’s sixth mass extinction already arrived? Nat Nat. 2011;471:51–7.CAS 
    Article 

    Google Scholar 
    21.Pascual U, Balvanera P, Díaz S, Pataki G, Roth E, Stenseke M, et al. Valuing nature’s contributions to people: the IPBES approach. Curr Opin Environ Sustain. 2017;26–27:7–16.Article 

    Google Scholar 
    22.Cavicchioli R, Ripple WJ, Timmis KN, Azam F, Bakken LR, Baylis M, et al. Scientists’ warning to humanity: microorganisms and climate change. Nat Rev Microbiol. 2019;17:569–86.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    23.Annamalai J, Namasivayam V. Endocrine disrupting chemicals in the atmosphere: Their effects on humans and wildlife. Environ Int. 2015;76:78–97.CAS 
    PubMed 
    Article 

    Google Scholar 
    24.Jumpponen A, Jones KL. Seasonally dynamic fungal communities in the Quercus macrocarpa phyllosphere differ between urban and nonurban environments. N Phytol. 2010;186:496–513.CAS 
    Article 

    Google Scholar 
    25.Imperato V, Kowalkowski L, Portillo-Estrada M, Gawronski SW, Vangronsveld J, Thijs S. Characterisation of the Carpinus betulus L. Phyllomicrobiome in urban and forest areas. Front Microbiol. 2019;10:1110.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    26.Bowers RM, McLetchie S, Knight R, Fierer N. Spatial variability in airborne bacterial communities across land-use types and their relationship to the bacterial communities of potential source environments. ISME J. 2011;5:601–12.CAS 
    PubMed 
    Article 

    Google Scholar 
    27.Lymperopoulou DS, Adams RI, Lindow SE. Contribution of vegetation to the microbial composition of nearby outdoor air. Appl Environ Microbiol. 2016;82:3822–33.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    28.De Kempeneer L, Sercu B, Vanbrabant W, Van Langenhove H, Verstraete W. Bioaugmentation of the phyllosphere for the removal of toluene from indoor air. Appl Microbiol Biotechnol. 2004;64:284–8.PubMed 
    Article 
    CAS 

    Google Scholar 
    29.Hanski I, Hertzen Lvon, Fyhrquist N, Koskinen K, Torppa K, Laatikainen T, et al. Environmental biodiversity, human microbiota, and allergy are interrelated. Proc Natl Acad Sci. 2012;109:8334–9.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    30.Smets W, Wuyts K, Oerlemans E, Wuyts S, Denys S, Samson R, et al. Impact of urban land use on the bacterial phyllosphere of ivy (Hedera sp.). Atmos Environ. 2016;147:376–83.CAS 
    Article 

    Google Scholar 
    31.Laforest-Lapointe I, Messier C, Kembel SW. Tree Leaf Bacterial Community Structure and Diversity Differ along a Gradient of Urban Intensity. mSystems. 2017;2:e00087–17.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    32.Espenshade J, Thijs S, Gawronski S, Bové H, Weyens N, Vangronsveld J. Influence of urbanization on epiphytic bacterial communities of the platanus × hispanica tree leaves in a Biennial Study. Front Microbiol. 2019;10:675.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    33.Wuyts K, Smets W, Lebeer S, Samson R. Green infrastructure and atmospheric pollution shape diversity and composition of phyllosphere bacterial communities in an urban landscape. FEMS Microbiol Ecol 2020;96:fiz173.CAS 
    PubMed 
    Article 

    Google Scholar 
    34.Zhao D, Liu G, Wang X, Daraz U, Sun Q. Abundance of human pathogen genes in the phyllosphere of four landscape plants. J Environ Manag. 2020;255:109933.CAS 
    Article 

    Google Scholar 
    35.Gandolfi I, Canedoli C, Imperato V, Tagliaferri I, Gkorezis P, Vangronsveld J, et al. Diversity and hydrocarbon-degrading potential of epiphytic microbial communities on Platanus x acerifolia leaves in an urban area. Environ Pollut. 2017;220:650–8.CAS 
    PubMed 
    Article 

    Google Scholar 
    36.Weyens N, van der Lelie D, Taghavi S, Vangronsveld J. Phytoremediation: plant–endophyte partnerships take the challenge. Curr Opin Biotechnol. 2009;20:248–54.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    37.Afzal M, Khan QM, Sessitsch A. Endophytic bacteria: prospects and applications for the phytoremediation of organic pollutants. Chemosphere. 2014;117:232–42.CAS 
    PubMed 
    Article 

    Google Scholar 
    38.Siciliano SD, Fortin N, Mihoc A, Wisse G, Labelle S, Beaumier D, et al. Selection of specific endophytic bacterial genotypes by plants in response to soil contamination. Appl Environ Microbiol. 2001;67:2469–75.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    39.Barac T, Taghavi S, Borremans B, Provoost A, Oeyen L, Colpaert JV, et al. Engineered endophytic bacteria improve phytoremediation of water-soluble, volatile, organic pollutants. Nat Biotechnol. 2004;22:583–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    40.Sandhu A, Halverson LJ, Beattie GA. Bacterial degradation of airborne phenol in the phyllosphere. Environ Microbiol. 2007;9:383–92.CAS 
    PubMed 
    Article 

    Google Scholar 
    41.Weyens N, Thijs S, Popek R, Witters N, Przybysz A, Espenshade J, et al. The role of plant–microbe interactions and their exploitation for phytoremediation of air pollutants. Int J Mol Sci. 2015;16:25576–604.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    42.Essl F, Dullinger S, Rabitsch W, Hulme PE, Hülber K, Jarošík V, et al. Socioeconomic legacy yields an invasion debt. Proc Natl Acad Sci. 2011;108:203–7.CAS 
    PubMed 
    Article 

    Google Scholar 
    43.Walther G-R, Roques A, Hulme PE, Sykes MT, Pyšek P, Kühn I, et al. Alien species in a warmer world: risks and opportunities. Trends Ecol Evol. 2009;24:686–93.PubMed 
    Article 

    Google Scholar 
    44.Blüthgen N, Menzel F, Blüthgen N. Measuring specialization in species interaction networks. BMC Ecol. 2006;6:9.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    45.Cobian GM, Egan CP, Amend AS. Plant–microbe specificity varies as a function of elevation. ISME J. 2019;13:2778–88.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    46.Bálint M, Bartha L, O’Hara RB, Olson MS, Otte J, Pfenninger M, et al. Relocation, high-latitude warming and host genetic identity shape the foliar fungal microbiome of poplars. Mol Ecol. 2015;24:235–48.PubMed 
    Article 
    CAS 
    PubMed Central 

    Google Scholar 
    47.Vacher C, Cordier T, Vallance J. Phyllosphere fungal communities differentiate more thoroughly than bacterial communities along an elevation gradient. Micro Ecol. 2016;72:1–3.Article 

    Google Scholar 
    48.Callaway RM, Brooker RW, Choler P, Kikvidze Z, Lortie CJ, Michalet R, et al. Positive interactions among alpine plants increase with stress. Nature. 2002;417:844–8.CAS 
    PubMed 
    Article 
    PubMed Central 

    Google Scholar 
    49.Bever JD. Feeback between plants and their soil communities in an old field. Community Ecol. 1994;75:1965–77.Article 

    Google Scholar 
    50.Bever JD. Soil community feedback and the coexistence of competitors: conceptual frameworks and empirical tests. N Phytol. 2003;157:465–73.Article 

    Google Scholar 
    51.Klironomos JN. Feedback with soil biota contributes to plant rarity and invasiveness in communities. Nature. 2002;417:67–70.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    52.Reinhart KO, Callaway RM. Soil biota and invasive plants. N Phytol. 2006;170:445–57.Article 

    Google Scholar 
    53.Callaway RM, Thelen GC, Rodriguez A, Holben WE. Soil biota and exotic plant invasion. Nature. 2004;427:731–3.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    54.Brown CD, Vellend M. Non-climatic constraints on upper elevational plant range expansion under climate change. Proc R Soc B Biol Sci. 2014;281:20141779.Article 

    Google Scholar 
    55.Carteron A, Parasquive V, Blanchard F, Guilbeault‐Mayers X, Turner BL, Vellend M, et al. Soil abiotic and biotic properties constrain the establishment of a dominant temperate tree into boreal forests. J Ecol. 2020;108:931–44.Article 

    Google Scholar 
    56.Williamson M. Biological invasions. 1996. Springer Netherlands.57.Mitchell CE, Power AG. Release of invasive plants from fungal and viral pathogens. Nature. 2003;421:625–7.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    58.Ramirez KS, Snoek LB, Koorem K, Geisen S, Bloem LJ, ten Hooven F, et al. Range-expansion effects on the belowground plant microbiome. Nat Ecol Evol. 2019;3:604–11.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    59.Diez JM, Dickie I, Edwards G, Hulme PE, Sullivan JJ, Duncan RP. Negative soil feedbacks accumulate over time for non-native plant species. Ecol Lett. 2010;13:803–9.PubMed 
    Article 

    Google Scholar 
    60.Lenssen NJL, Schmidt GA, Hansen JE, Menne MJ, Persin A, Ruedy R, et al. Improvements in the GISTEMP uncertainty model. J Geophys Res Atmos. 2019;124:6307–26.Article 

    Google Scholar 
    61.O’brien RD, Lindow SE. Effect of plant species and environmental conditions on ice nucleation activity of pseudomonas syringae on leaves. Appl Environ Microbiol. 1988;54:2281–6.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    62.Klinkert B, Narberhaus F. Microbial thermosensors. Cell Mol Life Sci. 2009;66:2661–76.CAS 
    PubMed 
    Article 

    Google Scholar 
    63.Velásquez AC, Castroverde CDM, He SY. Plant-pathogen warfare under changing climate conditions. Curr Biol CB. 2018;28:R619–R634.PubMed 
    Article 
    CAS 

    Google Scholar 
    64.Compant S, van der Heijden MGA, Sessitsch A. Climate change effects on beneficial plant-microorganism interactions. FEMS Microbiol Ecol. 2010;73:197–214.CAS 
    PubMed 

    Google Scholar 
    65.Cheng YT, Zhang L, He SY. Plant-microbe interactions facing environmental challenge. Cell Host Microbe. 2019;26:183–92.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    66.Guerra CA, Delgado‐Baquerizo M, Duarte E, Marigliano O, Görgen C, Maestre FT, et al. Global projections of the soil microbiome in the Anthropocene. Glob Ecol Biogeogr. 2021;30:987–99.PubMed 
    Article 

    Google Scholar 
    67.Frindte K, Pape R, Werner K, Löffler J, Knief C. Temperature and soil moisture control microbial community composition in an arctic–alpine ecosystem along elevational and micro-topographic gradients. ISME J. 2019;13:2031–43.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    68.Cordier T, Robin C, Capdevielle X, Fabreguettes O, Desprez-Loustau M-L, Vacher C. The composition of phyllosphere fungal assemblages of European beech (Fagus sylvatica) varies significantly along an elevation gradient. N Phytol. 2012;196:510–9.Article 

    Google Scholar 
    69.Tedersoo L, Bahram M, Toots M, Diédhiou AG, Henkel TW, Kjøller R, et al. Towards global patterns in the diversity and community structure of ectomycorrhizal fungi. Mol Ecol. 2012;21:4160–70.PubMed 
    Article 

    Google Scholar 
    70.Gomes T, Pereira JA, Benhadi J, Lino-Neto T, Baptista P. Endophytic and epiphytic phyllosphere fungal communities are shaped by different environmental factors in a Mediterranean ecosystem. Micro Ecol. 2018;76:668–79.Article 

    Google Scholar 
    71.Peñuelas J, Rico L, Ogaya R, Jump AS, Terradas J. Summer season and long-term drought increase the richness of bacteria and fungi in the foliar phyllosphere of Quercus ilex in a mixed Mediterranean forest. Plant Biol Stuttg Ger. 2012;14:565–75.Article 

    Google Scholar 
    72.Rico L, Ogaya R, Terradas J, Peñuelas J. Community structures of N2 -fixing bacteria associated with the phyllosphere of a Holm oak forest and their response to drought. Plant Biol Stuttg Ger. 2014;16:586–93.CAS 
    Article 

    Google Scholar 
    73.Grady KL, Sorensen JW, Stopnisek N, Guittar J, Shade A. Assembly and seasonality of core phyllosphere microbiota on perennial biofuel crops. Nat Commun. 2019;10:1–10.Article 
    CAS 

    Google Scholar 
    74.Redford AJ, Fierer N. Bacterial Succession on the Leaf Surface: A Novel System for Studying Successional Dynamics. Micro Ecol. 2009;58:189–98.Article 

    Google Scholar 
    75.Edwards JA, Santos-Medellín CM, Liechty ZS, Nguyen B, Lurie E, Eason S, et al. Compositional shifts in root-associated bacterial and archaeal microbiota track the plant life cycle in field-grown rice. PLOS Biol. 2018;16:e2003862.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    76.Parmesan C, Yohe G. A globally coherent fingerprint of climate change impacts across natural systems. Nature. 2003;421:37–42.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    77.Zhao C, Liu B, Piao S, Wang X, Lobell DB, Huang Y, et al. Temperature increase reduces global yields of major crops in four independent estimates. Proc Natl Acad Sci. 2017;114:9326–31.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    78.Ray DK, Mueller ND, West PC, Foley JA. Yield trends are insufficient to double global crop production by 2050. PLOS ONE. 2013;8:e66428.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    79.Angel R, Soares MIM, Ungar ED, Gillor O. Biogeography of soil archaea and bacteria along a steep precipitation gradient. ISME J. 2010;4:553–63.PubMed 
    Article 

    Google Scholar 
    80.Kaisermann A, Vries FTde, Griffiths RI, Bardgett RD. Legacy effects of drought on plant–soil feedbacks and plant–plant interactions. N Phytol. 2017;215:1413–24.CAS 
    Article 

    Google Scholar 
    81.Hawkes CV, Kivlin SN, Rocca JD, Huguet V, Thomsen MA, Suttle KB. Fungal community responses to precipitation. Glob Change Biol. 2011;17:1637–45.Article 

    Google Scholar 
    82.Lau JA, Lennon JT. Rapid responses of soil microorganisms improve plant fitness in novel environments. Proc Natl Acad Sci. 2012;109:14058–62.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    83.Sheik CS, Beasley WH, Elshahed MS, Zhou X, Luo Y, Krumholz LR. Effect of warming and drought on grassland microbial communities. ISME J. 2011;5:1692–700.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    84.Bradford MA. Thermal adaptation of decomposer communities in warming soils. Front Microbiol. 2013;4:333.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    85.Li F, Deng J, Nzabanita C, Li Y, Duan T. Growth and physiological responses of perennial ryegrass to an AMF and an Epichloë endophyte under different soil water contents. Symbiosis. 2019;79:151–61.CAS 
    Article 

    Google Scholar 
    86.Ibekwe AM, Ors S, Ferreira JFS, Liu X, Suarez DL, Ma J, et al. Functional relationships between aboveground and belowground spinach (Spinacia oleracea L., cv. Racoon) microbiomes impacted by salinity and drought. Sci Total Environ. 2020;717:137207.CAS 
    PubMed 
    Article 

    Google Scholar 
    87.Prosser JI, Bohannan BJM, Curtis TP, Ellis RJ, Firestone MK, Freckleton RP, et al. The role of ecological theory in microbial ecology. Nat Rev Microbiol. 2007;5:384–92.CAS 
    PubMed 
    Article 

    Google Scholar 
    88.Shoemaker WR, Locey KJ, Lennon JT. A macroecological theory of microbial biodiversity. Nat Ecol Evol. 2017;1:0107.Article 

    Google Scholar 
    89.Ratzke C, Denk J, Gore J. Ecological suicide in microbes. Nat Ecol Evol. 2018;2:867–72.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    90.Shade A, Dunn RR, Blowes SA, Keil P, Bohannan BJM, Herrmann M, et al. Macroecology to unite all life, large and small. Trends Ecol Evol. 2018;33:731–44.PubMed 
    Article 

    Google Scholar 
    91.Grilli J. Macroecological laws describe variation and diversity in microbial communities. Nat Commun. 2020;11:4743.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    92.Knief C, Ramette A, Frances L, Alonso-Blanco C, Vorholt JA. Site and plant species are important determinants of the Methylobacterium community composition in the plant phyllosphere. ISME J. 2010;4:719–28.CAS 
    PubMed 
    Article 

    Google Scholar 
    93.Redford AJ, Bowers RM, Knight R, Linhart Y, Fierer N. The ecology of the phyllosphere: geographic and phylogenetic variability in the distribution of bacteria on tree leaves: Biogeography of phyllosphere bacterial communities. Environ Microbiol. 2010;12:2885–93.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    94.Remus-Emsermann MNP, Tecon R, Kowalchuk GA, Leveau JHJ. Variation in local carrying capacity and the individual fate of bacterial colonizers in the phyllosphere. ISME J. 2012;6:756–65.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    95.Kembel SW, O’Connor TK, Arnold HK, Hubbell SP, Wright SJ, Green JL. Relationships between phyllosphere bacterial communities and plant functional traits in a neotropical forest. Proc Natl Acad Sci. 2014;111:13715–20.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    96.Maignien L, DeForce EA, Chafee ME, Eren AM, Simmons SL. Ecological succession and stochastic variation in the assembly of Arabidopsis thaliana phyllosphere communities. mBio. 2014;5:e00682–13.PubMed 
    PubMed Central 
    Article 
    CAS 

    Google Scholar 
    97.Wagner MR, Lundberg DS, del Rio TG, Tringe SG, Dangl JL, Mitchell-Olds T. Host genotype and age shape the leaf and root microbiomes of a wild perennial plant. Nat Commun. 2016;7:12151.CAS 
    PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    98.Carlström CI, Field CM, Bortfeld-Miller M, Müller B, Sunagawa S, Vorholt JA. Synthetic microbiota reveal priority effects and keystone strains in the Arabidopsis phyllosphere. Nat. Ecol Evol. 2019;3:1445–54.
    Google Scholar 
    99.Lajoie G, Maglione R, Kembel SW. Adaptive matching between phyllosphere bacteria and their tree hosts in a neotropical forest. Microbiome. 2020;8:70.PubMed 
    PubMed Central 
    Article 

    Google Scholar 
    100.Massoni J, Bortfeld-Miller M, Jardillier L, Salazar G, Sunagawa S, Vorholt JA. Consistent host and organ occupancy of phyllosphere bacteria in a community of wild herbaceous plant species. ISME J. 2020;14:245–58.CAS 
    PubMed 
    Article 

    Google Scholar 
    101.Lajoie G, Kembel SW. Host neighborhood shapes bacterial community assembly and specialization on tree species across a latitudinal gradient. Ecol Monogr. 2021;91:e01443.Article 

    Google Scholar 
    102.Vellend M. Conceptual synthesis in community ecology. Q Rev Biol. 2010;85:183–206.Article 
    PubMed 

    Google Scholar 
    103.Bernhardt ES, Rosi EJ, Gessner MO. Synthetic chemicals as agents of global change. Front Ecol Environ. 2017;15:84–90.Article 

    Google Scholar  More

  • in

    Preventing spillover as a key strategy against pandemics

    CORRESPONDENCE
    14 September 2021

    Preventing spillover as a key strategy against pandemics

    Neil M. Vora

     ORCID: http://orcid.org/0000-0002-4989-3108

    0
    ,

    Nigel Sizer

    1
    &

    Aaron Bernstein

    2

    Neil M. Vora

    Conservation International, Arlington, Virginia, USA.

    View author publications

    You can also search for this author in PubMed
     Google Scholar

    Nigel Sizer

    Preventing Pandemics at the Source Coalition, Mount Kisco, New York, USA.

    View author publications

    You can also search for this author in PubMed
     Google Scholar

    Aaron Bernstein

    Boston Children’s Hospital, Boston, Massachusetts, USA.

    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

    Download PDF

    Most new infectious diseases result from the spillover of pathogens from animals, particularly wildlife, to people. Spillover prevention should not be dismissed in discussions on how best to address pandemics (see Nature 596, 332–335; 2021).The belief that we are powerless to prevent spillover is, unfortunately, endorsed by many in public health and government. Improved management of farmed animals, regulations on wildlife trade and conservation of tropical forests have all helped to prevent spillover and subsequent outbreaks, as well as boosting greenhouse-gas mitigation and wildlife conservation (see go.nature.com/2uqwx1u). Moreover, preventing spillover is cheap compared with the costs of a single pandemic (A. P. Dobson et al. Science 369, 379–381; 2020).Outbreak containment measures will always be necessary, especially for the most vulnerable people in resource-limited settings, because spillover can never be completely eliminated. But if prioritized alongside post-spillover initiatives, outcomes will be more cost-effective, scientifically informed and equitable.

    Nature 597, 332 (2021)
    doi: https://doi.org/10.1038/d41586-021-02427-4

    Competing Interests
    The authors declare no competing interests.

    Related Articles

    See more letters to the editor

    Subjects

    Diseases

    Policy

    Conservation biology

    Latest on:

    Diseases

    How COVID is derailing the fight against HIV, TB and malaria
    News 10 SEP 21

    Cells of the human intestinal tract mapped across space and time
    Article 08 SEP 21

    Burden and characteristics of COVID-19 in the United States during 2020
    Article 26 AUG 21

    Policy

    Afghanistan: conflict risks local and global health
    Correspondence 14 SEP 21

    Climate science is supporting lawsuits that could help save the world
    News Feature 08 SEP 21

    Indonesia’s science super-agency must earn researchers’ trust
    Editorial 08 SEP 21

    Jobs

    Post-Doctoral Position

    McGuire Research Institute, Inc.
    Richmond, VA, United States

    Postdoctoral Research Associate

    Washington University in St. Louis (WUSTL)
    Saint Louis, MO, United States

    Division Head, General Internal Medicine and Geriatrics, University Health Network and Sinai Health Systems

    University Health Network (UHN), U of T
    Toronto, Ontario, Canada

    Director Protein Science

    Bright Peak Therapeutics
    Basel, Switzerland More