Ecology

Insect preparationWe collected 200–300 larvae and pupae of the diamondback moth from cabbage and cauliflower fields in Wuhan, Beijing, and Shenyang in late September from 2008 to 2012. We mixed all collections into one stock colony because of no geographic differentiation in this species from these sites59. We reared all individuals on cabbage leaves spread evenly across five screen cages (35 × 35 × 15 cm) in growth chambers at constant temperature (25 ± 1 °C) with 15-h light:9-h dark photoperiod, and relative humidity set at 60 ± 10%. We moved any new pupae to new screen cages for adult emergence. Emerged adults were fed with 10% honey solution in cotton balls. To collect eggs, we dipped four small pieces of laboratory film (7 × 5 cm) in fresh cabbage juice for 3–5 s and hung the treated film pieces on the top of each screen cage. To further enlarge the population for our experiments, we reared these insects in the artificial diet in plastic boxes at 25 ± 1 °C. We transferred 200 eggs to the surface of 120 g artificial diet (Southland Products Incorporated, USA) in each plastic box (10 × 10 × 9 cm). The hatched larvae dropped to the surface of the artificial diet and fed on it. Once individuals developed into 3rd or 4th instar larvae, pupae or adults, they were exposed to 10 °C for 24 h (to simulate gradually reduced temperatures in late autumn and allow a thermal acclimation) just before they were placed to the low-temperature regimes for overwintering tests. Overall, we obtained >7000 larvae and >8000 pupae for the laboratory experiment, and >8000 larvae, >8000 pupae, >4000 adults for the field experiment. We have compared the life history traits of insects reared on the artificial diet with natural host plants (cabbage leaves), they performed similarly (Peng and Li, unpublished data).Laboratory experiment of winter survivalSite selectionTo identify what factor determines winter survival under different winter thermal conditions, we conducted a laboratory experiment that simulated temperature regimes of 10 selected sites across a latitudinal gradient in China (Fig. 1, Supplementary Table 1) at which this species is known to occur and damage cruciferous crops during the growing season.Temperature treatmentTo simulate the winter temperatures in the 10 geographically distinct sites (Fig. 1a, Supplementary Table 1), we collected daily mean temperatures during winter (November to next April of 1966–2010) at each site from China Meteorological Data Service Centre (http://data.cma.cn/en). Then, we fitted a polynomial model to the temporal changes of winter daily mean temperatures for each site (Fig. 1b). To simplify the logistics of temperature control procedures, we set all temperature regimes in combinations of linear decline, horizontal maintenance and linear increase to mimic the polynomial changes of winter temperatures in the 10 sites, and adjusted temperature every 10 days as needed (Fig. 1c). We controlled the winter temperature changes of the 10 sites with climate chambers (RXZ-280B, Jiangnan Ltd., Ningbo, China) and refrigerators (Royalstar BCD-246GER) according to curves in Fig. 1c.Experimental protocolsWe conducted a winter survival experiment with 10 low-temperature regimes (Fig. 1c). We exposed 6050 larvae and 7150 pupae to 10 low-temperature regimes according to the experiment design (Fig. 1c). Then we sampled 55 larvae and 65 pupae every 10 days from each temperature regime resulting in 11 sampling points. Sampled larvae were placed at 25 °C for 1.5 days to observe the survival based on if their body kept fresh green60 and appendage moved after touching with a brush34. The pupae were placed at 25 °C, RH 70–80% and photoperiod of 16 L:8D for emergence to determine the survival (emergence rate). These samples were not returned to the temperature treatments. Thus, no individual was measured more than once and each sample interval represents an independent observation.Field survival experiments across 12 geographic sitesTo verify the cold survivals from the laboratory simulation and identify the best predictor under natural conditions, we conducted field experiments to explore the winter survival for multiple years at various geographic sites in China (Fig. 1a, Supplementary Table 1). The diamondback moth overwinters either in remaining cabbage plants or in fallen leaves (post-harvest conditions) in regions without standing cabbage crops in the winter. We tested the winter survival of larvae, pupae and adults in the caged cabbage plants or post-harvest conditions in fallen leaves on the soil surface at each site for 3–4 months. We transferred 30 larvae, 30 pupae or 30 adults from our stock rearing to a cabbage plant in the field. Then each plant was covered with a screen cage to avoid disturbance and contain focal individuals (see Supplementary Fig. 2). We set 6–8 cages for larvae, pupae and adults, respectively, in a field in November or early December. After an exposure of 1, 2, 3 and 4 months, we collected 2 cages of larvae, 2 cages of pupae and 2 cages of adults from the field at each sampling point and kept individuals in the laboratory (25 ± 1 °C, RH 65–75%, L:D = 16:8 h) for two days. We checked the survival status of the larvae based on the change in body coloration (i.e. if the larval body kept fresh green colour)60, pupal survival based on whether adults could emerge from the pupae, and adult survival based on if their appendage moved after touching with a brush.To simulate the field microenvironment of post-harvest conditions in winter, we filled half of a glass jar (diameter = 5.5 cm, height = 14 cm) with moist soil. Then, we transferred 30 larvae or 30 pupae to the soil surface, covered the insects with leaves, and then covered the glass jar with a nylon net (see Supplementary Fig. 2). We buried 6–8 jars for larvae and pupae, respectively, and kept the top of the jar at ground surface level at each site in November and early December. Because almost all adults died in few days within the jar, we did not test the adult survival in post-harvest conditions. After an exposure of 1, 2, 3 and 4 months, we took 2 jars of larvae and 2 jars of pupae per sampling period from the field and placed them in the laboratory with 25 ± 1 °C, RH 65–75%, L:D = 16:8 h for 2 days. The survival status of the larvae and pupae was checked with the same procedures as the overwintering tests on caged cabbage plants. Note that as in the standing plant experiment, no individual was tested more than once assuming that each observation is independent at the replicate level.Modelling and predicting winter survivalModel developmentOur goal was to identify key metrics that best predict the winter survival of the diamondback moth across a climatic gradient. To achieve this goal, we took several steps. First, we fit a set of predictive models to the laboratory experiments to identify which metric and model best describes survival under controlled conditions. We focused on three alternative predictors: the lowest daily mean temperature (MinDTmean), mean temperature (DTmean) combined with exposure days, and low-temperature degree-days (LTDD). The MinDTmean model assumes that survival can simply be predicted as a function of the lowest temperature an individual experienced during its exposure time. The DTmean model assumes that survival depends on both the average temperature individuals experience below the cold threshold for survival (11 °C)32 and exposure duration (note that exposure time varied systematically in 10-day increments). Finally, the LTDD model predicts survival depending on coldness below the cold threshold. We calculated LTDD by summing up negative deviations of daily mean temperatures from the cold threshold (11.0 °C) during each exposure period for each simulated geographic site (Fig. 1c). To detect potential relationships, we fit each model using three different functions, i.e. linear, exponential and sigmoid models to describe the survival probability (Supplementary Table 2, Fig. 1). We estimated parameters of models in SigmaStat 3.5 and compared model fit using R2 and AIC values (see detailed models in Supplementary Table 2).Field validation of survival modelsTo validate winter survival models derived from the laboratory (see models in Supplementary Table 2) for complex and variable field conditions, we compared model predictions to observed survivals in field experiments across 12 different geographic sites over 5 years (Fig. 1a, Supplementary Table 1). To make the connection, we first collected daily mean temperatures recorded at the nearest weather stations to our field sites from China Meteorological Data Service Centre. We then calculated MinDTmean, DTmean and exposure days, and LTDD for each site for each treated period and input these values into these laboratory models to predict winter survival. Note, that because the coefficients were calculated from the laboratory experiment, predictions are completely independent of survival observed under field conditions. During the model validation, we excluded the field data of south China, e.g. Guangzhou, Changsha and Wuhan where the warmer temperatures allowed moths to continue their regular life cycle during the whole winter, resulting in unrealistic winter survival. We also excluded replicates in which glass jars were filled with water and destroyed the tested insects. We used linear regression to compare predicted survival with field observations. The validity of each model was evaluated based on the variance explained, slopes of linear regressions and prediction bias (i.e. deviation from unity slope). Finally, we selected the exponential model driven by LTDD as the model to predict the global distribution of winter survival due to its lowest AIC value (Supplementary Table 2) and the least bias (Supplementary Table 3, Fig. 2) among all models.Global prediction of overwintering range shiftTo extrapolate our winter survival predictions to a global scale under present and future climate conditions, we downloaded global historical daily mean temperature data for 50 years (1967–2016) from Berkeley Earth (1° × 1° grid, http://berkeleyearth.org/data/). We added 1, 2, 3, 4, 5 and 6 °C to mean temperatures of 2012–2016, respectively, to represent the different future warming scenarios37. Then, we calculated the annual LTDD in the northern hemisphere with Eq. (1) and in the southern hemisphere with Eq. (2). For xi,j  x0, we excluded the xi,j for the calculation LTDD. For Eq. (1), we started the calculation of LTDD from July 1st (Julian date 182), ended on June 30th of next year (Julian date 181) to cover the whole low-temperature season in the northern hemisphere cross the calendar year. We used LTDD for every year during past conditions to our validated survival model (LTDD-dependent exponential model) and further calculated the expected corresponding yearly winter survival and 5-year mean survival. Since the diamondback moth only feeds on Brassicaceae plants61, we incorporated host availability to refine the pest distributions. We retrieved Brassicaceae occurrence data during 1967–2016 (3,720,971 records) from the Global Biodiversity Information Facility (GBIF) database (www.gbif.org), and excluded unknown and duplicate records; 919,808 records were retained to model the global distribution of host plants. We used a dataset of eight selected bioclimatic variables as described in a previous Brassicaceae biogeographic study62, including isothermality (bio3), temperature seasonality (bio4), min temperature of coldest month (bio6), mean temperature of wettest quarter (bio8), mean temperature of driest quarter (bio9), precipitation seasonality (bio15), precipitation of warmest quarter (bio18), precipitation of coldest quarter (bio19) from Worldclim dataset63 (http://worldclim.org). We ran the species distribution model using the Maxent algorithm in R package dismo64. Model outputs were presented in grid ranks of host plant presence probability from 0 (unsuitable) to 1 (most suitable). Based on the known distribution of Brassicaceae, we only included grid cells with Brassicaceae presence probability ≥0.3 for our final survival and distribution analysis to ensure the presence of the host plant and mapped them with Arcmap 10.2 (Environmental Systems Research Institute) (see Fig. 3a, e). To show spatial-temporal changes in the geographic distribution of winter survival, we quantified the historical change (expansions or contractions) in the overwintering range based on the total numbers of grids for each year between 1967 and 2016 relative to the baseline area in 1967 (see Fig. 3f) and further calculated average changes of every 5 years (see Fig. 3b). We selected sites in the overwintering marginal belt (with winter survival between 1 and 5%) in the baseline year (1967), calculated the annual LTDD of these sites from 1967 to 2016, and built the linear trend of annual LTDD for years 1967–2016 (see Fig. 3g). We predicted distribution changes for future scenarios (added 1, 2, 3, 4, 5 and 6 °C to the current mean temperatures of 2012–2016) relative to the baseline area of 1967–1971 (see Fig. 3c, d).Meta-analysis linking pesticide resistance to overwintering typeData preparation: literature search and selection criteriaWe performed a comprehensive literature survey to collect data on pesticide resistance of the diamondback moth worldwide. We searched for publications in databases of ISI Web of Science, Scopus and China National Knowledge Infrastructure (CNKI) using keywords “pesticide resistance” in combination with “diamondback moth” or “Plutella xylostella” and expanded references in the selected papers. We reviewed titles, abstracts and in many cases the full articles for relevance and agreement with our inclusion criteria. Studies were included if they (1) monitored the pesticide resistance of field populations, (2) used the leaf dip bioassay method to test pesticide resistance which is the most commonly used method recommended by Insecticide Resistance Action Committee (IRAC, http://www.irac-online.org); (3) provided resistance ratio of field populations. Resistance ratio (abbreviated as RR) is the magnitude of pesticide resistance and is commonly calculated by dividing the median lethal concentration (LC50) of a tested field population by LC50 of the susceptible population (without exposure to pesticide). The LC50 is commonly estimated from a concentration-mortality curve of a given pesticide. The preliminary literature search resulted in 2151 studies out of which 62 matched these criteria. A PRISMA diagram describing details of our literature search is available in Supplementary Fig. 4.Data preparation: data extractionWe extracted data from each selected publication, the names of pesticides, sampling locations and years of field populations, number of tested individuals in a bioassay, resistance ratio of field populations (RR), LC50 of field populations (LC50field) and susceptible populations (LC50susceptible), and 95% confidence intervals (CIs) of LC50field and LC50susceptible. Some studies generated results from multiple types of pesticides with the same field population, each of which was considered as a different entry. Finally, we gathered 1806 entries for pesticide resistance of field populations of the diamondback moth.Data preparation: calculation of the weighted effect sizeWe conducted a meta-analysis to test if pesticide resistance levels vary across different types of overwintering sites. To account for differences in sample sizes and variances in resistance ratios across studies, we calculated the corrected (weighted) resistance ratio for each study following the method in Hedges et al.65. We calculated the logarithm of resistance ratio (logRR) to present the effect size for each entry and further calculated the weighted effect size (wlogRR) by$${{{{{rm{wlogRR}}}}}}={{{{{rm{logRR}}}}}}times w$$
(3)
where w is the weighting factor of each entry, with w = 1/sqrt(VlogRR)66. To consider the contribution from both field and susceptible population, the pooled variance VlogRR was calculated as follows65:$${{{{{{rm{V}}}}}}}_{{{{{{rm{logRR}}}}}}}=frac{{{{{{{{rm{SE}}}}}}}_{{{{{{rm{field}}}}}}}}^{2}}{{n}_{{{{{{rm{field}}}}}}}times {{{{{{{rm{LC50}}}}}}}_{{{{{{rm{field}}}}}}}}^{2}}+frac{{{{{{{{rm{SE}}}}}}}_{{{{{{rm{susceptible}}}}}}}}^{2}}{{n}_{{{{{{rm{susceptible}}}}}}}times {{{{{{{rm{LC50}}}}}}}_{{{{{{rm{susceptible}}}}}}}}^{2}}$$
(4)
where LC50field and LC50susceptible, SEfield and SEsusceptible, nfield and nsusceptible, are LC50, the standard error of LC50 and sample size for field population and susceptible population, respectively. SEfield and SEsusceptible can be calculated from their own confidence intervals (95% CI)67:$${{{{{rm{SE}}}}}}=frac{{{{{{{rm{CI}}}}}}}_{{{{{{rm{upper}}}}}}{{{{{rm{limit}}}}}}}-{{{{{{rm{CI}}}}}}}_{{{{{{rm{lower}}}}}}{{{{{rm{limit}}}}}}}}{2times 1.96}$$
(5)
where CIupper limit is the upper limit and CIlower limit is the lower limit of the 95% CI for LC50.We used the prognostic method68 to estimate VlogRR for entries that miss either 95% CI or LC50 based on the average VlogRR of the other complete entries.Data preparation: potential moderator variablesSeveral factors could influence pesticide resistance besides overwintering temperatures. The effective temperature degree-days (ETDD) may change the annual number of generations, the intensity of pesticide application, and thus the selection stress47, e.g. between 7.4 and 33 °C for the diamondback moth32. In addition, the variety of pesticides used in a study may also affect the resistance levels through their mode of actions (the lethal mechanism) and cross-resistance69,70. To account for these potentially confounding factors, we collected the mode of action for each variety of pesticides from IRAC, and calculated LTDD, ETDD and overwintering type for each of the 1806 original records. We collected data for daily mean temperatures for each site from Berkeley Earth. For each location, we calculated the mean annual LTDD, ETDD and winter survival average across the 5 years before the sample. We split the sampling sites into three types based on predicted winter survival of the diamondback moth: (1) the permanent (overwintering) sites: locations with the mean winter survivals ≥5%, (2) marginal sites: locations with the mean winter survivals 1–5%, (3) transient (non-overwintering) sites: locations with the mean winter survivals More

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Field survey and samplingField data were collected from 130 study sites spanning a latitudinal gradient of 35.89−50.70° N and a longitudinal gradient of 76.62−122.41°E and covering five provinces across the temperate region in northern China (Xinjiang Autonomous Region, Qinghai Province, Gansu Province, Ningxia Autonomous Region, and Inner Mongolia Autonomous Region; Fig. 2a). Locations for the field study target natural drylands, delineated as areas with aridity level above 0.35 (ref. 30), and represent a large aridity gradient including dry-subhumid (N = 12), semiarid (N = 42), arid (N = 56), and hyperarid (N = 20) regions (Fig. 2a), which are highly vulnerable to expected increases in aridity with human activity and climate change33,71. The aridity level of each site was calculated as 1 – AI, where AI is the ratio of precipitation to potential evapotranspiration38. We obtained AI from the Global Aridity Index and Potential Evapotranspiration Climate database (https://cgiarcsi.community/). The selection of the field sites aimed to minimize the potential impacts of human activity and other disturbances on soil, vegetation, and geomorphological characteristics based on the following three criteria: (i) sites were at least 1 km away from major roads and >50 km from human habitations; (ii) sites were under pristine or unmanaged conditions without visible signs of domestic animal grazing, grass/wood collection, engineering restoration plantings, and infrastructure construction; and (iii) the soil was dry without experiencing rainfall events for at least 3 days prior to sampling. Collectively, our field survey involved a wide range of the abiotic and biotic features of dryland ecosystems across northern China. These sites encompass the 14 soil types, i.e., arenosols, calcisols, cambisols, chernozems, fluvisols, gleysols, greyzems, gypsisols, kastanozems, leptosols, luvisols, phaeozems, solonchaks, and solonetz, and the four main vegetation types44, i.e., typical grassland (dominated by Stipa spp., Leymus spp., Cleistogenes spp., and Agropyron spp.), desert grassland (dominated by Stipa spp., Cleistogenes spp., Suaeda spp., and Artemisia spp.), alpine grassland (dominated by Stipa spp., Leymus spp., Carex spp., and Festuca spp.), and desert (dominated by Reaumuria spp., Salsola spp., Calligonum spp., and Nitraria spp.). Elevation, mean annual temperature, and mean annual precipitation (1970–2000; https://www.worldclim.org/) of the sites varied from 204 to 3,570 m a.s.l. (mean, 1,294 m a.s.l.), from –4.3 to 12.8 °C (mean, 5.0 °C), and from 21 to 453 mm (mean, 195 mm), respectively (Supplementary Table 1).Field sampling was conducted between June and September from 2015 to 2017 (each site was visited once over this period) following well-established standardized protocols as described in refs. 13,34. In brief, three 30 m × 30 m quadrats were established at each site to represent the local vegetation and soil types that covered an area of no less than 10,000 m2. The cover of perennial vegetation was estimated and all perennial plant species were listed by walking steadily along four 1.5 m × 30 m parallel transects (spaced 8 m apart) located within each quadrat using the belt transect method72. Site-level estimate for perennial plant cover was obtained by averaging the values measured in the 12 transects established. After vegetation survey, we located five 1 m × 1 m (for typical grassland, desert grassland, and alpine grassland) or five 5 m × 5 m (for desert) plots within each quadrat (at each corner and the center of the quadrat) to measure site-level plant aboveground and root biomass (g m−2). In each 1 m × 1 m plot, all grasses and dwarf shrubs were harvested to ground level for measurement of aboveground biomass. Five soil cores (7 cm diameter; 0–40 cm depth) per 1-m2 plot were collected randomly, and the roots were removed using a 1-mm sieve and washed cleanly to measure root biomass. All shoot and root samples were dried to constant weight at 65 °C. In each 5 m × 5 m plot, we recorded the number of individuals per dominant shrub species and canopy cover and height of each individual, thereby estimating aboveground and root biomass according to the allometric models developed in previous studies that were conducted in the same regions as sampled here (see Supplementary Table 9 for details). Based on these measurements, we further estimated BNPP. However, BNPP is typically difficult to observe and measure, especially over large spatial scales and environmental gradients as in this study, because the root system is subject to simultaneous growth and turnover73,74. Across our survey areas, ~77–98% of the precipitation occurs between June and September (during the peak-growing season) corresponding to the period of the highest plant above- and belowground biomass34,35,41,75. Therefore, we argue that BNPP can be estimated approximately at each site by the following equation:$$frac{{{{{{rm{Aboveground}}}}}},{{{{{rm{biomass}}}}}}}{{{{{{rm{Root}}}}}},{{{{{rm{biomass}}}}}}}cong frac{{{{{{rm{Aboveground}}}}}},{{{{{rm{net}}}}}},{{{{{rm{primary}}}}}},{{{{{rm{productivity}}}}}},({{{{{rm{ANPP}}}}}})}{{{{{{rm{BNPP}}}}}}}$$
(1)
where both aboveground and root biomass are site-level measurements (g m−2). We used normalized difference vegetation index (NDVI) as a metric for ANPP as explained in recent studies in drylands14,33,70. NDVI data were obtained from the moderate resolution imaging spectroradiometer aboard NASA’s Terra satellites (https://neo.sci.gsfc.nasa.gov/). We used the average NDVI values during our sampling dates as a proxy for ANPP at the site level as described in ref. 14.Five soil cores (0–20 cm depth) per quadrat were then taken randomly under the canopies of the dominant perennial plant species and in bare areas devoid of perennial vegetation, respectively, and then were mixed as one sample for vegetation areas and the other sample for bare ground. When more than one dominant perennial plant species was observed, another three composite samples were collected under the canopies of co-dominant perennial plant species. All vegetation and soil surveys were carried out during the wet season (June to September) when biological activity and productivity are maximal; as such, we do not expect the different sampling times and years or seasonality to be a major factor influencing our conclusions. Collectively, 6–21 soil samples per site were collected, and in total 864 samples were taken and analyzed for each of the seven individual soil functions (see below) and multifunctionality. All soil functions evaluated in the field study were calculated at site level by using a weighted average of the mean values observed in vegetated areas and bare ground by their respective cover13,14,38. After field sampling, the visible pieces of plant material were removed carefully from the soil, which was sieved and divided into three portions. The first portion was air-dried and used for soil organic C, total N, total P, available P, and pH analyses. The second portion was immediately mixed with 2 M KCl and stored at 4 °C for soil ammonium and nitrate analyses. The third portion was immediately frozen at –80 °C for assessing soil microbial diversity.Microcosm experimentIn addition to the large-scale field study described above, we manipulated soil water availability in a microcosm experiment to evaluate the linkages between moisture content, soil microbial diversity, and multifunctionality. It is important to note that our intention is not to directly compare results between these two different approaches [i.e., in the field, measures of soil functions are related to nutrient pools, which we use to associate soil multifunctionality with both plant and soil microbial diversity, whereas in the microcosm experiment the measures of soil functions are related to process rates such as respiration rate and key enzyme activities (see below), which we use to associate soil multifunctionality with microbial diversity in the absence of plants]. Rather, by using an experimental microcosm approach, we aimed to complement the field study and thus further verify the potential increases in aridity to alter the relationship between soil microbial diversity and multifunctionality in the absence of plants. In parallel with the sampling protocols described above, we collected a greater mass of soil (c. 30 kg) under vegetation canopies from one site [i.e., Jingtai country (37.40°N, 104.26°E; Gansu Province, China)]. Soil type, mean annual temperature, mean annual precipitation, and aridity level (1970–2000; https://www.worldclim.org/) of the site is calcisols, 7.9 °C, 205 mm and 0.81, respectively. Following field sampling, the soil was stored immediately at 4 °C until subsequent processing in the laboratory.In brief, a total of 30 experimental microcosms composed of 10 moisture levels with three replicates were established under sterile conditions in a closed incubation chamber (Supplementary Fig. 1a). Each microcosm was filled with 1 kg of soil. These microcosms were incubated at 18.5 °C [the annual mean land surface temperature (1981–2010) for the sampling site; http://data.cma.cn/en], and moisture contents were adjusted and artificially maintained at the ten levels respectively equivalent to 3, 5, 8, 10, 20, 40, 60, 80, 100, and 120% field capacity (27.6%) during the duration of the experiment for 30 days. The corresponding moisture content (%) measured at the end of the experiment varied from 2.03 ± 0.034 to 33.57 ± 1.94, which matched well with differences in moisture conditions among a subset of field soil samples (N = 521; Supplementary Fig. 1b). After incubation, the soil was removed from each microcosm; a portion of the soil was immediately frozen at –80 °C for molecular analysis, and the other fraction was air-dried, sieved, and stored at –20 °C for assessing multiple soil functions as described below.DNA extraction, PCR amplification, and amplicon sequencingFor both the field and experimental studies, we assessed the diversity of soil archaea, bacteria, and fungi using Illumina-based sequencing. Genomic DNA was extracted from 0.5 g of each defrosted soil sample (N = 864 for the field study and N = 30 for the experimental study) using the PowerSoil® DNA Isolation Kit (MO BIO Laboratories, USA) according to the manufacturer’s instructions. For our field study, extracted DNA was pooled at site level, ultimately resulting in 130 composite DNA samples under canopies of vegetation and in bare ground, respectively. Pooling DNA samples may outperform the commonly used method that extracts genomic DNA from mixed soil samples, which could remove large amounts of information on the diversity of soil microorganisms14,22. Negative controls (deionized H2O in place of soil) underwent identical procedures during the extraction to ensure zero contamination in downstream analyses.The V3−V5 regions of the archaeal 16S rRNA gene were amplified using the primer pair Arch344F and Arch915R. Thermal conditions were composed of an initial denaturation of 3 min at 95 °C, ten cycles of touchdown PCR (95 °C for 30 s, annealing temperatures starting at 60 °C for 30 s then decreasing 0.5 °C per cycles, and 72 °C for 1 min), followed by 25 cycles at 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 1 min, with a final extension at 72 °C for 10 min. The primer pair 338F and 806R was used for amplification of the V3−V4 regions of the bacterial 16S rRNA gene. Thermocycling conditions consisted of 3 min at 95 °C and then subjected to 30 amplification cycles of 30 s denaturation at 95 °C, 30 s annealing at 55 °C, followed by 72 °C for 45 s, and a final extension of 72 °C for 10 min. The fungal internal transcribed spacer (ITS) region 1 was amplified using the primer pair ITS1F and ITS2. The amplification conditions involved denaturation at 95 °C for 3 min, 35 cycles of 94 °C for 1 min, 51 °C for 1 min, and 72 °C for 1 min and a final extension at 72 °C for 10 min. Details of primers for each microbial taxa were given in Supplementary Table 10. These primers contained variable length error-correcting barcodes unique to each sample. All amplification reactions were performed in a total volume of 20 μl containing 4 μl of 5× FastPfu Buffer, 2 μl of 2.5 mM dNTPs, 0.8 μl of both the forward and reverse primers, 10 ng of template DNA, and 0.4 μl of FastPfu DNA Polymerase (TransGen Biotech., China). To mitigate individual PCR reaction biases each sample was amplified in triplicate and pooled together. All PCRs were done with the ABI GeneAmp® 9700 Thermal Cycler (Thermo Fisher Scientific, USA). PCR products were evaluated on 2.0% agarose gel with ethidium bromide staining to ensure correct amplicon length, and were gel-purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, USA). Purified amplicons were combined at equimolar concentrations and paired-end sequenced (2 × 300 bp) on an Illumina MiSeq platform (Illumina, USA) at the Majorbio Bio-pharm Technology Co., Ltd. (Shanghai, China) according to standard protocols.Sequence processingInitial sequence processing was conducted with the QIIME pipeline76. Briefly, reads were quality-trimmed with a threshold of an average quality score higher than 20 over 10 bp moving-window sizes and a minimum length of 50 bp. Paired-end reads with at least 10 bp overlap and 2 indicate that the models are different; Supplementary Table 2]. We further assessed whether soil multifunctionality responded more rapidly to aridity than did any individual soil functions. To this end, we explored the presence of aridity thresholds for those relationships that were better fitted by nonlinear regressions (Fig. 2b–i) using the standard protocols developed in ref. 33. The presence of an aridity threshold means that once an aridity level is reached, a given variable either changes abruptly its value (i.e., discontinuous threshold) or its relationship with aridity (i.e., continuous threshold). Hence, a lower aridity threshold indicates that a given variable is more vulnerable to increasing aridity than are others33. We further fitted step (a linear regression that modifies only intercept at a given aridity level) and stegmented (showing changes both in intercept and slope at a given aridity level) regressions for the determination of discontinuous thresholds, and segmented (exhibiting changes only in slope at a given aridity level) regressions for continuous thresholds. Each of these models yields a change point (i.e., threshold) describing the aridity level that evidences the shift in a given nonlinear relationship evaluated. We also used AIC to choose the best threshold model and the corresponding threshold in each case (Supplementary Table 2).We then employed analysis of variance based on type-I sum of squares in a linear mixed-effects model (Eq. (2); Table 1) to test the relationships between the multiple biotic (BNPP, plant species richness, and the soil microbial diversity index) and abiotic (aridity, soil pH, and soil clay content) factors and soil multifunctionality:$${{{{{rm{Soil}}}}}},{{{{{rm{multifunctionality}}}}}} sim; {{{{{rm{Year}}}}}}+{{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}\ quad+,{{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}\ quad+{{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}times {{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}+{{{{{rm{Aridity}}}}}}\ quad+,{{{{{rm{Aridity}}}}}}times {{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}\ quad+,{{{{{rm{Aridity}}}}}}times {{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}+{{{{{rm{BNPP}}}}}}+{{{{{rm{Soil}}}}}},{{{{{rm{pH}}}}}}+{{{{{rm{Soil}}}}}},{{{{{rm{clay}}}}}},{{{{{rm{content}}}}}}\ quad+,{{{{{rm{Elevation}}}}}}+{{{{{rm{Latitude}}}}}}+{{{{{rm{Longitude}}}}}}+(1|{{{{{rm{Soil}}}}}},{{{{{rm{type}}}}}})+(1|{{{{{rm{Vegetation}}}}}},{{{{{rm{type}}}}}})$$
(2)
where × indicates an interaction term. We obtained information on soil clay content (%) from the SoilGrids system (https://soilgrids.org/), and eliminated variation due to different sampling years by first entering the term “Year” into the statistical model41. The elevation, latitude, and longitude of the study sites were included to account for the spatial structure of our dataset13,70. To account for the similarities of soil and vegetation types among study sites we included “Soil type” and “Vegetation type” as random terms.We further simplified the Eq. (2) to focus only on the relationships between aridity, biodiversity, and soil multifunctionality (Eq. (3); Supplementary Fig. 5). We did so because excluding additional biotic and abiotic factors did not change qualitatively the main results presented here (Table 1 and Supplementary Fig. 5), and therefore we used the simplest model to test our hypotheses more clearly. Our simplified model was:$${{{{{rm{Soil}}}}}},{{{{{rm{multifunctionality}}}}}} sim {{{{{rm{Year}}}}}}+{{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}\ quad+,{{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}\ quad+,{{{{{rm{Aridity}}}}}}+{{{{{rm{Aridity}}}}}}times {{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}\ quad+,{{{{{rm{Aridity}}}}}}times {{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}\ quad+,{{{{{rm{Aridity}}}}}}times {{{{{rm{Plant}}}}}},{{{{{rm{species}}}}}},{{{{{rm{richness}}}}}}times {{{{{rm{Soil}}}}}},{{{{{rm{microbial}}}}}},{{{{{rm{diversity}}}}}},{{{{{rm{index}}}}}}\ quad+,(1|{{{{{rm{Soil}}}}}},{{{{{rm{type}}}}}})+(1|{{{{{rm{Vegetation}}}}}},{{{{{rm{type}}}}}})$$
(3)
To evaluate how the biodiversity–multifunctionality relationships varied along aridity gradients, we conducted a moving-window analysis as detailed in ref. 69. Briefly, we performed the linear mixed-effects model described in Eq. (3) for a subset window of 60 study sites with the lowest aridity values (this number of sites provided sufficient statistical power for our model), and repeated the same calculations as many times as sites remained (i.e., 70). We then bootstrapped the standardized coefficients of each fixed term within each subset window, which was matched to the average value of aridity across the 60 sites. We fitted linear and nonlinear regressions to the bootstrapped coefficients of biodiversity and its interaction with aridity along aridity gradients (Fig. 3a, b and Supplementary Table 2), and identified the aridity thresholds for the changes in the coefficients of biodiversity (Fig. 3a and Supplementary Table 2) using the same procedure already described above. To provide further support for the aridity thresholds identified here, we also assessed the significance of the bootstrapped standardized coefficients of biodiversity and its interaction with aridity at 95% confidence intervals for each subset window (Fig. 3e). Before fitting threshold regressions, we evaluated whether the variables followed either a unimodal or bimodal distribution using the fitgmdist function in MATLAB (The MathWorks Inc., USA). Our results showed that all variables used for threshold detection presented unimodal distributions (Supplementary Table 11), suggesting that the three threshold regressions mentioned above (i.e., segmented, step, and stegmented) are appropriate in all cases33. We used the chngpt and gam packages in R (http://cran.r-project.org/) to fit segmented/step/stegmented and GAM regressions, respectively. To further check the validity of the thresholds identified, we bootstrapped linear regressions at both sides of each threshold for each variable. We then used the nonparametric Mann–Whitney U-test to compare the slope and the predicted value evaluated before and after each threshold. In all cases, we found significant differences in both of these two parameters (Fig. 3c, d and Supplementary Figs. 2, 3, 6).Given a clear shift in the relationships between plant or microbial diversity and soil multifunctionality occurring at a threshold around an aridity level of 0.8 (Fig. 3), we further used OLS regressions to clarify the relationships between each component of plant or microbial diversity and soil multifunctionality in less and more arid regions separately, as well as across all sites (Fig. 4). To do so, we split our study sites into two groups: sites with aridity 0.8 (more arid regions; N = 76). Moreover, we fitted the mixed-effects model described in Eq. (2) for less and more arid regions separately to ensure the robustness of these bivariate correlations when accounting for multiple biotic and abiotic factors simultaneously, with the exception of using all components of microbial diversity metrics (i.e., soil archaeal, bacterial, and fungal richness) instead of the soil microbial diversity index in the models (Supplementary Table 4). All linear mixed-effects models were performed using the R package lme4. We used a variance inflation factor (VIF) to evaluate the risk of multicollinearity, and selected variables with VIF  0.05). Finally, we also used SEMs to compare the hypothesized direct and indirect relationships between moisture content, microbial diversity, and soil multifunctionality at low and high moisture levels (see an a priori model in Supplementary Figs. 17b and 31c, d). Test of goodness-of-fit for SEMs were same as described above. All the SEM analyses were conducted using AMOS 21.0 (IBM SPSS Inc., USA). Data and code used to perform above analyses are available in figshare98.Reporting SummaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

Collection of the samplesTen (10) wood pellet samples were purchased from a different location in B&H, of known suppliers from the market (supermarkets, garden shops, and gas stations). The samples were accompanied by a declaration describing that nine of them were originated from B&H, and one of them was from Italy. Characteristics of collected wood pellet samples (type of wood, energetic value, declared moisture, declared and determined ash amount) are listed in Table 1. All of the samples were analyzed for moisture and ash content. Additionally, in ash samples of mentioned wood pellets, heavy metal concentration (Cd, Co, Cr, Cu, Fe, Mn, Ni, Pb, and Zn) was determined.Table 1 Characteristic of analyzed samples wood pellets.Full size tableAll pellet samples were originated from B&H, purchased from different cities, often used for house heating, instead of sample S3 which was from Italy.Ash determination of wood biomass samplesThe wood pellet samples were oven-dried at 105 °C for 24 h. The content of ash was determined by gravimetric method according to the procedure published by Pan and Eberhardt18 as follows: pellet samples, 1 g (± 0.1 mg) of each was weighed into a previously annealed ceramic pot (m1) and burned in a muffle furnace (Nabertherm) for one hour at 300 °C, following by increasing the temperature to 400 °C for one hour more and then burning the samples for next six hours at 550 °C. The procedure is repeated until a constant mass (m2) was reached. The ash content is determined by the Eq. (1):$${text{Ash content}}, % = frac{{{text{(m}}_{2} – {text{m}}_{{1}} {)}}}{{{text{m}}_{{{text{sample}}}} }} times {100 }{text{.}}$$
(1)
Preparation of samplesThe chemical determinations of the heavy metals in wood pellet ashes (Table 2) were made by wet digestion by soaking the samples in 25 mL of 65% HNO3 in polytetrafluoroethylene (PTFE) vessels. After evaporation of the nitrogen oxides, the vessels were closed and allowed to react for 14 h at 80 °C, following by cooling to room temperature. Then, the digest was filtered, transferred to a 25 mL volumetric flask, and filled up with redistilled water to the mark. All samples and blank were prepared in three replicates19,20,21.Table 2 Heavy metal concentrations (mg kg−1 d.w.) in the wood pellet ashes.Full size tableHeavy metal analysisMetal analyses in ash samples of mentioned wood pellets were performed using a flame atomic absorption spectrometry (Varian AA240FS) for Mn, Fe, Pb, and Zn and graphite furnace (Varian AA240Z) for Cd, Co, Cr, Cu, and Ni. A blank probe was prepared using the same digestion method to avoid the matrix effect. Standard metal solutions used for the calibration graphs were prepared by diluting 1000 mg L−1 stock single-element atomic absorption standard solutions of Cd, Co, Cr, Cu, Fe, Mn, Ni, Pb, and Zn (Certipur Grade, Merck, Germany). Linear calibration graphs with correlation coefficients  > 0.99 were obtained for all analyzed metals. The accuracy of the method was evaluated using the standard reference materials: Fine Fly Ash (CTA-FFA-1, Institute of Nuclear Chemistry and Technology Poland) and Fly Ash from pulverized coal (BCR-038, Institute of reference materials and measurements-IRMM, Belgium). The obtained results were in the range of the reference materials. The detection limit (LOD) and limit of quantification (LOQ) for the nine analyzed metals were calculated based on Xb + 3 SDb and Xb + 10 SDb, respectively, where Xb is the mean concentration of the blank sample (n = 8) and SDb is the standard deviation of the blank for eight readings22. The values of the LOD were: Cd (0.61 µg L−1), Co (0.49 µg L−1), Cr (0.67 µg L−1), Cu (20.10 µg L−1), Fe (83.85 µg L−1), Mn (6.42 µg L−1), Ni (1.12 µg L−1), Pb (23.77 µg L−1), Zn (58.68 µg L−1), and LOQ values were: Cd (1.25 µg L−1), Co (1.41 µg L−1), Cr (1.42 µg L−1), Cu (47.66 µg L−1), Fe (111.2 µg L−1), Mn (16.14 µg L−1), Ni (2.70 µg L−1), Pb (47.73 µg L−1) and Zn (71.05 µg L−1).Pollution evaluationThe metal pollution index (MPI) as the geometric mean of the concentration of all metals found in ashes of wood samples was calculated by the following Eq. (2)23:$${text{MPI}} = left( {{text{C}}_{1} cdot {text{C}}_{2} cdot cdots {text{C}}_{{text{k}}} } right)^{{1/{text{k}}}} ,$$
(2)
where C1 is the concentration value of the first metal, C2 is the concentration value of the second metal, Ck is the concentration value of the kth metal.Evaluation of the presence and the grade of anthropogenic activity were demonstrated through the calculation of the enrichment factor (EF), widely used in environmental issues24. To understand which elements were relatively enriched in the different wood pellet ash samples, the heavy metal enrichment factor was calculated relative to soil values according to Eq. (3)25.$${text{EF}} = frac{{left( {frac{{{text{C}}_{{text{k}}} }}{{{text{E}}_{{{text{ref}}}} }}} right)_{{{text{ashes}}}} }}{{left( {frac{{{text{C}}_{{text{k}}} }}{{{text{E}}_{{{text{ref}}}} }}} right)_{{{text{soil}}}} }},$$
(3)
where Ck is the concentration of the element in the sample or the soil, Eref the concentration of the reference element used for normalization. A reference element is an element commonly stable in the soil characterized by the absence of vertical mobility and/or degradation phenomena. As in many studies as a reference element were Fe, Al, Mn, Sc, or total organic carbon used26,27. Therefore Fe has been chosen as reference material in this study. Iron is one of the major constituents of soil, as well as the average chemical constituent of the upper continental crust26.Health risk assessmentThe general exposure equations used in this study were adapted according to the US Environmental Protection Agency guidance28,29,30. The daily exposure (D) to heavy metals via wood pellet ash was calculated for the three main routes of exposure: (i) direct ingestion of ash particles (Ding); (ii) inhalation of suspended particles via mouth and nose (Dinh); and (iii) dermal absorption to skin adhered ash particles (Ddermal). Equations (4) to (6) were used to calculate exposure via ingestion, inhalation, and dermal route, respectively22,31.$${text{D}}_{{{text{ing}}}} = {text{ C }} cdot frac{{{text{ IngR }} cdot {text{ EF }} cdot {text{ ED}}}}{{{text{BW }} cdot {text{ AT}}}}{ } cdot {text{CF}}1{, }$$
(4)
$${text{D}}_{{{text{inh}}}} = {text{ C }} cdot frac{{{text{ InhR}} cdot {text{ EF }} cdot {text{ ED}}}}{{{text{PEF }} cdot {text{ BW }} cdot {text{ AT}}}}{, }$$
(5)
$${text{D}}_{{{text{dermal}}}} = {text{ C }} cdot frac{{{text{ SA }} cdot {text{ SL }} cdot {text{ABS }} cdot {text{EF }} cdot {text{ ED}}}}{{{text{BW }} cdot {text{ AT}}}}{ } cdot {text{CF}}1{, }$$
(6)

where c (mg kg−1) is the heavy metals concentrations in ash samples; IngR (mg day−1) is the conservative estimates of dust ingestion rates, 50 for adults, 200 for children30,32; InhR (m3 h−1) is the inhalation rate, 2.15 for adults, 1.68 for children32; EF (h year−1) is the exposure frequency, 1225 for adults and children22; ED (years) is the exposure duration, 70 for adults, 6 for children22; BW (kg) is the body weight, 80 for adults, 18.60 for children32; AT (days) is the averaging time, 25,550 for adults, 2190 for children22; PEF is the particle emission factor (m3 kg−1), 6.80 × 108 for adults and children31; SA (cm3) is the exposed skin area, 6840 for adults, 2550 for children32; SL (mg cm−2) is the skin adherence factor, 0.22 for adults, 0.27 for children32; ABS is the dermal absorption factor, 0.001 for adults and children31; CF1 is the unit conversation factor, 10–6 for adults and children22.The potential non-carcinogenic risk for each metal was estimated using the Hazard coefficient (HQ), as suggested by US EPA33. The HQ under various routes of exposure such as ingestion (HQing), inhalation (HQinh), and dermal (HQdermal) was calculated as a ratio of daily exposure (D) to reference dose of each metal (RfD) according to Eq. (7)32.$${text{HQ}}_{{text{k}}} = frac{{{text{D}}_{{text{k}}} }}{{{text{RfD}}}},$$
(7)

where k is ingestion, inhalation, or dermal route. The total hazard index (HI) of heavy metal for all routes of exposure was calculated as a sum of HQing, HQinh, and HQdermal as given in Eq. (8)34.$${text{HI}} = {text{ HQ}}_{{text{ing }}} + {text{ HQ}}_{{text{inh }}} + {text{ HQ}}_{{text{dermal }}} .$$
(8)
The carcinogenic risk (Risk) for potential carcinogenic metals was calculated by multiplying the doses by the corresponding slope factor (SF), as given in Eq. (9)35. The carcinogenic oral, inhalation, and dermal SF, as well as dermal absorption toxicity values, were provided from the Integrated Risk Information System30. The reference doses for Pb were taken from the Guidelines for Drinking Water Quality published by the World Health Organization36.$${text{Risk}} = { }mathop sum limits_{{{text{k}} = 1}}^{{text{n}}} {text{D}}_{{text{k}}} cdot {text{ SF}}_{{text{k}}} ,$$
(9)
where SF is the cancer slope factor for individually metal and k route of exposure (ingestion, inhalation, or dermal path). The total cancer risk (Risktotal) of potential carcinogens was calculated as the sum of the individual risk values using the following Eq. (10).$${text{Risk}}_{{{text{total}}}} = {text{Risk}}_{{{text{ing}}}} + {text{Risk}}_{{{text{inh}}}} + {text{Risk}}_{{{text{dermal}}}} .$$
(10) More

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NEWS AND VIEWS
08 September 2021

Pollination advantage of rare plants unveiled

An analysis of plant–pollinator interactions reveals that the presence of abundant plant species favours the pollination of rare species. Such asymmetric facilitation might promote the coexistence of species in diverse plant communities.

Marcelo A. Aizen

 ORCID: http://orcid.org/0000-0001-9079-9749

0

Marcelo A. Aizen

Marcelo A. Aizen is at the Research Institute for Biodiversity and the Environment (INIBIOMA), National University of Comahue – CONICET, 8400 San Carlos de Bariloche, Río Negro, Argentina, and at the Institute for Advanced Study, Berlin, Germany.

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Species diversification results from the balance between the formation of new species (speciation) and the loss of existing ones (extinction). The tremendous proliferation of different life forms on Earth can be attributed to both high rates of speciation and low rates of extinction. Flowering plants — a group called angiosperms — are one of the most diverse groups of non-mobile organism. There are approximately 352,000 plant species, nearly 90% of which depend, to various extents, on insects and other animals for pollination and seed production1. These animal pollinators have been key to the unstoppable diversification of the angiosperms, starting at least 120 million years ago, with pollinators promoting speciation by acting as potent selection agents for a plethora of flower traits2,3. Pollinators also aid species persistence by enabling pollen transfer between relatively distant individuals in sparse plant populations4. Writing in Nature, Wei et al.5 report that, for plant species that flower at the same time, pollinators mediate interactions that might facilitate species coexistence in diverse plant communities.

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doi: https://doi.org/10.1038/d41586-021-02375-z

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