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    Carbon response of tundra ecosystems to advancing greenup and snowmelt in Alaska

    Study sites: site description and climatic limitIn this study, we focused on seven flux tower sites located in Alaska, United States, including six AmeriFlux sites and one site of the Korea Polar Research Institute (KOPRI) (Fig. S1A, Table S1). Over the seven study sites, the annual mean temperature was between −10.09 and −0.55 °C and the annual total precipitation ranged from 287 to 540 during 2001–2018 based on the North American Regional Reanalysis (NARR41, 0.3-degree resolution every 3 h). The annual mean temperature increased from 2001 to 2018 at all sites, with rates between 0.5 and 2.2 °C per decade (p  0.05 at all sites). Our study sites were mostly dominated by wet sedges, grasses, moss, lichens, and dwarf shrubs. For example, the dominant plants at the US-Atq site (at a higher latitude) are herbaceous sedges (Carex aquatilis, Eriophorum russeolum, and Eriophorum angustifolium) and shrubs (Salix rotundifolia), with abundant mosses (Calliergon richardsonii and Cinclidium subrotundum) and lichens (Peltigera sp.)11. At the KOPRI site (at a lower latitude), mosses (Sphagnum magellanicum, Sphagnum angustifolium, and Sphagnum fuscum), lichens (Cladonia mitis, Cladonia crispata, and Cladonia stellaris), and tundra tussock cottongrass (Eriophorum vaginatum) are abundant39. The active layer thickness is between 0.33 and 1.0 m, according to field data and radar-based estimates.Climatic limits imposed by temperature, water, and radiation were quantified following Nemani et al.42 at each site during the GS between 2001 and 2018 (Fig. S2) using the NARR data. In this study, we defined the GS as from May to Oct., early GS as between May and Jun., peak GS as between Jul. and Aug., and late GS as between Sep. and Oct. For a temperature limit scalar, the monthly mean temperature from −5  to 5 °C was linearly scaled between 100% (i.e., no growth) and 0% (i.e., no reduction in growing days). The monthly ratio of precipitation to potential evapotranspiration (PET by the Priestley-Taylor method43), ranging between 0 and 0.75, was linearly scaled from 100 to 0% as a water limit scalar. A radiation limit scalar was estimated as a 0.5% reduction in growing days for every 1% increase in monthly cloudiness above the 10% threshold (monthly cloudiness (n) was estimated44 as (R={R}_{0}(1-0.75{n}^{3.4})), where R and R0 are the monthly mean incoming radiation and clear-sky radiation45, respectively).The carbon flux response to climatic variations at each site was further analyzed using a forward stepwise multiple regression analysis11 between the NEE and meteorological variables (temperature, PAR, and VPD) using tower data during the GS. Interaction terms among the variables are also included to consider the convolved effects of the variables (Eq. (1)).$${Y}_{{{NEE}}}= {beta }_{0}+{beta }_{1}{X}_{{{T}}}+{beta }_{2},{X}_{{{VPD}}}+{beta }_{3}{X}_{{{PAR}}}+{beta }_{4}{X}_{{{T}}}* {X}_{{{VPD}}}+ldots$$$$qquad;;; {beta }_{5}{X}_{{{T}}}* {X}_{{{PAR}}}+{beta }_{6}{X}_{{{VPD}}}* {X}_{{{PAR}}}+{beta }_{7}{X}_{{{T}}}* {X}_{{{VPD}}}* {X}_{{{PAR}}}$$
    (1)
    where YNEE is the daily average NEE (µmol m−2 s−1), and XT, XVPD, and XPAR are daily average air temperature (°C), VPD (ha), and PAR (µmol Photon m−2 s−1), respectively. Regression coefficients (({beta }_{0},ldots ,{beta }_{7})), standard errors, significance (P-value), and R2 value of the final regression model are summarized in Table S2.MODIS: long-term trends of snowmelt and greenup timingsWe collected the gridded MODIS snow cover (MOD10A1.V00646 at a 500-m resolution every day) and phenology (MCD12Q2.V00634 at a 500-m resolution yearly) from the NASA Earthdata (https://earthdata.nasa.gov/). We estimated the snowmelt and snowpack timings at each site as the date when a logistic fit to the MODIS snow cover (quality flags of good and best) passed 0.1 each year. We rejected those snowmelt timings when the gaps in the daily MODIS snow cover were longer than 2 weeks around the time of the snowmelt event. The greenup and dormancy timings with a quality flag of best were taken from the MODIS phenology. Based on the spatial representativeness assessment (see Supplementary Note), we decided to use the snowmelt timing and greenup timing within a 1 × 1 pixel window.The significance of the long-term trends in greenup and snowmelt timings at each site was determined by Spearman’s rho and Mann-Kendall tests (Fig. S3). We further estimated the 95% confidence intervals of the trends from 3000 timing sets generated by bootstrap resampling from a normal distribution47 (mean equal to each greenup or snowmelt timing with three standard deviation set to 10 or 6.6 days, respectively, i.e., the root-mean-squared values between the ground data-based estimates and MODIS values in a 1 × 1 pixel window, Figs. S9 and S10).SGSI: snowmelt-growing season indexGrowing season index (GSI)48 is one of the novel phenology models49 and has been widely applied for the phenological representations of various ecosystems50,51. GSI is a product of three indices of climatic variables (Eq. (2), Fig. S5): daylength, VPD, and growing-degree-days (GDD)52. As a phenological measure for a given meteorological condition, we calculated the daily GSI for spring (from Jan. 1 to Jul. 31) and fall (from Aug. 1 to Dec. 31), respectively. For the spring-GSI, GDD is the degree sum when the daily mean temperature rises above −5 °C after Jan. 1. For the fall-GSI, GDD is the degree sum when the daily mean temperature falls below 20 °C after Aug. 1. We then revised the GSI by multiplying it by a snowmelt index (iS) and referred to this as the snowmelt-GSI (SGSI, Eq. (3), Fig. S5). This guarantees that vegetation greenup does not start unless snow is melted, even if the meteorological conditions are sufficient to trigger leaf-out. The iS was estimated to be 0 when the snow cover fraction (snowfac variable53 in ED2) was above 0.1 and 1 otherwise.$${{{GSI}}}={{iX}}_{1} times {{iX}}_{2} times {{iX}}_{3}$$
    (2)
    $${{{SGSI}}}={{{GSI}}} times {iS}$$
    (3)
    where iX (X1, X2, and X3 represent daylength, VPD, and GDD, respectively) is 0 (X ≤ Xmin), 1 (X ≥ Xmax), and (X − Xmin)/(Xmax − Xmin) otherwise. Xmax and Xmin are the maximum and minimum thresholds of each index, respectively. For the spring-GSI, Xmin was calculated as the minimum value among the values on the greenup day (from MCD12Q2.V006) for the study period of 2001–2018 at each study site, and Xmax was calculated as the minimum value among the values on the maturity day. For the fall-GSI, similarly, Xmin was the minimum value for the dormancy timings, and Xmax was the minimum value for the senescence timings. We incorporated GSI (or SGSI) into ED2 by multiplying it to the optimal value of leaf biomass on the day, where it operates as an upper limit of the leaf biomass.In this study, it was assumed that phenological stages are driven by meteorological conditions, not by other factors (e.g., no assumption of fixed phenological periods6,54,55). The development of a robust phenological model for the tundra ecosystem would be enabled by an increasing amount of ground-based phenology data (e.g., PhenoCam data37).Case study: flux data analysisThere were three sites where flux data is available for >5 years in Alaska; US-Atq site (flux data during 2004–2008 with delayed snowmelt in 2005), US-EML site (flux data during 2009–2017 with delayed snowmelt in 2017), and US-BZF (flux data during 2012–2018 with delayed snowmelt in 2017 and 2018). We first calculated two timings that are related to the meteorological conditions (0.1-GSI timing and half-max GSI timing51, Fig. S6) using the NARR data. The 0.1-GSI timing and half-max GSI timing were calculated on the day when the GSI passed 0.1 and the half-max value (i.e., 0.5), respectively, each year. To calculate two timings regarding the flux seasonal profile51 (source-sink transition timing and half-max productivity timing, Fig. S6), we used 30-min gap-filled FLUXNET201556 data (NEE and GPP; quality flags of measured or good) at the US-Atq site to calculate daily NEP (i.e., negative NEE) and daily GPP. At the US-EML and the US-BZF sites, we applied an open-source code called ONEFlux (Open Network-Enabled Flux processing pipeline for eddy-covariance data)57 using the ERA5 data (European Centre for Medium-Range Weather Forecast Reanalysis v558) which was downscaled with a quantile mapping method59. Using the gap-filled 30-min NEP and GPP data from the ONEFlux, we calculated the corresponding daily values and fitted a smoothing spline to the daily NEP and the daily GPP each year. The source-sink transition timing was defined as the day when the smoothing spline of the daily NEP passed zero in each year. The half-max productivity timing was set to the day when the smoothing spline of the daily GPP passed the half-max value in that year51.Further, we investigated whether the delayed snowmelt altered the relationships between meteorological conditions and the flux-threshold timings at each site based on (1) the correlation between the 0.1-GSI timing and the source-sink transition timing and (2) the correlation between the half-max GSI timing and the half-max productivity timing.ED2: model implementationWe used NARR data41 (0.3-degree resolution every 3 h; temperature, precipitation rate, pressure, v- and u-wind speed, downward longwave and shortwave radiation flux, and relative and specific humidity) for single-point ED2 implementation at each study site from 2001 to 2018. Vegetation structure (LAI, leaf and structural biomass, diameter at breast height, and population density) was initialized for each site by using the maximum annual LAI of cold-adapted shrubs and Arctic C3 grass from the Ent Global Vegetation Structure Dataset v1.0b (Ent-GVSD v1.0b) with the allometric equations in ED2. Ent-GVSD v1.0b provides plant functional types (from the MODIS land cover, MCD12C1.V00560) and maximum annual LAI values (from the MODIS LAI, MOD15A2.V00461,62) in subgrid cover fractions. We did not use canopy heights from Ent-GVSD v1.0b because of the absence of trees at the study sites. Soil texture (the ratio of sand:silt:clay) was set following the Harmonized World Soil Database v1.163 of the Food and Agriculture Organization of the United Nations (UN FAO). Soil carbon was initialized using the UN FAO Global Soil Organic Carbon Map64, and soil nitrogen was estimated using the soil C/N ratio of moist tundra (mean: 18.4)65.The prior distribution of each key variable was based on previous studies (Table S4), and 10,000 parameter sets were randomly generated from the prior distributions (the so-called Monte Carlo method). The best parameter set was selected based on statistical measures (r2 and root-mean-squared error) when compared to the data at the US-Atq site, i.e., NEP flux data for 2004–2006 and MODIS LAI data for 2003–2010 (MCD15A3H.V00666 at a 500-m resolution every 4 days) (Table S5). We then validated the performance of ED2 with this best parameter set by focusing on key ecosystem processes, such as NEP, ecosystem respiration, soil temperature, snowmelt timing, greenup timing, and the LAI at all sites (Table S5). The ED2 LAI was overestimated by 0.15–0.16 compared to the field-measured LAI values (Jul.–Aug. in 2006 at Barrow67 and Jun.–Aug. in 1996 at Toolik68).It is worth noting that the accuracy of the MODIS LAI has not been extensively evaluated at high latitudes because of limited ground measurements and few valid MODIS data points due to inadequate sun-sensor geometry, illumination conditions, and cloud contamination69,70. Furthermore, the heterogeneous landscapes of the region at the scale of remote sensing data (from hundreds of meters to a few kilometers) are also a major challenge that must be addressed before the data can be evaluated against ground measurements. According to the spatial representativeness assessment (see Supplementary Note, and Figs. S7 and S8), the landscapes around the flux towers generally have heterogeneity at a level similar to, or smaller than, the tower footprint size (200–300 m) during the early GS and peak GS in the MODIS 1 × 1 pixel window (i.e., 500 × 500 m2), but mostly higher than in the 3 × 3 pixel window (Table S6). This indicates that it is desirable to evaluate the MODIS 1 × 1 pixel LAI values against ground measurements, as both MODIS greenup and snowmelt timings agreed more with the ground data at the 1 × 1 pixel window scale than at the 3 × 3 pixel window scale (Figs. S9 and S10). A more thorough evaluation of both MODIS LAI data and ED2 LAI values is required in the coming years with the increase in ground data availability (e.g., National Ecological Observatory Network, NEON, LAI measurements).Correlation analysis: the effects of early or delayed snowmelt timingTo analyze the net and lagged effects of early or late snowmelt timing, it is necessary to constrain the contribution of interannual meteorological variation. Therefore, we compared only the years when meteorological conditions were similar, i.e., when the weekly mean GSI value was within one standard deviation of the weekly mean GSI during 2001–2018 (at the US-Hva site, weekly values during 1994–2018); meteorological conditions appeared similar for 8 or 9 years at each study site, except the US-BZF site, where the similarities were found for 10 years. We also limited the effect of greenup timing changes by excluding the years when greenup was earlier or later by one standard deviation of the mean of greenup timings during the study period. For the years satisfying the constraints, a least-squares linear regression was applied between the snowmelt timing change (deviation in snowmelt timing each year from the mean snowmelt timing) and the seasonal deviation (the difference of the seasonal mean from the mean value over the years) of each process from the ED2 results.To analyze the net and lagged effects of delayed snowmelt, we implemented the ED2 model in two schemes, (1) following the meteorologically-determined phenological index (i.e., the GSI, Eqs. (2) and (2) constraining leaf-out by snowmelt (i.e., the SGSI, Eq. (3)). For the years when unmelted snow delayed greenup, we took the difference between the modeled results (i.e., GSI and SGSI) for each process and applied a least-squares linear regression between the difference of each process and the delayed snowmelt days. More

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    Parasitoid vectors a plant pathogen, potentially diminishing the benefits it confers as a biological control agent

    Insect rearingA CLas negative colony of ACP was initially collected from CLas-free Murraya exotica L. growing in the ornamental landscape of South China Agricultural University (SCAU, Guangzhou, China) in May 2014. Then it was reared on potted M. exotica in a greenhouse at SCAU. M. exotica plants were pruned regularly to promote the growth flushes necessary to stimulate ACP oviposition. The ACP populations were periodically (at least once a month) tested to ensure the colony was CLas-free using nested quantitative PCR detection according to the method described by Coy et al.30.The parasitoid T. radiata used in the current study was initially collected from ACP hosts on M. exotica plants in the above-mentioned location during June 2015. Its population was maintained in rearing cages (60 × 60 × 60 cm) using a CLas-free ACP-M. exotica rearing system under laboratory conditions (26 ± 1 °C, RH 80 ± 10% with L:D = 14:10 photoperiods in insect incubators).Host plantsCLas-free and CLas-infected plants of Citrus reticulata Blanco cv. Shatangju were used in the current study. Both plant types were obtained from The Citrus Research Institute of Zhaoqing University (Guangdong, China). The CLas-infected plants were inoculated by shoot grafting. All plants were approximately 4-year old and 1.2−1.5 m in height, separated in nylon net greenhouses (70 mesh per inch2) at two different locations about 2.2 km apart in SCAU. Again, nested qPCR detection was performed periodically (at least once a month) to test for the presence or absence of CLas in the citrus plants according to the method described by Coy et al.30.Acquisition and persistence of CLas in Tamarixia radiata
    When new shoots of CLas-infected C. reticulata plants were grown to 5–8 cm, 20 pairs of 1 week-old ACP adults were introduced into one nylon bag covering one fresh shoot to lay eggs for 48 h. When the progeny of ACP developed through to 4th or 5th instar nymph (CLas-donor ACP), which are the stages preferred by T. radiata parasitoids, 150 of the ACP nymphs were randomly selected and the remaining ones were removed. Following this, 10 pairs of 3-day old T. radiata adults, randomly selected from the population that has been tested to be CLas-free, were introduced into the nylon bag in order to parasitize the 4th or 5th instar ACP nymphs for 48 h before being recaptured. Then the potentially parasitized ACP nymphs together with the citrus plants were cultured in a plant growth chamber (Jiangnan Instrument Company, RXZ-500D, at 26 ± 1 °C, 60 ± 2% RH and 14:10 h L:D photoperiod of 3,000 lx illumination).When the progeny of T. radiata (considered F0 generation) developed to 3-day egg, 1st to 4th instar larvae, pupae, and adult stages respectively, they were identified and collected with the assistance of a stereomicroscope. DNA of each stage sample was extracted using the TIANamp Genomic DNA Kit (TIANGEN, Beijing, China) for CLas qPCR detection and titer quantification. Thirty eggs, 20 individuals of 1st or 2nd instar, 10 individuals of 3rd or 4th instar larvae or pupa, as well as three individuals of female or male adults were subsequently ground together to represent each life stage in qPCR, and each stage qPCR detection was repeated three times.The primers used for CLas qPCR detection were LJ900 primers, (F5′-GCCGTTTTAAC ACAAAAGATGAATATC-3′, R5′-ATAAATCAATTTGTTCTAGTTTAC GAC-3′), and 18S rRNA gene of T. radiata (F5′-AAACGGCTACCACATCCA-3′, R5′-ACCAGACT TGCCCTC CA-3′)31 was used as an internal control for DNA normalization and quantification. In order to normalize the qPCR values, each qPCR reaction was performed in three independent runs using SYBR Premix Ex Taq (Takara, Dalian, China) in Bio-Rad CFX Connect™ Real-Time PCR Detection System, with a protocol of initial denaturation at 95 °C for 3 min, followed by 40 cycles at 95 °C for 10 s, 60 °C for 20 s and 72 °C for 30 s.To monitor the CLas persistence in T. radiata, newly emerged female adults of T. radiata (considered F1 generation) were collected from the above experiment and fed with 20% honey water. After 1, 5, 10, and 15 days, 10 parasitoids were recaptured, subsequently ground for DNA extraction and CLas titer detection and quantification using qPCR. The protocol of DNA extraction and qPCR reaction was the same as above, and qPCR quantification was repeated three times for each treatment.Localization patterns of CLas in Tamarixia radiata
    Localization patterns of CLas in different instars of T. radiataFluorescent in situ hybridization (FISH) was used to visualize the distribution of CLas in T. radiata exposed to CLas positive ACP, following the method of Gottlieb et al.32 with a slight modification. Eggs and different larval instars of T. radiata were collected and fixed in Carnoy’s solution (chloroform-ethanol-glacial acetic acid [6:3:1,vol/vol] formamide) overnight at 4 °C. After fixation, the samples were washed three times in 50% ethanol with 1× phosphate buffered saline (PBS) for 5 min. Then the samples were decolorized in 6% H2O2 in ethanol for 12 h, after which they were hybridized overnight in 1 ml hybridization buffer (20 mM Tris-HCl pH 8.0, 0.9 M NaCl, 0.01% sodium dodecyl sulfate, 30% formamide) containing 10 pmol of fluorescent probes/ml in a 37 °C water bath under dark conditions. The CLas probe used for FISH was 5′-Cy3-GCCTCGCGACTTCGCAACCCAT-3′. Finally, the stained T. radiata samples were washed three times in a washing buffer (0.3 M NaCl, 0.03 M sodium citrate, 0.01% sodium dodecyl sulfate, 10 min per time). After the samples were whole mounted and stained, the slides were observed and photographed using a Nikon eclipse Ti-U inverted microscope. For each stage sample, approximately 20 individuals were examined to confirm the results.Localization patterns of CLas in different organs of T. radiataDifferent organs (gut, fat body, ovary, poison sac, salivary glands, spermatheca, and chest muscle) were dissected from newly emerged adults of T. radiata in 1× phosphate buffered saline (PBS) under a stereomicroscope using a depression microscope slide and a fine anatomical needle. After a sufficient number of each tissue sample was collected (20 or more), the tissues were washed three times with 1 × PBS, followed by the fixation, decolorization, and hybridization procedures as outlined above, except that this time of decolorization was 2 h. After hybridization, nuclei in the different organs were counterstained with DAPI (0.1 mg/ml in 1 × PBS) for 10 min, then the samples were transferred to slides, mounted whole in hybridization buffer, and viewed using confocal microscopy (Nikon, Japan).Maternal transmission of CLas between Tamarixia generationsFive groups of experiments were used to clarify whether CLas can be transmitted vertically between different T. radiata generations. In the first group, 60 pairs of newly emerged T. radiata adults from the CLas-infected ACP colony (potential CLas-acquired parasitoid adults, F0 generation) were introduced into 60 nylon bags (one female per cage). Each bag covered one fresh citrus plant shoot with one marked CLas-free 4th instar nymph of ACP, the parasitoid females were given 24 h to oviposit, then transferred to another four groups successively to oviposit with intervals of 24 h before they were recaptured for CLas-PCR detection (58/60 and 56/60 T. radiata females and males respectively were CLas-infected). Only the progeny (F1 generation) in which parasitoid parents were both CLas-infected continued to be investigated.When the F1 progeny of CLas-infected parasitoid females developed to egg, larval, pupal, and adult stages respectively, they were collected and divided into two groups; in one group samples were used for the qPCR detection of the CLas titer, and the other group was used for the FISH visualization of CLas. The qPCR and FISH analysis protocols of CLas as well as the number of tested individuals were the same as previously outlined. Each stage was repeated three times.
    CLas detection in T. radiata-inoculated ACPQuantitative PCR detection of CLasApproximately 60 newly emerged parasitoid adult females from CLas-infected ACP hosts (potential CLas-acquired parasitoid adults) were collected using an aspirator. They were first starved for 5 h, then released into finger tubes (diameter 6 mm × length 30 mm); one female per tube containing one 4th instar nymph of CLas-free ACP (this was treated as one experimental replicate). The probing behavior of the parasitoids was observed under a stereomicroscope, after which the parasitoids were recaptured for CLas PCR detection (similar to the above experiment, approximately 95% were CLas-infected). Only those 4th instar ACP nymphs, probed for egg-laying by a CLas-infected parasitoid but survived from the probing (the averaged proportion of such samples was 5.36 ± 0.47% and were 100% CLas infected), were transferred onto fresh CLas-free M. exotica shoots to complete their development (hereafter referred as “T. radiata-inoculated ACP”). The experiment was repeated in 32 parallel replicates (Supplementary Table 1), in which 103 T. radiata-inoculated ACP nymphs were finally obtained.Following the above, thirty T. radiata-inoculated ACP nymphs were collected when they developed into 5th instar nymphs (the stage when infection proliferation might have just begun since the infection was introduced at the 4th instar). In addition, thirty 8-day old adults that developed from the T. radiata-inoculated ACP nymphs were also collected. This was because the results in Wu et al.28 revealed that the proportion of CLas-infected ACP individuals exceeds 90% at the 12th day after infection acquisition, while ACP takes 4 days to develop into an adult from 5th instar stage. Their alimentary canals and salivary glands were dissected under a stereomicroscope using the methods of Ammar et al.33, and hemolymphs were collected with a 10 μl pipette tip using the method of Killiny et al.34. The DNA of the alimentary canals, salivary glands and hemolymphs were extracted using TIANamp Micro DNA Kit (Tiangen, Beijing, China), and the relative titers of CLas in each tissue of ACP nymphs and adults were detected by qPCR with of LJ900. The β-actin gene of ACP (F 5′-CCCTGGACTTTGAACAGGAA-3′; R 5′-CTCGTGGATACCGCAAGATT-3′) was selected as an internal control for data normalization and quantification35. For each sample, qPCR detection was repeated three times.FISH visualization of CLasThe alimentary canals and salivary glands of 5th instar nymphs and 8-day old adults of T. radiata-inoculated ACP were dissected as described above, and the distribution of CLas was visualized by FISH and confocal microscopy. The alimentary canals and salivary glands of CLas-infected ACP nymphs and adults (collected from CLas-infected citrus plants) were used as a positive control, and five to ten samples were detected by FISH for each tissue.
    CLas transmission from T. radiata-inoculated ACP to citrus plantsAccording to the above experimental results, if the CLas could be detected in the salivary glands of the 8-day old ACP adults (T. radiata-inoculated ACP), 30 more of these adults were randomly selected to inoculate on fresh shoots of CLas-free citrus. ACP adults that acquired CLas from plants and CLas-free ACP adults were used as positive and negative controls respectively.After 20, 30, 40, and 50 days of feeding samples of the citrus leaves fed on by T. radiata-inoculated ACP (named as CLas-recipient citrus leaves), fed on by ACP that acquired CLas from plants (positive control), and fed on by CLas-free ACP (negative control) were cut (1 cm2). Their DNAs were extracted using DNAsecure Plant Kit (Tiangen, Beijing, China). The infections of CLas in these plants were detected by nested PCR based on the methods of Jagoueix et al.36 and Deng et al.37. The experiment was repeated in six plants for each of 20, 30, 40, and 50 days feeding duration, and the infection rates of CLas were calculated.Localization of CLas in citrus plants fed on by T. radiata-inoculated ACPIn order to further confirm the infection of CLas in the recipient citrus leaves, FISH was used to visualize the localization of CLas. According to the above experimental results, after being fed on for 50 days by the T. radiata-inoculated ACP adults, citrus leaf sections containing the midrib were cross-sliced in 30 µ sections using a cryostat (CM1950, Leica, Germany). The leaf samples were prepared for FISH vitalization according to the protocol described by Gottlieb et al.32. Citrus leaves from the plant that had been fed on by ACP adults that acquired CLas from plants and CLas-free ACP adults were used as positive and negative controls, respectively. Five to 10 leaf samples were detected by FISH for each treatment.Phylogenetic analysis of CLas bacteria in different ACP populations and citrus plantsTo assess the identity of the CLas bacteria in CLas donor ACP, CLas vectored parasitoids, T. radiata-inoculated ACP and recipient citrus leaves, the outer membrane protein gene (omp) of CLas was PCR amplified with the primers HP1asinv (5′-GATGATAGG TGCATAAAAGTACAGAAG-3′) and Lp1c (5′-AATACCCTTATGGGATACAAAAA-3′) following the procedure described in Bastianel et al.38. Then the PCR products were sent for sequencing after visualizing the expected bands on 1% agarose gels.All the DNA sequences of CLas omp gene were edited and aligned manually using Clustal X1.8339 in Mega 640. The best model and partitioning scheme were chosen using the Bayesian information criterion in PartitionFinder v.1.0.141. Phylogenetic analysis was undertaken using a maximum likelihood (ML) method with 1000 non-parametric bootstrap replications in RAxML42. Escherichia coli was used as an outgroup.Statistics and reproducibilityTaking 18S rRNA gene of T. radiata and the β-actin gene of ACP as housekeeping genes, the relative titers of CLas in different stages and different tissues of T. radiata and ACP were calculated using the method of 2[−ΔΔct 43. For the parallel experiments that had more than three replicates the differences were compared using analysis of variance (ANOVA) with SPSS 18.0 at a significance level α = 0.05; while for CLas titer, two-sample comparison between genders of Tamarixia adults analysis was performed using paired t-test. Fluorescent pictures were processed using Photoshop CS5 software.Reporting summaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

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    Skin irritation and potential antioxidant, anti-collagenase, and anti-elastase activities of edible insect extracts

    Insect extractsThai edible insects (Fig. 1) were extracted and yield of each extract is shown in Fig. 2. Hexane extracts of most insects, except for P. succincta, provided the highest yield, followed by ethanolic extracts, and aqueous extracts, respectively. The reason might be due to a high amount of fat content of insects. Since these fat components are hydrophobic, they could be extracted well using nonpolar solvent, e.g. hexane. Semi-polar solvent like ethanol could also be used to extract hydrophobic compounds but with less extraction efficacy5. Several previous studies reported that fat was abundant in biomass of insects, ranging from 4.2 to 77.2%, which was accounted for about 26.8% on average dried insects6,7.Figure 1External appearances of Thai edible insects, including (a) rice grasshopper (Euconocephalus sp.), (b) bamboo caterpillar (O. fuscidentalis), (c) house cricket (A. domesticus), (d) silkworm pupae (B. mori), (e) Bombay locust (P. succincta), and (f) giant water bug (L. indicus).Full size imageFigure 2Yields of insect extracts, including B. mori (BM), O. fuscidentalis (OF), Euconocephalus sp. (EU), P. succincta (PS), A. domesticus (AD), and L. indicus (LI). The data are expressed as mean ± SD (n = 3). The Greek alphabet letters (α, β, γ, and δ) indicate significant differences among hexane extracts, the capital letters (A, B, C, and D) indicate significant differences among ethanolic extracts, and the small case letters (a, b, and c) indicate significant differences among aqueous extracts. The data were analyzed using One-Way ANOVA followed by post hoc Tukey test (p  More

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    Wolbachia reduces virus infection in a natural population of Drosophila

    1.Weinert, L. A., Araujo-Jnr, E. V., Ahmed, M. Z. & Welch, J. J. The incidence of bacterial endosymbionts in terrestrial arthropods. Proc. R. Soc. B: Biol. Sci. 282, 20150249 (2015).
    Google Scholar 
    2.Werren, J. H. Biology of Wolbachia. Annu Rev. Entomol. 42, 587–609 (1997).CAS 
    PubMed 

    Google Scholar 
    3.Turelli, M. & Hoffmann, A. A. Rapid spread of an inherited incompatibility factor in California Drosophila. Nature 353, 440–442 (1991).CAS 
    PubMed 

    Google Scholar 
    4.Werren, J. H., Baldo, L. & Clark, M. E. Wolbachia: master manipulators of invertebrate biology. Nat. Rev. Microbiol. 6, 741–751 (2008).CAS 
    PubMed 

    Google Scholar 
    5.Teixeira, L., Ferreira, A. & Ashburner, M. The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster. Plos Biol. 6, e2 (2008).PubMed 

    Google Scholar 
    6.Hedges, L. M., Brownlie, J. C., O’Neill, S. L. & Johnson, K. N. Wolbachia and virus protection in insects. Science 322, 702 (2008).CAS 
    PubMed 

    Google Scholar 
    7.Rocha, M. N. et al. Pluripotency of Wolbachia against Arboviruses: the case of yellow fever. Gates Open Res. 3, 161 (2019).PubMed 
    PubMed Central 

    Google Scholar 
    8.Moreira, L. A. et al. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell 139, 1268–1278 (2009).PubMed 

    Google Scholar 
    9.Dutra, H. L. et al. Wolbachia blocks currently circulating zika virus isolates in Brazilian Aedes aegypti mosquitoes. Cell Host Microbe 19, 771–774 (2016).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    10.Aliota, M. T. et al. The wMel strain of Wolbachia reduces transmission of chikungunya virus in Aedes aegypti. PLoS Negl. Trop. Dis. 10, e0004677 (2016).PubMed 
    PubMed Central 

    Google Scholar 
    11.Schmidt, T. L. et al. Local introduction and heterogeneous spatial spread of dengue-suppressing Wolbachia through an urban population of Aedes aegypti. PLoS Biol. 15, e2001894 (2017).PubMed 
    PubMed Central 

    Google Scholar 
    12.Ryan, P. A. et al. Establishment of wMel Wolbachia in Aedes aegypti mosquitoes and reduction of local dengue transmission in Cairns and surrounding locations in northern Queensland, Australia. Gates Open Res. 3, 1547 (2020).PubMed 
    PubMed Central 

    Google Scholar 
    13.Indriani, C. et al. Reduced dengue incidence following deployments of Wolbachia-infected Aedes aegypti in Yogyakarta, Indonesia: a quasi-experimental trial using controlled interrupted time series analysis. Gates Open Res. 4, 50 (2020).PubMed 
    PubMed Central 

    Google Scholar 
    14.Zug, R. & Hammerstein, P. Bad guys turned nice? A critical assessment of Wolbachia mutualisms in arthropod hosts. Biol. Rev. Camb. Philos. Soc. 90, 89–111 (2015).PubMed 

    Google Scholar 
    15.Shi, M. et al. No detectable effect of Wolbachia wMel on the prevalence and abundance of the RNA virome of Drosophila melanogaster. Proc. Biol. Sci. https://doi.org/10.1098/rspb.2018.1165 (2018).16.Webster, C. L. et al. The discovery, distribution, and evolution of viruses associated with Drosophila melanogaster. PLoS Biol. 13, e1002210 (2015).PubMed 
    PubMed Central 

    Google Scholar 
    17.Pimentel, A. C., Cesar, C. S., Martins, M. & Cogni, R. The antiviral effects of the symbiont bacteria Wolbachia in insects. Front Immunol. 11, 626329 (2021).PubMed 
    PubMed Central 

    Google Scholar 
    18.Kriesner, P., Hoffmann, A. A., Lee, S. F., Turelli, M. & Weeks, A. R. Rapid sequential spread of two Wolbachia variants in Drosophila simulans. PLoS Pathog. 9, e1003607 (2013).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    19.Weeks, A. R., Turelli, M., Harcombe, W. R., Reynolds, K. T. & Hoffmann, A. A. From parasite to mutualist: rapid evolution of Wolbachia in natural populations of Drosophila. PLoS Biol. 5, e114 (2007).PubMed 
    PubMed Central 

    Google Scholar 
    20.Hoffmann, A. A. & Turelli, M. Unidirectional incompatibility in Drosophila simulans: inheritance, geographic variation and fitness effects. Genetics 119, 435–444 (1988).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    21.Cross, S. T. et al. Partitiviruses infecting Drosophila melanogaster and Aedes aegypti exhibit efficient biparental vertical transmission. J. Virol. https://doi.org/10.1128/jvi.01070-20 (2020).22.Webster, C. L., Longdon, B., Lewis, S. H. & Obbard, D. J. Twenty-five new viruses associated with the Drosophilidae (Diptera). Evolut. Bioinforma. online 12, 13–25 (2016).CAS 

    Google Scholar 
    23.Jousset, F. X. & Plus, N. Study of the vertical transmission and horizontal transmission of “Drosophila melanogaster” and “Drosophila immigrans” picornavirus (author’s transl). Ann. Microbiol. 126, 231–249 (1975).CAS 

    Google Scholar 
    24.Jousset, F. X., Plus, N., Croizier, G. & Thomas, M. Existence in Drosophila of 2 groups of picornavirus with different biological and serological properties. C. R. Acad. Hebd. Seances Acad. Sci. D. 275, 3043–3046 (1972).CAS 
    PubMed 

    Google Scholar 
    25.Kapun, M. et al. Genomic Analysis of European Drosophila melanogaster populations reveals longitudinal structure, continent-wide selection, and previously unknown DNA viruses. Mol. Biol. Evol. 37, 2661–2678 (2020).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    26.Medd, N. C. et al. The virome of Drosophila suzukii, an invasive pest of soft fruit. Virus Evol. 4, vey009 (2018).PubMed 
    PubMed Central 

    Google Scholar 
    27.Longdon, B. et al. The evolution, diversity, and host associations of rhabdoviruses. Virus Evol. 1, vev014 (2015).PubMed 
    PubMed Central 

    Google Scholar 
    28.Schoonvaere, K., Smagghe, G., Francis, F. & de Graaf, D. C. Study of the metatranscriptome of eight social and solitary wild bee species reveals novel viruses and bee parasites. Front. Microbiol. 9, 177 (2018).PubMed 
    PubMed Central 

    Google Scholar 
    29.Pettersson, J. H., Shi, M., Eden, J. S., Holmes, E. C. & Hesson, J. C. Meta-transcriptomic comparison of the RNA viromes of the mosquito vectors Culex pipiens and Culex torrentium in Northern Europe. Viruses https://doi.org/10.3390/v11111033 (2019).30.Mahar, J. E., Shi, M., Hall, R. N., Strive, T. & Holmes, E. C. Comparative analysis of RNA virome composition in rabbits and associated ectoparasites. J. Virol. https://doi.org/10.1128/jvi.02119-19 (2020).31.Martinez, J. et al. Symbionts commonly provide broad spectrum resistance to viruses in insects: a comparative analysis of Wolbachia strains. PLoS Pathogens https://doi.org/10.1371/journal.ppat.1004369 (2014).32.Cross, S. T. et al. Galbut virus infection minimally influences Drosophila melanogaster fitness traits in a strain and sex-dependent manner. Preprint at bioRxiv https://doi.org/10.1101/2021.05.18.444759 (2021).33.Yampolsky, L. Y., Webb, C. T., Shabalina, S. A. & Kondrashov, A. S. Rapid accumulation of a vertically transmitted parasite triggered by relaxation of natural selection among hosts. Evolut. Ecol. Res. 1, 581–589 (1999).
    Google Scholar 
    34.Wilfert, L. & Jiggins, F. M. The dynamics of reciprocal selective sweeps of host resistance and a parasite counter-adaptation in Drosophila. Evolution 67, 761–773 (2013).CAS 
    PubMed 

    Google Scholar 
    35.Chrostek, E., Martins, N., Marialva, M. S. & Teixeira, L. Wolbachia conferred antiviral protection is determined by developmental temperature. mBio 12, e0292320 (2021).PubMed 

    Google Scholar 
    36.Ortiz-Baez, A. S., Shi, M., Hoffmann, A. A. & Holmes, E. C. RNA virome diversity and Wolbachia infection in individual Drosophila simulans flies. J. Gen. Virol. 102, 001639 (2021).
    Google Scholar 
    37.Haine, E. R. Symbiont-mediated protection. Proc. Biol. Sci. 275, 353–361 (2008).PubMed 

    Google Scholar 
    38.Martinez, J. et al. Addicted? Reduced host resistance in populations with defensive symbionts. Proc. Biol. Sci. https://doi.org/10.1098/rspb.2016.0778 (2016).39.Cogni, R. et al. Variation in Drosophila melanogaster central metabolic genes appears driven by natural selection both within and between populations. P R. Soc. B-Biol. Sci. 282, 20142688 (2015).CAS 

    Google Scholar 
    40.Cogni, R. et al. On the long-term stability of clines in some metabolic genes in Drosophila melanogaster. Sci. Rep. https://doi.org/10.1038/srep42766 (2017).41.Longdon, B. et al. The causes and consequences of changes in virulence following pathogen host shifts. PLoS Pathogens https://doi.org/10.1371/journal.ppat.1004728 (2015).42.Longdon, B., Hadfield, J. D., Webster, C. L., Obbard, D. J. & Jiggins, F. M. Host phylogeny determines viral persistence and replication in novel hosts. PLoS Pathog. 7, e1002260 (2011).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    43.Meyer, M. & Kircher, M. Illumina sequencing library preparation for highly multiplexed target capture and sequencing. Cold Spring Harb. Protoc. 2010, pdb.prot5448 (2010).PubMed 

    Google Scholar 
    44.Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    45.Quast, C. et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 41, D590–D596 (2013).CAS 
    PubMed 

    Google Scholar 
    46.Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    47.Grabherr, M. G. et al. Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat. Biotechnol. 29, 644–652 (2011).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    48.Buchfink, B., Xie, C. & Huson, D. H. Fast and sensitive protein alignment using DIAMOND. Nat. Methods 12, 59–60 (2015).CAS 

    Google Scholar 
    49.Notredame, C., Higgins, D. G. & Heringa, J. T-Coffee: a novel method for fast and accurate multiple sequence alignment. J. Mol. Biol. 302, 205–217 (2000).CAS 
    PubMed 

    Google Scholar 
    50.Guindon, S. et al. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst. Biol. 59, 307–321 (2010).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    51.Coyle, M. C., Elya, C. N., Bronski, M. & Eisen, M. B. Entomophthovirus: an insect-derived iflavirus that infects a behavior manipulating fungal pathogen of dipterans. Preprint at bioRxiv https://doi.org/10.1101/371526 (2018).52.Longdon, B. et al. Vertically transmitted rhabdoviruses are found across three insect families and have dynamic interactions with their hosts. P Roy Soc B-Biol Sci https://doi.org/10.1098/rspb.2016.2381 (2017).53.Untergasser, A. et al. Primer3-new capabilities and interfaces. Nucleic Acids Res. 40, e115 (2012).CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    54.Ye, J. et al. Primer-BLAST: A tool to design target-specific primers for polymerase chain reaction. BMC Bioinforma. 13, 134 (2012).CAS 

    Google Scholar 
    55.Lefever, S., Pattyn, F., Hellemans, J. & Vandesompele, J. Single-nucleotide polymorphisms and other mismatches reduce performance of quantitative PCR assays. Clin. Chem. 59, 1470–1480 (2013).CAS 
    PubMed 

    Google Scholar 
    56.Hadfield, J. D. MCMC Methods for Multi-Response Generalized Linear Mixed Models: The MCMCglmm R Package. 2010 33, 22, (2010).57.Cogni, R., Ding, S. D., Pimentel, A. C., Day, J. P. & Jiggins, F. M. https://doi.org/10.5281/zenodo.5525967 (Zenodo 2021). More

  • in

    Increased microbial expression of organic nitrogen cycling genes in long-term warmed grassland soils

    1.Schmidt MWI, Torn MS, Abiven S, Dittmar T, Guggenberger G, Janssens IA, et al. Persistence of soil organic matter as an ecosystem property. Nature. 2011;478:49–56.CAS 
    PubMed 

    Google Scholar 
    2.Bond-Lamberty B, Bailey VL, Chen M, Gough CM, Vargas R. Globally rising soil heterotrophic respiration over recent decades. Nature. 2018;560:80–3.CAS 
    PubMed 

    Google Scholar 
    3.Bradford MA. Thermal adaptation of decomposer communities in warming soils. Front Microbiol. 2013;4:1–16.
    Google Scholar 
    4.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 

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

    Google Scholar 
    6.Liu L, Greaver TL. A global perspective on belowground carbon dynamics under nitrogen enrichment. Ecol Lett. 2010;13:819–28.PubMed 

    Google Scholar 
    7.Knicker H. Soil organic N – An under-rated player for C sequestration in soils? Soil Biol Biochem. 2011;43:1118–29.CAS 

    Google Scholar 
    8.Soong JL, Fuchslueger L, Marañon-Jimenez S, Torn MS, Janssens IA, Peñuelas J, et al. Microbial carbon limitation: The need for integrating microorganisms into our understanding of ecosystem carbon cycling. Glob Chang Biol. 2020;26:1953–61.
    Google Scholar 
    9.Mooshammer M, Wanek W, Hämmerle I, Fuchslueger L, Hofhansl F, Knoltsch A, et al. Adjustment of microbial nitrogen use efficiency to carbon:nitrogen imbalances regulates soil nitrogen cycling. Nat Commun. 2014;5:1–7.
    Google Scholar 
    10.Geisseler D, Horwath WR, Joergensen RG, Ludwig B. Pathways of nitrogen utilization by soil microorganisms – a review. Soil Biol Biochem. 2010;42:2058–67.CAS 

    Google Scholar 
    11.Wang X, Wang C, Cotrufo MF, Sun L, Jiang P, Liu Z, et al. Elevated temperature increases the accumulation of microbial necromass nitrogen in soil via increasing microbial turnover. Glob Chang Biol. 2020;26:5277–89.PubMed 

    Google Scholar 
    12.Simpson AJ, Simpson MJ, Smith E, Kelleher BP. Microbially derived inputs to soil organic matter: Are current estimates too low? Environ Sci Technol. 2007;41:8070–6.CAS 
    PubMed 

    Google Scholar 
    13.Kuypers MMM, Marchant HK, Kartal B. The microbial nitrogen-cycling network. Nat Rev Microbiol. 2018;16:263–76.CAS 
    PubMed 

    Google Scholar 
    14.Walker TWN, Kaiser C, Strasser F, Herbold CW, Leblans NIW, Woebken D, et al. Microbial temperature sensitivity and biomass change explain soil carbon loss with warming. Nat Climate Change. 2018;8:885–9.CAS 

    Google Scholar 
    15.Marañón-Jiménez S, Peñuelas J, Richter A, Sigurdsson BD, Fuchslueger L, Leblans NIW, et al. Coupled carbon and nitrogen losses in response to seven years of chronic warming in subarctic soils. Soil Biol Biochem. 2019;134:152–61.
    Google Scholar 
    16.Nguyen TTH, Myrold DD, Mueller RS. Distributions of extracellular peptidases across prokaryotic genomes reflect phylogeny and habitat. Front Microbiol. 2019;10:1–14.
    Google Scholar 
    17.Zimmerman AE, Martiny AC, Allison SD. Microdiversity of extracellular enzyme genes among sequenced prokaryotic genomes. ISME J. 2013;7:1187–99.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    18.Beier S, Bertilsson S. Bacterial chitin degradation-mechanisms and ecophysiological strategies. Front Microbiol. 2013;4:1–12.
    Google Scholar 
    19.Kielak AM, Cretoiu MS, Semenov AV, Sørensen SJ, Van, Elsas JD. Bacterial chitinolytic communities respond to chitin and pH alteration in soil. Appl Environ Microbiol. 2013;79:263–72.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    20.Weintraub MN, Schimel JP. Seasonal protein dynamics in Alaskan arctic tundra soils. Soil Biol Biochem. 2005;37:1469–75.CAS 

    Google Scholar 
    21.Boer VM, De Winde JH, Pronk JT, Piper MDW. The genome-wide transcriptional responses of Saccharomyces cerevisiae grown on glucose in aerobic chemostat cultures limited for carbon, nitrogen, phosphorus, or sulfur. J Biol Chem. 2003;278:3265–74.CAS 
    PubMed 

    Google Scholar 
    22.Kolkman A, Daran-Lapujade P, Fullaondo A, Olsthoorn MMA, Pronk JT, Slijper M, et al. Proteome analysis of yeast response to various nutrient limitations. Mol Syst Biol. 2006;2:1–16.
    Google Scholar 
    23.Silberbach M, Hüser A, Kalinowski J, Pühler A, Walter B, Krämer R, et al. DNA microarray analysis of the nitrogen starvation response of Corynebacterium glutamicum. J Biotechnol. 2005;119:357–67.CAS 
    PubMed 

    Google Scholar 
    24.Merrick MJ, Edwards RA. Nitrogen control in bacteria. Microbiol Rev. 1995;59:604–22.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    25.Daebeler A, Abell GCJ, Bodelier PLE, Bodrossy L, Frampton DMF, Hefting MM, et al. Archaeal dominated ammonia-oxidizing communities in Icelandic grassland soils are moderately affected by long-term N fertilization and geothermal heating. Front Microbiol. 2012;3:1–14.
    Google Scholar 
    26.Yeager CM, Kornosky JL, Housman DC, Grote EE, Belnap J, Kuske CR. Diazotrophic community structure and function in two successional stages of biological soil crusts from the colorado plateau and Chihuahuan Desert. Appl Environ Microbiol. 2004;70:973–83.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    27.Malik AA, Swenson T, Weihe C, Morrison EW, Martiny JBH, Brodie EL, et al. Drought and plant litter chemistry alter microbial gene expression and metabolite production. ISME J. 2020;14:2236–47.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    28.Tveit A, Schwacke R, Svenning MM, Urich T. Organic carbon transformations in high-Arctic peat soils: Key functions and microorganisms. ISME J. 2013;7:299–311.CAS 
    PubMed 

    Google Scholar 
    29.Geisen S, Tveit AT, Clark IM, Richter A, Svenning MM, Bonkowski M, et al. Metatranscriptomic census of active protists in soils. ISME J. 2015;9:2178–90.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    30.Urich T, Lanzén A, Qi J, Huson DH, Schleper C, Schuster SC. Simultaneous assessment of soil microbial community structure and function through analysis of the meta-transcriptome. PLoS ONE. 2008;3:1–13.
    Google Scholar 
    31.Kallenbach CM, Frey SD, Grandy AS. Direct evidence for microbial-derived soil organic matter formation and its ecophysiological controls. Nat Commun. 2016;7:1–10.
    Google Scholar 
    32.Walker TWN, Janssens IA, Weedon JT, Sigurdsson BD, Richter A, Peñuelas J, et al. A systemic overreaction to years versus decades of warming in a subarctic grassland ecosystem. Nat Ecol Evol. 2020;4:101–8.PubMed 

    Google Scholar 
    33.Sigurdsson BD, Wallander H, Gunnarsdóttir GE, Richter A, Sigurðsson P, Leblans NIW, et al. Geothermal ecosystems as natural climate change experiments: the ForHot research site in Iceland as a case study. Icelandic Agric Sci. 2016;29:53–71.
    Google Scholar 
    34.Söllinger A, Séneca J, Dahl MB, Prommer J, Verbruggen E, Sigurdsson BD, et al. Downregulation of the microbial protein biosynthesis machinery in response to weeks, years and decades of soil warming. 2021 Research Square preprint. https://doi.org/10.21203/rs.3.rs-132190/v235.Leblans N. Natural gradients in temperature and nitrogen: Iceland represents a unique environment to clarify long-term global change effects on carbon dynamics. Joint doctoral dissertation. Antwerp University and Agricultural University of Iceland, Reykjavik, Iceland; 2016:1–229.36.Angel R, Claus P, Conrad R. Methanogenic archaea are globally ubiquitous in aerated soils and become active under wet anoxic conditions. ISME J. 2012;6:847–62.CAS 
    PubMed 

    Google Scholar 
    37.Hyatt D, Chen G-L, Locascio PF, Land ML, Larimer FW, Hauser LJ. Prodigal: prokaryotic gene recognition and translation initiation site identification. BMC Bioinformatics. 2010;11:5–11.
    Google Scholar 
    38.Gillespie CS. Fitting heavy tailed distributions: the poweRlaw Package. J Stat Softw. 2015;64:1–16.
    Google Scholar 
    39.El-Gebali S, Mistry J, Bateman A, Eddy SR, Luciani A, Potter SC, et al. The Pfam protein families database in 2019. Nucleic Acids Res. 2018;47:427–32.
    Google Scholar 
    40.Eddy SR. Accelerated profile HMM searches. PLOS Comput Biol. 2011;7:e1002195.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    41.Petersen TN, Brunak S, von Heijne G, Nielsen H. SignalP 4.0: discriminating signal peptides from transmembrane regions. Nat Methods. 2011;8:785–6.CAS 
    PubMed 

    Google Scholar 
    42.Bendtsen JD, Kiemer L, Fausbøll A, Brunak S. Non-classical protein secretion in bacteria. BMC Microbiol. 2005;5:1–13.
    Google Scholar 
    43.Yu NY, Wagner JR, Laird MR, Melli G, Rey S, Lo R, et al. PSORTb 3.0: improved protein subcellular localization prediction with refined localization subcategories and predictive capabilities for all prokaryotes. Bioinformatics. 2010;26:1608–15.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    44.Orsi WD. MetaProt: an integrated database of predicted proteins for improved annotation of metaomic datasets. Open Data LMU. 2020. https://doi.org/10.5282/ubm/data.18345.Lombard V, Golaconda Ramulu H, Drula E, Coutinho PM, Henrissat B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 2013;42:490–5.
    Google Scholar 
    46.Buchfink B, Xie C, Huson DH. Fast and sensitive protein alignment using DIAMOND. Nat Methods. 2014;12:59–60.PubMed 

    Google Scholar 
    47.Oksanen AJ, Blanchet FG, Kindt R, Legen- P, Minchin PR, Hara RBO, et al. vegan: Community Ecology Package. 2019. https://cran.r-project.org/package=vegan48.Lê S, Josse J, Husson F. FactoMineR: an R package for multivariate analysis. J Stat Softw. 2008;25:1–18.
    Google Scholar 
    49.Kolde R. pheatmap: pretty heatmaps. 2019. https://cran.r-project.org/package=pheatmap50.Noll L, Zhang S, Zheng Q, Hu Y, Wanek W. Wide-spread limitation of soil organic nitrogen transformations by substrate availability and not by extracellular enzyme content. Soil Biol Biochem. 2019;133:37–49.CAS 
    PubMed 
    PubMed Central 

    Google Scholar 
    51.Schimel JP, Bennett J. Nitrogen mineralization: challenges of a changing paradigm. Ecology. 2004;85:591–602.
    Google Scholar 
    52.Wild B, Ambus P, Reinsch S, Richter A. Resistance of soil protein depolymerization rates to eight years of elevated CO2, warming, and summer drought in a temperate heathland. Biogeochemistry. 2018;140:255–67.CAS 

    Google Scholar 
    53.Wanek W, Mooshammer M, Blöchl A, Hanreich A, Richter A. Determination of gross rates of amino acid production and immobilization in decomposing leaf litter by a novel 15N isotope pool dilution technique. Soil Biol Biochem. 2010;42:1293–302.CAS 

    Google Scholar 
    54.Liang C, Schimel JP, Jastrow JD. The importance of anabolism in microbial control over soil carbon storage. Nat Microbiol. 2017;2:1–6.
    Google Scholar 
    55.Vranova V, Rejsek K, Formanek P. Proteolytic activity in soil: a review. Appl Soil Ecol. 2013;70:23–32.
    Google Scholar 
    56.Schimel JP, Weintraub MN. The implications of exoenzyme activity on microbial carbon and nitrogen limitation in soil: a theoretical model. Soil Biol Biochem. 2003;35:549–63.CAS 

    Google Scholar 
    57.Rawlings ND, Waller M, Barrett AJ, Bateman A. MEROPS: The database of proteolytic enzymes, their substrates and inhibitors. Nucleic Acids Res. 2014;42:503–9.
    Google Scholar 
    58.Vollmer W, Joris B, Charlier P, Foster S. Bacterial peptidoglycan (murein) hydrolases. FEMS Microbiol Rev. 2008;32:259–86.CAS 
    PubMed 

    Google Scholar 
    59.Vermassen A, Leroy S, Talon R, Provot C, Popowska M, Desvaux M. Cell wall hydrolases in bacteria: Insight on the diversity of cell wall amidases, glycosidases and peptidases toward peptidoglycan. Front Microbiol. 2019;10:1–27.
    Google Scholar 
    60.Donhauser J, Qi W, Bergk-Pinto B, Frey B. High temperatures enhance the microbial genetic potential to recycle C and N from necromass in high-mountain soils. Glob Chang Biol. 2020;27:1365–86.61.Vollmer W, Blanot D, De Pedro MA. Peptidoglycan structure and architecture. FEMS Microbiology Reviews. 2008;32:149–67.CAS 
    PubMed 

    Google Scholar 
    62.Semchenko M, Leff JW, Lozano YM, Saar S, Davison J, Wilkinson A, et al. Fungal diversity regulates plant-soil feedbacks in temperate grassland. Science Adv. 2018;4.63.Saary P, Mitchell AL, Finn RD. Estimating the quality of eukaryotic genomes recovered from metagenomic analysis. Genome Biol. 2020;21:244.PubMed 
    PubMed Central 

    Google Scholar  More

  • in

    Climate and land-use changes reduce the benefits of terrestrial protected areas

    1.Watson, J. E. M., Dudley, N., Segan, D. B. & Hockings, M. The performance and potential of protected areas. Nature 515, 67–73 (2014).CAS 

    Google Scholar 
    2.Juffe-Bignoli, D. et al. Protected Planet Report 2014 (UNEP-WCMC, 2014).3.Gray, C. L. et al. Local biodiversity is higher inside than outside terrestrial protected areas worldwide. Nat. Commun. 7, 12306 (2016).4.Xu, W. et al. Strengthening protected areas for biodiversity and ecosystem services in China. Proc. Natl Acad. Sci. USA 114, 1601–1606 (2017).CAS 

    Google Scholar 
    5.Naidoo, R. et al. Evaluating the impacts of protected areas on human well-being across the developing world. Sci. Adv. 5, eaav3006 (2019).CAS 

    Google Scholar 
    6.Geldmann, J. et al. Effectiveness of terrestrial protected areas in reducing habitat loss and population declines. Biol. Conserv. 161, 230–238 (2013).
    Google Scholar 
    7.Cazalis, V. et al. Effectiveness of protected areas in conserving tropical forest birds. Nat. Commun. 11, 4461 (2020).8.Elsen, P. R., Monahan, W. B., Dougherty, E. R. & Merenlender, A. M. Keeping pace with climate change in global terrestrial protected areas. Sci. Adv. 6, eaay0814 (2020).
    Google Scholar 
    9.Hoffmann, S., Irl, S. D. H. & Beierkuhnlein, C. Predicted climate shifts within terrestrial protected areas worldwide. Nat. Commun. 10, 4787 (2019).10.Batllori, E., Parisien, M. A., Parks, S. A., Moritz, M. A. & Miller, C. Potential relocation of climatic environments suggests high rates of climate displacement within the North American protection network. Glob. Change Biol. 23, 3219–3230 (2017).
    Google Scholar 
    11.Ward, M. et al. Just ten percent of the global terrestrial protected area network is structurally connected via intact land. Nat. Commun. 11, 4563 (2020).CAS 

    Google Scholar 
    12.Jones, K. R. et al. One-third of global protected land is under intense human pressure. Science 360, 788–791 (2018).CAS 

    Google Scholar 
    13.Parks, S. A., Carroll, C., Dobrowski, S. Z. & Allred, B. W. Human land uses reduce climate connectivity across North America. Glob. Change Biol. 26, 2944–2955 (2020).
    Google Scholar 
    14.McGuire, J. L., Lawler, J. J., McRae, B. H., Nuñez, T. A. & Theobald, D. M. Achieving climate connectivity in a fragmented landscape. Proc. Natl Acad. Sci. USA 113, 7195–7200 (2016).CAS 

    Google Scholar 
    15.Watson, J. E. M., Iwamura, T. & Butt, N. Mapping vulnerability and conservation adaptation strategies under climate change. Nat. Clim. Change 3, 989–994 (2013).
    Google Scholar 
    16.Pecl, G. T. et al. Biodiversity redistribution under climate change: impacts on ecosystems and human well-being. Science 355, eaai9214 (2017).
    Google Scholar 
    17.Jones, C., Giorgi, F. & Asrar, G. The coordinated regional downscaling experiment: CORDEX–an international downscaling link to CMIP5. CLIVAR Exch. 16, 34–40 (2011).
    Google Scholar 
    18.Hurtt, G. C. et al. Harmonization of global land-use change and management for the period 850-2100 (LUH2) for CMIP6. Geosci. Model Dev. 13, 5425–5464 (2020).CAS 

    Google Scholar 
    19.Loarie, S. R. et al. The velocity of climate change. Nature 462, 1052–1055 (2009).CAS 

    Google Scholar 
    20.Ordonez, A., Martinuzzi, S., Radeloff, V. C. & Williams, J. W. Combined speeds of climate and land-use change of the conterminous US until 2050. Nat. Clim. Change 4, 811–816 (2014).
    Google Scholar 
    21.UN General Assembly Resolution A/RES/70/1 (UN, 2015).22.Harrop, S. R. ‘Living in harmony with nature’? Outcomes of the 2010 Nagoya conference of the convention on biological diversity. J. Environ. Law 23, 117–128 (2011).
    Google Scholar 
    23.Maxwell, S. L. et al. Area-based conservation in the twenty-first century. Nature 586, 217–227 (2020).CAS 

    Google Scholar 
    24.Schloss, C. A., Nuñez, T. A. & Lawler, J. J. Dispersal will limit ability of mammals to track climate change in the Western Hemisphere. Proc. Natl Acad. Sci. USA 109, 8606–8611 (2012).CAS 

    Google Scholar 
    25.Chen, I. C., Hill, J. K., Ohlemüller, R., Roy, D. B. & Thomas, C. D. Rapid range shifts of species associated with high levels of climate warming. Science 333, 1024–1026 (2011).CAS 

    Google Scholar 
    26.Schwalm, C. R., Glendon, S. & Duffy, P. B. RCP8.5 tracks cumulative CO2 emissions. Proc. Natl Acad. Sci. USA 117, 19656–19657 (2020).CAS 

    Google Scholar 
    27.Ando, A. W. & Mallory, M. L. Optimal portfolio design to reduce climate-related conservation uncertainty in the Prairie Pothole Region. Proc. Natl Acad. Sci. USA 109, 6484–6489 (2012).CAS 

    Google Scholar 
    28.Ackerly, D. D. et al. The geography of climate change: implications for conservation biogeography. Divers. Distrib. 16, 476–487 (2010).
    Google Scholar 
    29.Dobrowski, S. Z. & Parks, S. A. Climate change velocity underestimates climate change exposure in mountainous regions. Nat. Commun. 7, 12349 (2016).30.Hoegh-Guldberg, O. et al. in Special Report on Global Warming of 1.5°C (eds Masson-Delmotte, V. et al.) 175–311 (IPCC, WMO, 2018).31.Sandel, B. et al. The influence of late Quaternary climate-change velocity on species endemism. Science 334, 660–664 (2011).CAS 

    Google Scholar 
    32.Ordonez, A., Williams, J. W. & Svenning, J.-C. Mapping climatic mechanisms likely to favour the emergence of novel communities. Nat. Clim. Change 6, 1104–1109 (2016).
    Google Scholar 
    33.Carroll, C. et al. Scale-dependent complementarity of climatic velocity and environmental diversity for identifying priority areas for conservation under climate change. Glob. Change Biol. 23, 4508–4520 (2017).
    Google Scholar 
    34.Alexander, J. M. et al. Lags in the response of mountain plant communities to climate change. Glob. Change Biol. 24, 563–579 (2018).
    Google Scholar 
    35.Lawler, J. J. et al. Projected land-use change impacts on ecosystem services in the United States. Proc. Natl Acad. Sci. USA 111, 7492–7497 (2014).CAS 

    Google Scholar 
    36.Stein, B. A. et al. Preparing for and managing change: climate adaptation for biodiversity and ecosystems. Front. Ecol. Environ. 11, 502–510 (2013).
    Google Scholar 
    37.Elsen, P. R., Monahan, W. B. & Merenlender, A. M. Global patterns of protection of elevational gradients in mountain ranges. Proc. Natl Acad. Sci. USA 115, 6004–6009 (2018).CAS 

    Google Scholar 
    38.Burrows, M. T. et al. The pace of shifting climate in marine and terrestrial ecosystems. Science 334, 652–655 (2011).CAS 

    Google Scholar 
    39.Burrows, M. T. et al. Geographical limits to species-range shifts are suggested by climate velocity. Nature 507, 492–495 (2014).CAS 

    Google Scholar 
    40.Fitzpatrick, M. C., Gove, A. D., Sanders, N. & Dunn, R. R. Climate change, plant migration, and range collapse in a global biodiversity hotspot: the Banksia (Proteaceae) of Western Australia. Glob. Change Biol. 14, 1337–1352 (2008).
    Google Scholar 
    41.Dynesius, M. & Jansson, R. Evolutionary consequences of changes in species’ geographical distributions driven by Milankovitch climate oscillations. Proc. Natl Acad. Sci. USA 97, 9115–9120 (2000).CAS 

    Google Scholar 
    42.Geldmann, J., Manica, A., Burgess, N. D., Coad, L. & Balmford, A. A global-level assessment of the effectiveness of protected areas at resisting anthropogenic pressures. Proc. Natl Acad. Sci. USA 116, 23209–23215 (2019).CAS 

    Google Scholar 
    43.Tittensor, D. P. et al. Integrating climate adaptation and biodiversity conservation in the global ocean. Sci. Adv. 5, eaay9969 (2019).
    Google Scholar 
    44.Osorio, F., Vallejos, R. & Cuevas, F. SpatialPack: Package for Analysis of Spatial Data. R package version 0.2-3 (2014).45.Williams, K. D. et al. The Met Office Global Coupled model 2.0 (GC2) configuration. Geosci. Model Dev. 8, 1509–1524 (2015).
    Google Scholar 
    46.Giorgetta, M. A. et al. Climate and carbon cycle changes from 1850 to 2100 in MPI-ESM simulations for the Coupled Model Intercomparison Project Phase 5. J. Adv. Model. Earth Syst. https://doi.org/10.1002/jame.20038 (2013).47.Knudsen, E. M. & Walsh, J. E. Northern Hemisphere storminess in the Norwegian Earth System Model (NorESM1-M). Geosci. Model Dev. 9, 2335–2355 (2016).
    Google Scholar 
    48.Brito-Morales, I. et al. Climate velocity can inform conservation in a warming world. Trends Ecol. Evol. 33, 441–457 (2018).
    Google Scholar 
    49.García Molinos, J., Schoeman, D. S., Brown, C. J. & Burrows, M. T. VoCC: an R package for calculating the velocity of climate change and related climatic metrics. Methods Ecol. Evol. 10, 2195–2202 (2019).
    Google Scholar 
    50.UNEP‐WCMC & IUCN Protected Planet: The World Database on Protected Areas (WDPA, 2018).51.Visconti, P. et al. Protected area targets post-2020. Science 364, eaav6886 (2019).
    Google Scholar 
    52.Farr, T. G. et al. The shuttle radar topography mission. Rev. Geophys. https://doi.org/10.1029/2005RG000183 (2007).53.Olson, D. M. et al. Terrestrial ecoregions of the world: a new map of life on Earth. BioScience 51, 933–938 (2001).54.Ellis, E. C., Antill, E. C. & Kreft, H. All is not loss: plant biodiversity in the anthropocene. PLoS ONE 7, 30535 (2012).55.Asamoah, E. F. Climate Velocity and Land-use Instability 1971–2100 (Figshare, 2021); https://doi.org/10.6084/m9.figshare.14852955.v4 More

  • in

    Trees outside of forests as natural climate solutions

    1.Chao, S. Forest Peoples: Numbers Across the World (Forest Peoples Programme, 2021).2.Zomer, R. J. et al. Sci. Rep. 6, 29987 (2016).CAS 
    Article 

    Google Scholar 
    3.Schnell, S., Altrell, D., Ståhl, G. & Kleinn, C. Environ. Monit. Assess. 187, 4197 (2015).Article 

    Google Scholar 
    4.Brandt, M. et al. Nature 587, 78–82 (2020).Article 

    Google Scholar 
    5.Baccini, A. et al. Nat. Clim. Chang. 2, 182–185 (2012).CAS 
    Article 

    Google Scholar 
    6.Miller, D. C., Muñoz-Mora, J. C. & Christiaensen, L. Forest Policy Econ. 84, 47–61 (2017).Article 

    Google Scholar 
    7.Mbow, C., Smith, P., Skole, D., Duguma, L. & Bustamante, M. Curr. Opin. Environ. Sustain. 6, 8–14 (2014).Article 

    Google Scholar 
    8.Brandt, M. et al. Nat. Geosci. 11, 328–333 (2018).CAS 
    Article 

    Google Scholar 
    9.Mbow, C. et al. in Climate Change and Agriculture (ed. Deryng, D.) Ch. 10 (Burleigh Dodds Science Publishing, 2020).10.Akinyemi, F. O., Ghazaryan, G. & Dubovyk, O. Land Degrad. Dev. 32, 158–172 (2021).Article 

    Google Scholar 
    11.Sitch, S. et al. Biogeosciences 12, 653–679 (2015).Article 

    Google Scholar 
    12.Schnell, S., Kleinn, C. & Ståhl, G. Environ. Monit. Assess. 187, 600 (2015).Article 

    Google Scholar 
    13.Hansen, M. C. et al. Science 342, 850–853 (2013).CAS 
    Article 

    Google Scholar 
    14.Kuyah, S. et al. Agrofor. Syst. 86, 267–277 (2012).Article 

    Google Scholar 
    15.Nationally Determined Contributions Under the Paris Agreement Synthesis Report by the Secretariat FCCC/PA/CMA/2021/2/Add 2 (UNFCCC, 2021); https://unfccc.int/documents/26857316.Lohbeck, M. et al. Sci. Rep. 10, 15038 (2020).CAS 
    Article 

    Google Scholar 
    17.Chomba, S., Sinclair, F., Savadogo, P., Bourne, M. & Lohbeck, M. Front. For. Glob. Chang. 3, 571679 (2020).Article 

    Google Scholar 
    18.Griscom, B. W. et al. Proc. Natl Acad. Sci. USA 114, 11645–11650 (2017).CAS 
    Article 

    Google Scholar  More

  • in

    Reduced deforestation and degradation in Indigenous Lands pan-tropically

    1.Weisse, M. & Goldman, E. D. We Lost a Football Pitch of Primary Rainforest Every 6 Seconds in 2019 (World Resources Institute, 2020); https://www.wri.org/blog/2020/06/global-tree-cover-loss-data-20192.Gibson, L. et al. Primary forests are irreplaceable for sustaining tropical biodiversity. Nature 478, 378–381 (2011).CAS 

    Google Scholar 
    3.Pan, Y. et al. A large and persistent carbon sink in the world’s forests. Science 333, 988–993 (2011).CAS 

    Google Scholar 
    4.State of the World’s Indigenous Peoples: Rights to Lands, Territories and Resources (UN, 2021).5.Curtis, P. G., Slay, C. M., Harris, N. L., Tyukavina, A. & Hansen, M. C. Classifying drivers of global forest loss. Science 361, 1108–1111 (2018).CAS 

    Google Scholar 
    6.Larsen, P. B. et al. Understanding and responding to the environmental human rights defenders crisis: the case for conservation action. Conserv. Lett. 14, e12777 (2020).
    Google Scholar 
    7.Tauli-Corpuz, V., Alcorn, J., Molnar, A., Healy, C. & Barrow, E. Cornered by PAs: adopting rights-based approaches to enable cost-effective conservation and climate action. World Dev. 130, 104923 (2020).
    Google Scholar 
    8.Dinerstein, E. et al. A global deal for nature: guiding principles, milestones, and targets. Sci. Adv. 5, eaaw2869 (2019).CAS 

    Google Scholar 
    9.Dudley, N. et al. The essential role of other effective area-based conservation measures in achieving big bold conservation targets. Glob. Ecol. Conserv. 15, e00424 (2018).
    Google Scholar 
    10.Zero Draft of the Post-2020 Global Biodiversity Framework CBD/WG2020/2/3 (Convention on Biological Diversity, 2020).11.NGO Concerns Over the Proposed 30% Target for Protected Areas and Absence of Safeguards for Indigenous Peoples and Local Communities (Rainforest Foundation UK, 2021).12.Reyes-García, V. et al. Recognizing Indigenous Peoples’ and local communities’ rights and agency in the post-2020 Biodiversity Agenda. Ambio https://doi.org/10.1007/s13280-021-01561-7 (2021).13.Territories of Life: 2021 Report 52 (ICCA Consortium, 2021); https://report.territoriesoflife.org14.Garnett, S. T. et al. A spatial overview of the global importance of Indigenous lands for conservation. Nat. Sustain. 1, 369–374 (2018).
    Google Scholar 
    15.Fa, J. E. et al. Importance of Indigenous Peoples’ lands for the conservation of intact forest landscapes. Front. Ecol. Environ. 18, 135–140 (2020).
    Google Scholar 
    16.Vergara-Asenjo, G. & Potvin, C. Forest protection and tenure status: the key role of indigenous peoples and protected areas in Panama. Glob. Environ. Change 28, 205–215 (2014).
    Google Scholar 
    17.Blackman, A. & Veit, P. Titled Amazon indigenous communities cut forest carbon emissions. Ecol. Econ. 153, 56–67 (2018).
    Google Scholar 
    18.Walker, W. S. et al. The role of forest conversion, degradation, and disturbance in the carbon dynamics of Amazon indigenous territories and protected areas. Proc. Natl Acad. Sci. USA 117, 3015–3025 (2020).CAS 

    Google Scholar 
    19.Nolte, C., Agrawal, A., Silvius, K. M. & Soares-Filho, B. S. Governance regime and location influence avoided deforestation success of protected areas in the Brazilian Amazon. Proc. Natl Acad. Sci. USA 110, 4956–4961 (2013).CAS 

    Google Scholar 
    20.Schleicher, J., Peres, C. A., Amano, T., Llactayo, W. & Leader-Williams, N. Conservation performance of different conservation governance regimes in the Peruvian Amazon. Sci. Rep. 7, 11318 (2017).
    Google Scholar 
    21.Jusys, T. Changing patterns in deforestation avoidance by different protection types in the Brazilian Amazon. PLoS ONE 13, e0195900 (2018).
    Google Scholar 
    22.State of the World’s Indigenous Peoples (UN, 2009).23.Jackson, J. E. & Warren, K. B. Indigenous movements in Latin America, 1992–2004: controversies, ironies, new directions. Annu. Rev. Anthropol. 34, 549–573 (2005).
    Google Scholar 
    24.Vancutsem, C. et al. Long-term (1990–2019) monitoring of forest cover changes in the humid tropics. Sci. Adv. 7, eabe1603 (2021).
    Google Scholar 
    25.Hansen, M. C. et al. High-resolution global maps of 21st-century forest cover change. Science 342, 850–853 (2013).CAS 

    Google Scholar 
    26.Stuart, E. A. & Rubin, D. B. in Best Practices in Quantitative Methods (ed. Osborne, J.) 155–176 (SAGE Publications, 2008).27.Pfaff, A., Robalino, J., Lima, E., Sandoval, C. & Herrera, L. D. Governance, location and avoided deforestation from protected areas: greater restrictions can have lower impact, due to differences in location. World Dev. 55, 7–20 (2014).
    Google Scholar 
    28.Leberger, R., Rosa, I. M. D., Guerra, C. A., Wolf, F. & Pereira, H. M. Global patterns of forest loss across IUCN categories of protected areas. Biol. Conserv. 241, 108299 (2020).
    Google Scholar 
    29.Borrini-Feyerabend, G. et al. Governance of Protected Areas: From Understanding to Action (IUCN, 2013).30.Who Owns the World’s Land? A Global Baseline of Formally Recognized Indigenous and Community Land Rights (Rights and Resources Initiative, 2015); https://rightsandresources.org/wp-content/uploads/GlobalBaseline_web.pdf31.Dubertret, F. & Alden Wily, L. Percent of Indigenous and Community Lands (Landmark, 2015).32.Under the Cover of COVID: New Laws in Asia Favor Business at the Cost of Indigenous Peoples’ and Local Communities’ Land and Territorial Rights (Rights and Resources Initiative, 2020).33.Domínguez, L. & Luoma, C. Decolonising conservation policy: how colonial land and conservation ideologies persist and perpetuate indigenous injustices at the expense of the environment. Land 9, 65 (2020).
    Google Scholar 
    34.Pyhälä, A., Orozco, A. O. & Counsell, S. Protected Areas in the Congo Basin: Failing both people and biodiversity? (FAO, 2016).35.Pearson, T. R. H., Brown, S., Murray, L. & Sidman, G. Greenhouse gas emissions from tropical forest degradation: an underestimated source. Carbon Balance Manag. 12, 3 (2017).
    Google Scholar 
    36.Barlow, J. et al. Anthropogenic disturbance in tropical forests can double biodiversity loss from deforestation. Nature 535, 144–147 (2016).CAS 

    Google Scholar 
    37.Hansen, A. J. et al. A policy-driven framework for conserving the best of Earth’s remaining moist tropical forests. Nat. Ecol. Evol. 4, 1377–1384 (2020).
    Google Scholar 
    38.Milodowski, D. T. et al. The impact of logging on vertical canopy structure across a gradient of tropical forest degradation intensity in Borneo. J. Appl. Ecol. 58, 1764–1775 (2021).
    Google Scholar 
    39.Benítez-López, A., Santini, L., Schipper, A. M., Busana, M. & Huijbregts, M. A. J. Intact but empty forests? Patterns of hunting-induced mammal defaunation in the tropics. PLoS Biol. 17, e3000247 (2019).
    Google Scholar 
    40.Miettinen, J., Stibig, H.-J. & Achard, F. Remote sensing of forest degradation in Southeast Asia—aiming for a regional view through 5–30 m satellite data. Glob. Ecol. Conserv. 2, 24–36 (2014).
    Google Scholar 
    41.Yuliani, E. L. et al. Keeping the land: indigenous communities’ struggle over land use and sustainable forest management in Kalimantan, Indonesia. Ecol. Soc. 23, art49 (2018).
    Google Scholar 
    42.Berkes, F. Sacred Ecology (Routledge, 2017).43.Sheil, D., Boissière, M. & Beaudoin, G. Unseen sentinels: local monitoring and control in conservation’s blind spots. Ecol. Soc. 20, 39 (2015).
    Google Scholar 
    44.Sasaoka, M. & Laumonier, Y. Suitability of local resource management practices based on supernatural enforcement mechanisms in the local social-cultural context. Ecol. Soc. 17, 6 (2012).
    Google Scholar 
    45.Asante, E. A., Ababio, S. & Boadu, K. B. The use of indigenous cultural practices by the Ashantis for the conservation of forests in Ghana. SAGE Open 7, 215824401668761 (2017).
    Google Scholar 
    46.Schwartzman, S. et al. The natural and social history of the indigenous lands and protected areas corridor of the Xingu River basin. Philos. Trans. R. Soc. B 368, 20120164 (2013).
    Google Scholar 
    47.Hayes, T. M. & Murtinho, F. Are indigenous forest reserves sustainable? An analysis of present and future land-use trends in Bosawas, Nicaragua. Int. J. Sustain. Dev. World Ecol. 15, 497–511 (2008).
    Google Scholar 
    48.Tellman, B. et al. Illicit drivers of land use change: narcotrafficking and forest loss in central America. Glob. Environ. Change 63, 102092 (2020).
    Google Scholar 
    49.Bryan, J. For Nicaragua’s indigenous communities, land rights in name only: delineating boundaries among indigenous and black communities in eastern Nicaragua was supposed to guaranteed their land rights. Instead, it did the opposite. NACLA Rep. Am. 51, 55–64 (2019).
    Google Scholar 
    50.Seymour, F., La Vina, T. & Hite, K. Evidence Linking Community-level Tenure and Forest Condition: An Annotated Bibliography (Climate and Land Use Alliance, 2014).51.Tseng, T.-W. J. et al. Influence of land tenure interventions on human well-being and environmental outcomes. Nat. Sustain. 4, 242–251 (2021).
    Google Scholar 
    52.Robinson, B. E. et al. Incorporating land tenure security into conservation: conservation and land tenure security. Conserv. Lett. 11, e12383 (2018).
    Google Scholar 
    53.Smith, D. A., Holland, M. B., Michon, A., Ibáñez, A. & Herrera, F. The hidden layer of indigenous land tenure: informal forest ownership and its implications for forest use and conservation in Panama’s largest collective territory. Int. For. Rev. 19, 478–494 (2017).
    Google Scholar 
    54.Larson, A. M. & Springer, J. Recognition and Respect for Tenure Rights (IUCN, CEESP, CIFOR, 2016).55.Arizona, Y., Wicaksono, M. T. & Vel, J. The role of indigeneity NGOs in the legal recognition of adat communities and customary forests in Indonesia. Asia Pac. J. Anthropol. 20, 487–506 (2019).
    Google Scholar 
    56.Malavasi, M. The map of biodiversity mapping. Biol. Conserv. 252, 108843 (2020).
    Google Scholar 
    57.Witter, R. & Satterfield, T. The ebb and flow of indigenous rights recognitions in conservation policy: indigenous rights recognitions in conservation policy. Dev. Change 50, 1083–1108 (2019).
    Google Scholar 
    58.Dutta, A. et al. Response to a “global safety net” to reverse biodiversity loss and stabilize Earth’s climate. Sci. Adv. 6, eabb2824 (2021).
    Google Scholar 
    59.Herrera, D., Pfaff, A. & Robalino, J. Impacts of protected areas vary with the level of government: comparing avoided deforestation across agencies in the Brazilian Amazon. Proc. Natl Acad. Sci. USA 116, 14916–14925 (2019).CAS 

    Google Scholar 
    60.Bebbington, A. J. et al. Resource extraction and infrastructure threaten forest cover and community rights. Proc. Natl Acad. Sci. USA 115, 13164–13173 (2018).CAS 

    Google Scholar 
    61.Johnson, C. J., Venter, O., Ray, J. C. & Watson, J. E. M. Growth‐inducing infrastructure represents transformative yet ignored keystone environmental decisions. Conserv. Lett. https://doi.org/10.1111/conl.12696 (2020).62.Davis, K. F., Yu, K., Rulli, M. C., Pichdara, L. & D’Odorico, P. Accelerated deforestation driven by large-scale land acquisitions in Cambodia. Nat. Geosci. 8, 772–775 (2015).CAS 

    Google Scholar 
    63.Conigliani, C., Cuffaro, N. & D’Agostino, G. Large-scale land investments and forests in Africa. Land Use Policy 75, 651–660 (2018).
    Google Scholar 
    64.Global Land Analysis & Discovery. Global 2010 Tree Cover (30m) (Department of Geographical Sciences, Univ. Maryland, 2013).65.Global Forest Watch. Tree Cover Loss version 1.6 (World Resources Institute, 2019).66.Hansen, M. C., Stehman, S. V. & Potapov, P. V. Quantification of global gross forest cover loss. Proc. Natl Acad. Sci. USA 107, 8650–8655 (2010).CAS 

    Google Scholar 
    67.Protected Planet: The World Database on Protected Areas (WDPA) (UNEP-WCMC & IUCN, accessed January 2020; www.protectedplanet.net68.Hanson, J. O. wdpar: Interface to the world database on protected areas (CRAN, 2020); https://CRAN.R-project.org/package=wdpar69.Global Forest Watch. Spatial Database of Planted Trees (World Resources Institute, data aaccessed May 2021).70.Transparent World & Global Forest Watch. Tree Plantations (World Resources Institute, date accessed May 2021).71.Nelson, A. & Chomitz, K. M. Effectiveness of strict vs. multiple use protected areas in reducing tropical forest fires: a global analysis using matching methods. PLoS ONE 6, e22722 (2011).CAS 

    Google Scholar 
    72.Joppa, L. N. & Pfaff, A. High and far: biases in the location of protected areas. PLoS ONE 4, e8273 (2009).
    Google Scholar 
    73.Global Forest Watch. Tree Cover 2000 version 1.2 (World Resources Institute, 2015).74.Amatulli, G. et al. A suite of global, cross-scale topographic variables for environmental and biodiversity modeling. Sci. Data 5, 180040 (2018).
    Google Scholar 
    75.Nelson, A. et al. A suite of global accessibility indicators. Sci. Data 6, 266 (2019).
    Google Scholar 
    76.Global Roads Open Access Data Set Version 1 (gROADSv1) (1980–2010) (NASA SEDAC, 2013).77.Lloyd, C. T., Sorichetta, A. & Tatem, A. J. High resolution global gridded data for use in population studies. Sci. Data 4, 170001 (2017).
    Google Scholar 
    78.GADM Database of Global Administrative Areas version 3.6 (FAO, 2018).79.Ho, D., Imai, K., King, G. & Stuart, E. matchIt: Nonparametric preprocessing for parametric causal inference (CRAN, 2018); https://CRAN.R-project.org/package=MatchIt80.Wood, S. mgcv: Mixed GAM computation vehicle with automatic smoothness estimation (CRAN, 2019); https://CRAN.R-project.org/package=mgcv More