Ecology

Bolliger, J., Kienast, F. & Zimmermann, N. E. Risk of global warming on montane and subalpine forests in Switzerland—A modeling study. Reg. Environ. Change 1, 99–111 (2000).
Google Scholar 
Bugmann, H. & Pfister, Ch. Impacts of interannual climate variability on past and future forest composition. Reg. Environ. Change 1, 112–125 (2000).
Google Scholar 
Becker, A. & Bugmann, H. (eds.) Global change and mountain regions: The Mountain Research Initiative. IHDP Report 13, GTOS Report 28 and IGBP Report 49, Stockholm (2001).Kullman, L. 20th Century climate warming and tree-limit rise in the southern Scandes of Sweden. Ambio 30, 72–80. https://doi.org/10.1579/0044-7447-30.2.72 (2001).CAS 
PubMed 

Google Scholar 
Körner, Ch. & Paulsen, J. A world-wide study of high altitude treeline temperatures. J. Biogeogr. 31, 713–732. https://doi.org/10.1111/j.1365-2699.2003.01043.x (2004).
Google Scholar 
Harsch, M. A. & Bader, M. Y. Treeline form—A potential key to understanding treeline dynamics. Global Ecol. Biogeogr. 20, 582–596. https://doi.org/10.1111/j.1466-8238.2010.00622.x (2011).
Google Scholar 
Tokarczyk, N. Forest encroachment on temperate mountain meadows: scale, drivers, and current research directions. Geogr. Pol. 90, 463–480 (2017).
Google Scholar 
Vitali, A. et al. Pine recolonization dynamics in Mediterranean human-disturbed treeline ecotones. For. Ecol. Manag. 435, 28–37. https://doi.org/10.1016/j.foreco.2018.12.039 (2019).
Google Scholar 
Heikkinen, O., Obrębska-Starkel, B. & Tuhkanen, S. Introduction: the timberline—A changing battlefront. Prace Geograficzne UJ 98, 7–16 (1995).
Google Scholar 
Mattson, J. Human impact on the timberline in the far North of Europe. Zeszyty Naukowe UJ, Prace Geogr. 98, 41–56 (1995).
Google Scholar 
Stanisci, A., Lavieri, D., Acosta, A. & Blasi, C. Structure and diversity trends at Fagus timberline in central Italy. Community Ecol. 1, 133–138 (2000).
Google Scholar 
Gehrig-Fasel, J., Guisan, A. & Zimmermann, N. E. Tree line shifts in the Swiss Alps: Climate change or land abandonment?. J. Veg. Sci. 18, 571–582 (2007).
Google Scholar 
Feurdean, A. et al. Long-term land-cover/use change in a traditional farming landscape in Romania inferred from pollen data, historical maps and satellite images. Reg. Environ. Change 17, 2193–2207. https://doi.org/10.1007/s10113-016-1063-7 (2017).
Google Scholar 
Burga, C. A., Bührer, S. & Klötzli, F. Mountain ash (Sorbus aucuparia) forests of the Central and Southern Alps (Grisons and Ticino, Switzerland-Prov. Verbano-Cusio-Ossola, N-Italy): Plant ecological and phytosociological aspects. Tuexenia 39, 121–138 (2019).
Google Scholar 
Slayter, R. O. & Noble, I. R. Dynamics of Montane Treelines. In Landscape Boundaries, Consequences for Biotic Diversity and Ecological Flows. Ecological Studies Vol. 92 (eds Hansen, A. J. & di Castri, F.) 346–359 (Springer-Verlag, 1992).
Google Scholar 
Bryn, A. Recent forest limit changes in south-east Norway: Effects of climate change or regrowth after abandoned utilisation?. Nor. Geogr. Tidsskr. 62(4), 251–270. https://doi.org/10.1080/00291950802517551 (2008).
Google Scholar 
Lu, X., Liang, E., Wang, Y., Babst, F. & Camarero, J. J. Mountain treelines climb slowly despite rapid climate warming. Glob. Ecol. Biogeogr. 30(1), 305–315. https://doi.org/10.1111/geb.13214 (2021).
Google Scholar 
Armand, A. D. Sharp and Gradual Mountain Timberlines as Result of species Interaction. Landscape Boundaries, Consequences for Biotic Diversity and Ecological Flows. In Ecological Studies Vol. 92 (eds Hansen, A. J. & di Castri, F.) 360–377 (Springer-Verlag, 1992).
Google Scholar 
Kucharzyk, S. Ekologiczne znaczenie drzewostanów w strefie górnej granicy lasu w Karpatach Wschodnich i ich wrażliwość na zmiany antropogeniczne [Ecological importance of stands at the upper forest limit in the Eastern Carpathians and their sensibility to anthropogenic changes]. Roczn. Bieszcz. 14, 15–43 (2006) (in Polish with English summary).
Google Scholar 
Surina, B. & Rakaj, M. Subalpine beech forest with Hairy alpenrose (Polysticho lonchitis-Fagetum Rhododendretosum hirsuti subass. nova) on Mt. Snežnik (Liburnian Karst, Dinaric Mts). Hacquetia 6, 195–208 (2007).
Google Scholar 
Kucharzyk, S. Zmiany przebiegu górnej granicy lasu w pasmie Szerokiego Wierchu w Bieszczadzkim Parku Narodowym [Changes of upper forest limit in the Szeroki Wierch range (Bieszczady National Park)]. Roczn. Bieszcz. 12, 81–102 (2004) (in Polish with English summary).
Google Scholar 
Kucharzyk, S. & Augustyn, M. Dynamika górnej granicy lasu w Bieszczadach Zachodnich – zmiany w ciągu półtora wieku [The upper forest limit dynamics in the Western Bieszczady Mts.—Changes over a century and a half]. Stud. Nat. 54, 133–156 (2008) (in Polish with English summary).
Google Scholar 
Kubijowicz, W. Życie pasterskie w Beskidach Wschodnich [La Vie Pastorale dans les Beskides Orientales]. Prace Instytutu Geograficznego UJ 5, 3–30 (1926) (in Polish).
Google Scholar 
Zarzycki, K. Lasy Bieszczadów Zachodnich [The forests of the Western Bieszczady Mts (Polish Eastern Carpathians)]. Acta Agr. et Silv. Ser. Leśna 3, 1–131 (1963) (in Polish with English summary).
Google Scholar 
Augustyn, M. Połoniny w Bieszczadach Zachodnich [Almen im westlichen Bieszczady-Gebirge]. Materiały Muzeum Budownictwa Ludowego w Sanoku 31, 88–98 (1993) (in Polish with German summary).
Google Scholar 
Winnicki, T. Zbiorowiska roślinne połonin Bieszczadzkiego Parku Narodowego (Bieszczady Zachodnie, Karpaty Wschodnie) [Plant communities of subalpine poloninas in the Bieszczady National Park (Western Bieszczady Mts, Eastern Carpathians)]. Monogr. Bieszczadzkie 4, 1–215 (1999) (in Polish with English summary).
Google Scholar 
Mróz, W. Zróżnicowanie szaty roślinnej przy górnej granicy lasu w Bieszczadach Wschodnich i Zachodnich [The diversity of vegetation near the upper timberline in the Eastern and the Western Bieszczady Mts]. Roczn. Bieszcz. 14, 45–62 (2006) (in Polish with English summary).
Google Scholar 
Augustyn, M. & Kucharzyk, S. Górna granica lasu na terenie wsi Ustrzyki Górne i Wołosate w końcu XVIII wieku [Timberline in the Western Bieszczady Mts.]. Roczn. Bieszcz. 20, 15–27 (2012) (in Polish with English summary).
Google Scholar 
Jeník, J. Succession on the Połonina Balds in the Western Bieszczady, the Eastern Carpathians. Tuexenia 3, 207–216 (1983).
Google Scholar 
Michalik, S. & Szary, A. Zbiorowiska leśne Bieszczadzkiego Parku Narodowego [The forest communities of the Bieszczady National Park]. Monogr. Bieszcz. 1, 1–175 (1997).
Google Scholar 
Zemanek, B. & Winnicki, T. Rośliny naczyniowe Bieszczadzkiego Parku Narodowego [Vascular plants of the Bieszczady National Park]. Monogr. Bieszcz. 3, 1–249 (1999) (in Polish with English summary).
Google Scholar 
Kucharzyk, S. & Augustyn, M. Trwałość polan reglowych w Bieszczadzkim Parku Narodowym [Stability of mountain glades in the Bieszczady National Park]. Roczn. Bieszcz. 18, 45–58 (2010) (in Polish with English summary).
Google Scholar 
Durak, T., Żywiec, M. & Ortyl, B. Rozprzestrzenianie się zarośli drzewiastych w piętrze połonin Bieszczad Zachodnich [Expansion of brushwood in the subalpine zone of the Western Bieszczady Mts]. Sylwan 157, 130–138 (2013) (in Polish with English summary).
Google Scholar 
Durak, T., Żywiec, M., Kapusta, P. & Holeksa, J. Impact of land use and climate changes on expansion of woody species on subalpine meadows in the Eastern Carpathians. For. Ecol. Manag. 339, 127–135. https://doi.org/10.1016/j.foreco.2014.12.014 (2015).
Google Scholar 
Durak, T., Żywiec, M., Kapusta, P. & Holeksa, J. Rapid spread of a fleshy-fruited species in abandoned subalpine meadows—Formation of an unusual forest belt in the eastern Carpathians. iForest – Biogeosci. For. 9, 337–343. https://doi.org/10.3832/ifor1470-008 (2015).
Google Scholar 
Wężyk, P. & Hawryło, P. Analiza struktury 3D drzewostanów Bieszczadzkiego PN na podstawie danych lotniczego skanowania laserowego oraz ortofotomap lotniczych CIR [3D structure analysis of stands of the Bieszczady National Park on the basis of airborne laser scanning data and CIR aerial ortho-photomaps] (ProGea Consulting, 2015) (in Polish).Anselin, L. Local indicators of spatial association—LISA. Geogr. Anal. 27, 93–115. https://doi.org/10.1111/j.1538-4632.1995.tb00338.x (1995).
Google Scholar 
Scott, L. M. & Janikas, M. V. Spatial Statistics in ArcGIS. In Handbook of Applied Spatial Analysis (eds Fischer, M. M. & Getis, A.) 27–41 (Springer, 2010).
Google Scholar 
Cui, H., Wu, L., Hu, S., Lu, R. & Wang, S. Research on the driving forces of urban hot spots based on exploratory analysis and binary logistic regression model. Trans. GIS 25(3), 1522–1541. https://doi.org/10.1111/tgis.12739 (2021).
Google Scholar 
Pierce, K. B., Lookingbill, T. & Urban, D. A simple method for estimating potential relative radiation (PRR) for landscape-scale vegetation analysis. Landsc. Ecol. 20, 137–147 (2005).
Google Scholar 
Riley, S. J., DeGloria, S. D. & Elliot, R. A terrain ruggedness index that quantifies topographic heterogeneity. Int. J. Sc. 5, 23–27 (1999).
Google Scholar 
Böhner, J. & Antonić, O. Land-surface parameters specific to topo-climatology. Geomorphometry – Concepts, Softw. Appl. Dev. Soil Sci. 33, 195–226. https://doi.org/10.1016/S0166-2481(08)00008-1 (2009).
Google Scholar 
R Core Team. R: A Language and Environment for Statistical Computing (R Core Team, 2021).
Google Scholar 
Agresti, A. An Introduction to Categorical Data Analysis 2nd edn. (Wiley & Sons Inc., 2007).MATH 

Google Scholar 
Cottrell, A. Gnu Regression, Econometrics and Time-series Library gretl. http://gretl.sourceforge.net/(2020).Hellevik, O. Linear versus logistic regression when the dependent variable is a dichotomy. Qual. Quant. 43, 59–74 (2009).
Google Scholar 
Azen, R. & Traxel, N. Using dominance analysis to determine predictor importance in logistic regression. J. Educ. Behav. Stat. 34, 319–347. https://doi.org/10.3102/1076998609332754 (2009).
Google Scholar 
Borcard, P., Legendre, P. & Drapeau, P. Partialling out the spatial component of ecological variation. Ecology 73, 1045–1055 (1992).
Google Scholar 
Przybylska, K. & Kucharzyk, S. Skład gatunkowy i struktura lasów Bieszczadzkiego Parku Narodowego [Species composition and structure of forest of the Bieszczady National Park. Monogr. Bieszcz. 6, 1–159 (1999) (in Polish with English summary).
Google Scholar 
Bader, M. Y. et al. A global framework for linking alpine-treeline ecotone patterns to underlying processes. Ecography 44(2), 265–292. https://doi.org/10.1111/ecog.05285 (2021).
Google Scholar 
Nowosad, M. Zarys klimatu Bieszczadzkiego Parku Narodowego i jego otuliny w świetle dotychczasowych badań [Outlines of climate of the Bieszczady National Park and its bufferzone in the light of previous studies]. Roczn. Bieszcz. 4, 163–183 (1995) (in Polish with English summary).
Google Scholar 
Nowosad, M. & Wereski, S. Warunki klimatyczne. Bieszczadzki Park Narodowy–40 lat ochrony [Climatic conditions. Bieszczady National Park–40 years of protection]. In Bieszczadzki Park Narodowy [The Bieszczady National Park] (eds Górecki, A. & Zemanek, B.) 31–38 (Wyd. Bieszczadzki Park Narodowy, 2016) (in Polish with English summary).
Google Scholar 
Kukulak, J. Neotectonics and planation surfaces in the High Bieszczady Mountains (Outer Carpathians, Poland). Ann. Soc. Geol. Pol. 74, 339–350 (2004).
Google Scholar 
Haczewski, G., Kukulak, J. & Bąk, K. Budowa geologiczna i rzeźba Bieszczadzkiego Parku Narodowego [Geology and relief of the Bieszczady National Park]. Prace monograficzne (Akademia Pedagogiczna im. Komisji Edukacji Narodowej w Krakowie) 468, 1–156 (2007) (in Polish with English summary).
Google Scholar 
Skiba, S., Drewnik, M., Kacprzak, A. & Kołodziejczyk, M. Gleby litogeniczne Bieszczadów i Beskidu Niskiego [Lithogenous soils of the Bieszczady and Beskid Niski Mts (Polish Carpathians)]. Roczn. Bieszcz. 7, 387–396 (1998) (in Polish with English summary).
Google Scholar 
Skiba, S. & Winnicki, T. Gleby zbiorowisk roślinnych bieszczadzkich połonin [Soils of the subalpine meadows plant communities in the Bieszczady Mts]. Roczn. Bieszcz. 4, 97–109 (1995) (in Polish with English summary).
Google Scholar 
Musielok, Ł, Drewnik, M., Szymański, W. & Stolarczyk, M. Classification of mountain soils in a subalpine zone—A case study from the Bieszczady Mountains (SE Poland). Soil Sci. Annu. 70, 170–177. https://doi.org/10.2478/ssa-2019-0015 (2019).CAS 

Google Scholar 
Spatz, G. Succession patterns on mountain pastures. Vegetatio 43, 39–41 (1980).
Google Scholar 
Kozak, J. Zmiany powierzchni lasów w Karpatach Polskich na tle innych gór świata [Changes in the Land Cover in the Polish Carpathians at the Turn of the 20th and 21st Century in Relation to Local Development Level]. Wydawnictwo Uniwersytetu Jagiellońskiego, Kraków (2005) (in Polish with English summary).Vitali, A., Urbinati, C., Weisberg, P. J., Urza, A. K. & Garbarino, M. Effects of natural and anthropogenic drivers on land-cover change and treeline dynamics in the Apennines (Italy). J. Veg. Sci. 29(2), 189–199. https://doi.org/10.1111/jvs.12598 (2018).
Google Scholar 
Micu, D. M., Dumitrescu, A., Cheval, S., Nita, I.-A. & Birsan, M.-V. Temperature changes and elevation-warming relationships in the Carpathian Mountains. Int. J. Climatol. 41, 2154–2172. https://doi.org/10.1002/joc.6952 (2020).
Google Scholar 
Rehman, A. Ziemie dawnej Polski. Cz. I. Karpaty [The lands of ancient Poland. Part I. The Carpathians]. (Gubrynowicz i Schmidt, Lwów) (1895) (in Polish).Frey, W. The influence of snow on growth and survival of planted trees. Arct. Alp. Res. 15, 241–251 (1983).
Google Scholar 
Malanson, G. P. et al. Alpine treeline of Western North America: Linking organism-to-landscape dynamics. Phys. Geogr. 28, 378–396. https://doi.org/10.2747/0272-3646.28.5.378 (2007).
Google Scholar 
Holtmeier, F. K. & Broll, G. Wind as an ecological agent at treelines in North America, the Alps, and the European Subarctic. Phys. Geogr. 31, 203–233. https://doi.org/10.2747/0272-3646.31.3.203 (2010).
Google Scholar 
Barclay, A. M. & Crawford, R. M. M. Winter desiccation stress and resting bud viability in relation to high altitude survival in Sorbus aucuparia L. Flora 172, 21–34 (1982).
Google Scholar 
Raspé, O., Findlay, C. & Jacquemart, A. L. Sorbus aucuparia L. J. Ecol. 88, 910–930 (2000).
Google Scholar 
Zerbe, S. On the ecology of Sorbus aucuparia (Rosaceae) with special regard to germination, establishment and growth. Pol. Bot. J. 46, 229–239 (2001).
Google Scholar 
Smith, W. K., Germino, M. J., Hancock, T. E. & Johnson, D. M. Another perspective on altitudinal limits of alpine timberlines. Tree Physiol. 23, 1101–1112 (2003).PubMed 

Google Scholar 
Trant, A., Higgs, E. & Starzomski, B. M. A century of high elevation ecosystem change in the Canadian Rocky Mountains. Sci. Rep. 10, 9698. https://doi.org/10.1038/s41598-020-66277-2 (2020).ADS 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Barbeito, I., Dawes, M. A., Rixen, C., Senn, J. & Bebi, P. Factors driving mortality and growth at treeline: A 30-year experiment of 92 000 conifers. Ecology 93(2), 389–401 (2012).PubMed 

Google Scholar 
Kullman, L. A 25-year survey of geoecological change in the scandes mountains of Sweden. Geogr. Ann. Ser. B 79, 139–165 (1997).
Google Scholar 
Pękala, K. Rzeźba Bieszczadzkiego Parku Narodowego [Relief of the Bieszczady National Park]. Roczn. Bieszcz. 6, 19–38 (1997) (in Polish with English summary).
Google Scholar 
Kullman, L. Temporal and spatial aspects of subalpine populations of Sorbus aucuparia in Sweden. Ann. Bot. Fenn. 23, 267–275 (1986).
Google Scholar 
Hoersch, B. Modelling the spatial distribution of montane and subalpine forests in the Central Alps using digital elevation models. Ecol. Model. 168, 267–282 (2003).
Google Scholar 
Resler, L. M., Butler, D. R. & Malanson, G. P. Topographic shelter and conifer establishment and mortality in an alpine environment, Glacier National Park, Montana. Phys. Geogr. 26, 112–125 (2005).
Google Scholar 
Kollmann, J. Regeneration window for fleshy-fruited plants during scrub development on abandoned grassland. Ecoscience 2, 213–222 (1995).
Google Scholar 
Lediuk, K. D., Damascos, M. A., Puntieri, J. G. & de Torres Curth, M. I. Population dynamics of an invasive tree, Sorbus aucuparia, in the understory of a Patagonian forest. Plant Ecol. 217, 899–911 (2016).
Google Scholar 
McCutchan, M. H. & Fox, D. G. Effect of elevation and aspect on wind, temperature and humidity. J. Appl. Meteorol. Climatol. 25(12), 1996–2013 (1986).ADS 

Google Scholar 
Stage, A. R. & Salas, C. Interactions of elevation, aspect, and slope in models of forest species composition and productivity. For. Sci. 53, 486–492 (2007).
Google Scholar 
Pocewicz, A. L., Gessler, P. & Robinson, A. P. The relationship between effective plant area index and Landsat spectral response across elevation, solar insolation, and spatial scales in a northern Idaho forest. Can. J. For. Res. 34, 465–480 (2004).
Google Scholar 
Kucharzyk, S. & Sugiero, D. Zróżnicowanie dynamiki procesów lasotwórczych w buczynach bieszczadzkich w zależności od wystawy i wzniesienia [Variability of the dynamics of forest development processes in the Bieszczady beech forests in relation to exposition and altitude]. Sylwan 7, 29–38 (2007) (in Polish with English summary).
Google Scholar 
Drewnik, M., Musielok, Ł, Stolarczyk, M., Mitka, J. & Gus, M. Effects of exposure and vegetation type on organic matter stock in the soils of subalpine meadows in the Eastern Carpathians. CATENA 147, 167–176. https://doi.org/10.1016/j.catena.2016.07.014 (2016).CAS 

Google Scholar 
Zheng, L. et al. Tree regeneration patterns on contrasting slopes at treeline ecotones in Eastern Tibet. Forests 12, 1605. https://doi.org/10.3390/f12111605 (2021).
Google Scholar  More

The studied viviparous clausiliids developed four types of morphological adaptations that facilitate the delivery of embryos through the shell aperture: (1) reduction of the clausiliar apparatus, (2) decrease of embryonic shell width, (3) widening of the shell canal, and (4) development of a flexible embryonic shell.Reduction of the clausiliar apparatusMembers of the Reinia genus, arboreal species from Japan (Fig. 1), show the most advanced adaptations to live-bearing compared to hypothetical ancestral Phaedusinae. The shell shape in these species is more conical than fusiform, the number of whorls decreases, and the aperture widens. One of the species, R. variegata, features almost full reduction of the clausiliar apparatus that consists of only vestigial folds (Fig. 1F). This species also lacks the clausilium, so the entrance through the aperture is unprotected.Figure 1Different stages of reduction of apertural barriers in members of genus Reinia: R. ashizuriensis (A–C; upper row) and R. variegata (D–F; lower row). (A,D) Adult shells; (B,C,E,F) adult shells with body whorl cut open dorsally in microCT visualisation. cp clausilium plate, il inferior lamella, pr principal plica, sc subcolumellar lamella, sl superior lamella, sp spiral lamella, upp upper palatal plica.Full size imageDecrease of embryonic shell widthAnother adaptation concerns the shape of the embryonic shell (“protoconch”), which becomes very narrow in some viviparous species. This feature is conspicuous because embryonic whorls remain in the adult shell as apical whorls. For instance in S. addisoni (Fig. 2A–D), the apical part being much narrower than the first whorls of the teleoconch is a clear evidence that the growth trajectory has changed abruptly after birth. Other examples include E. cylindrella and E. steetzneri, in which both the protoconch and the teleoconch are very narrow, yet at the borderline between these parts, the shell axis is slightly bent (Fig. 2E–L). We suppose that this feature develops as a result of obstruction during birth.Figure 2Width difference between protoconch and teleoconch in Stereophaedusa addisoni (A–D, upper row), Euphaedusa cylindrella (E–H, middle row), Euphaedusa steetzneri (I–L, lower row). (A,C,E,G,I,K) Adult shells with very narrow apical whorls; (B,F,J) X-rayed adults; (F,J) with retained embryos inside; (D,H,L) X-rays of apical part of adult shell with schematic drawings of a neonate.Full size imageWidening of the shell canalThe third type of adaptation is the widening of the shell canal in the body whorl, allowing for easier passage of the embryo between the lamellae and plicae of the apertural barriers. In this case, the outline of the shell changes only slightly giving the body whorl a more convex appearance. A substantial difference to egg-laying species concerns the apertural barriers: the clausiliar includes a broad clausilium plate and a spirally ascending inferior lamella (Fig. 3A–D). These modifications result in a spacious shell canal in the body whorl, for example in S. addisoni and E. sheridani, that can accommodate the transfer of a large embryo. Table 1 presents neonatal size in these species (shell width ca. 1.2 mm), which is very similar to their clausilium width (ca. 1.1–1.2 mm).Figure 3Two types of clausiliar apparatus occurring in Phaedusinae in microCT visualisation: with spirally ascending inferior lamella and wide clausilium plate (upper row), and with straight ascending inferior lamella and narrow clausilium plate (lower row). (A) T. sheridani adult shell with the body whorl cut open dorsally; (B) clausilium of T. sheridani; (C) clausilium of S. addisoni; (D) clausilium of R. ashizuriensis; (E) Zaptyx ventriosa adult shell with body whorl cut open dorsally; (F) clausilium of Z. ventriosa; (G,H) clausilia of O. miranda. Note, that all depicted species are viviparous.Full size imageTable 1 Shell size of studied Phaedusinae species.Full size tableMost viviparid clausiliids develop one of these three types of modification; some adaptations co-occur within a single species, for example a wide clausilium accompanies a narrow apex. Interestingly, the Reinia genus includes taxa with a gradual escalation of viviparity-related adaptations: R. ashizurensis, with a stout shell shape and a low number of whorls, has fully developed apertural barriers with a broad clausilium plate (Fig. 1A–C), while its congener, R. variegata, has reduced apertural barriers (Fig. 1D–F).Development of a flexible embryonic shellThe fourth type of adaptation found in Phaedusinae concerns the structure of the embryonic shells. We report this adaptation in O. miranda and Z. ventriosa.Oospira miranda is a dextral, often decollated, ground-dwelling species from Vietnam (Fig. 4A). The species is viviparous: during microCT scanning of museum specimens, we found embryos within a parental shell (Fig. 4B); in laboratory culture, we observed neonates immediately after live birth (Fig. 4C,D). Morphological characters recognized in the adult shell, i.e., a wide apex (= wide embryonic shell), straightly ascending inferior lamella, and a narrow clausilium plate (Fig. 3G,H), seemed to exclude the possibility of live-bearing reproduction, as embryos are too large to pass through the shell canal at the narrowest point. The height and width of the neonatal shell (mean values: 5.19 mm, 3.59 mm) evidently exceeds the width of the clausilium plate in this species (1.97 mm) (Table 1). However, under closer examination, we found the shell to be thin and delicate, which we refer to as a ‘soft shell’. In direct examination, the neonatal shell of O. miranda resembles cellophane, which may keep a given shape for a long time but becomes distorted already under slight pressure.Figure 4Viviparous clausiliids and their ‘soft-shelled’ neonates born in laboratory culture. (A–D) O. miranda: adult shell, X-rayed shell with embryo visible inside, neonates; (E–H) Z. ventriosa: adult shell, X-rayed shell with eggs visible inside, neonates.Full size imageA similar adaptation exists in Z. ventriosa, a Taiwanese species with a very wide apex, never decollated, a straight ascending inferior lamella, and a narrow clausilium plate (Figs. 3E,F, 4E,F). This species produces neonates in laboratory culture (Fig. 4G–H). The dimensions of the neonates (mean values: height 3.37 mm, width 2.51 mm) exceed at last twofold the width of the clausilium plate (1.08 mm). The shells of such freshly delivered juveniles, when gently touched with laboratory tweezers, became dented, but not fractured. More intense and stronger pressing can break this dentation.These initial observations, that we made during the maintenance of the laboratory culture, suggested that the neonatal shells of O. miranda and Z. ventriosa have flexible walls. These ‘soft-shells’ seem to be highly malleable during the entire embryonic development period and delivery through apertural barriers, hardening shortly after birth. We further investigated the physical properties of the embryonic shell by means of microcomputed tomography and scanning electron microscopy.Microcomputed tomographyWe scanned ‘soft-shelled’ neonates of O. miranda and Z. ventriosa, together with ‘hard-shelled’ embryos and neonates of S. addisoni and T. sheridani, in order to compare the density and thickness of the shells (Fig. 5).Figure 5Comparison of embryonic shell thickness in clausiliids: ‘soft-shelled’ neonates of Z. ventriosa (A,B,G,H) and O. miranda (C,D,I,J); “hard-shelled” neonate of S. addisoni (E,K) and embryo of T. sheridani (F,L) scanned inside a parental shell. Upper row—microCT visualisation of shell surface; middle row—microCT sections of those specimens; (M–O) X-ray photographs of S. addisoni (embryo from dissected adult) and Z. ventriosa (neonate) enlarged in (N,O), respectively, showing the difference in shell density and thickness; (P) microCT based volume rendering of O. miranda (left) and S. addisoni (right) neonates, showing difference between relative density of their shells.Full size imagePreliminary observations using the two-dimensional X-ray photographs showed a difference in thickness and density between S. addisoni and Z. ventriosa (Fig. 5M, enlarged in N and O, respectively). The 3D visualization of O. miranda and S. addisoni (the same microCT scanning and reconstruction parameters) confirmed the difference between density and shell thickness of these two species (Fig. 5P).Due to variations in wall thickness within the neonatal shell (e.g., between the first and the second whorls), it is not possible to precisely determine the thickness of the shell wall. The accuracy of the measurement is also limited by the resolution of the microCT scans, especially in the case of the relatively large neonates of O. miranda and Z. ventriosa. When scanning the whole embryonic shell of Z. ventriosa (approximately 3.5 mm in height), the size of the voxel was approximately 1 µm. Thus, we cannot determine the shell thickness down to the nearest micron, but we can estimate it from a few to a dozen microns. A direct comparison between virtual microCT sections of specimens scanned under the same conditions shows a clear difference between the ‘soft-shelled’ and ‘hard-shelled’ taxa (Fig. 5G–L). The ’hard-shelled’ neonates have a shell wall of 30–40 µm thick. We examined the sequence of three ’soft-shelled’ O. miranda specimens that differed in size (the exact time of birth of each of the cultured neonates is unknown, ca. 1–2 days). The larger (older) the neonate was, the thicker the shell. The shell of the largest of the studied O. miranda was up to 20 µm thick. However, the shell wall of this relatively large juvenile (several millimeters in height) still did not reach the thickness of the small ’hard-shelled’ T. sheridani embryo, which was already about 30–40 µm thick, stiff and rigid during the retention in the genital tract. The neonates of O. miranda and Z. ventriosa were much larger than the embryos and neonates of S. addisoni and R. variegata (Table 1), however, the former taxa has much thinner shells.Scanning electron microscopyAfter the non-invasive microCT scan, we scanned embryos and neonates using SEM (Fig. 6). The different properties of the shells of Z. ventriosa and O. miranda vs. S. addisoni and R. variegata were already visible during the preparation of the analysis. Under vacuum conditions, the soft shells of Z. ventriosa and O. miranda shrank and crumpled, creating a cellophane-like surface (Fig. 6A). Embryos and neonates of S. addisoni and R. variegata did not require any special preparation and their shell shape remained unchanged under the vacuum conditions applied during the SEM examination (Fig. 6D,E). To reduce the shell deformations, we freeze-dried the next group of thin-shelled neonates prior to SEM analyses (Fig. 6B,C).Figure 6Neonates of O. miranda (A,B,F,I,L,M,O) and Z. ventriosa (C,G,J,P) in direct comparison with hard-shelled embryos and neonates of R. variegata (D,N,Q) and S. addisoni (E,H,K); SEM microphotographs. The vacuum conditions in SEM led to the shrinkage of the thin O. miranda shell (A); freeze-drying of ‘soft-shelled’ neonates prior to SEM imaging reduced the level of deformity (B,C). Contrastingly, R. variegata and S. addisoni shells do not require special preparation and retain their shape (D,E). (F) The dented surface of O. miranda neonate and SEM-close-up (I) on a cross-section of the shell just a few micrometers thick (arrow in F indicates the region enlarged in I). (G,J) Shell of Z. ventriosa in comparison with similarly ornamented fragment of S. addisoni (H,K); note several times thicker shell in the latter (arrows in G,H indicate the regions enlarged in J,K, respectively). (L,M) Inner surface of intact periostracum which still connects two fragments of broken aragonite shell of O. miranda (the arrow in M indicates the region enlarged in L); note the difference between shell thickness in O. miranda (L,M) and R. variegata (N). All observed specimens have similar crossed-lamellar microstructure (L–Q). However, just as shell thickness, also the number of lamellar layers of alternate orientation within the shell differs (L,M,O,P vs N,Q).Full size imageThe SEM studies allowed for complementary measurements of the shells. In the broken fragments of Z. ventriosa and O. miranda, the thickness of the shell wall ranged from 2–3 µm (Fig. 6F,G,I,J,L,M) to 18 µm in the largest neonate of O. miranda (Fig. 6O). The shells of S. addisoni (Fig. 6H,K) and R. variegata (Fig. 6N) are several times thicker.All analyzed samples have a thin ( More

Alerstam, T., Hedenström, A. & Åkesson, S. Long-distance migration: evolution and determinants. Oikos 103, 247–260 (2003).Article 

Google Scholar 
Bauer, S., Lisovski, S. & Hahn, S. Timing is crucial for consequences of migratory connectivity. Oikos 125, 605–612 (2016).Article 

Google Scholar 
Bauer, S. & Hoye, B. J. Migratory animals couple biodiversity and ecosystem functioning worldwide. Science 344, 1242552 (2014).Article 
CAS 
PubMed 

Google Scholar 
Fricke, E. C., Ordonez, A., Rogers, H. S. & Svenning, J. C. The effects of defaunation on plants’ capacity to track climate change. Science 214, 210–214 (2022).Article 

Google Scholar 
Tucker, M. A. et al. Moving in the Anthropocene: global reductions in terrestrial mammalian movements. Science 359, 466–469 (2018).Article 
CAS 
PubMed 

Google Scholar 
Wilcove, D. S. & Wikelski, M. Going, going, gone: is animal migration disappearing? PLoS Biol. 6, e188 (2008).Article 
PubMed 
PubMed Central 

Google Scholar 
Parmesan, C. & Yohe, G. A globally coherent fingerprint of climate change. Nature 421, 37–42 (2003).Article 
CAS 
PubMed 

Google Scholar 
Walther, G. et al. Ecological responses to recent climate change. Nature 4126, 389–395 (2002).Article 

Google Scholar 
Teitelbaum, C. S. et al. Experience drives innovation of new migration patterns of whooping cranes in response to global change. Nat. Commun. 7, 12793 (2016).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Oestreich, W. K., Chapman, M. S. & Crowder, L. B. A comparative analysis of dynamic management in marine and terrestrial systems. Front. Ecol. Environ. 18, 496–504 (2020).Article 

Google Scholar 
Senzaki, M. et al. Sensory pollutants alter bird phenology and fitness across a continent. Nature 587, 605–609 (2020).Article 
CAS 
PubMed 

Google Scholar 
Guerra, A. S. Wolves of the sea: managing human–wildlife conflict in an increasingly tense ocean. Mar. Policy 99, 369–373 (2019).Article 

Google Scholar 
Abrahms, B. Human–wildlife conflict under climate change. Science 373, 484–485 (2021).Article 
CAS 
PubMed 

Google Scholar 
Both, C., Bouwhuis, S., Lessells, C. M. & Visser, M. E. Climate change and population declines in a long-distance migratory bird. Nature 441, 81–83 (2006).Article 
CAS 
PubMed 

Google Scholar 
Post, E. & Forchhammer, M. C. Climate change reduces reproductive success of an Arctic herbivore through trophic mismatch. Phil. Trans. R. Soc. B Biol. Sci. 363, 2369–2375 (2008).Article 

Google Scholar 
Winkler, D. W. et al. Cues, strategies, and outcomes: how migrating vertebrates track environmental change. Mov. Ecol. 2, 10 (2014).Article 

Google Scholar 
Xu, W. et al. The plasticity of ungulate migration in a changing world. Ecology 102, e03293 (2021).Article 
PubMed 

Google Scholar 
McNamara, J. M., Barta, Z., Klaassen, M. & Bauer, S. Cues and the optimal timing of activities under environmental change. Ecol. Lett. 14, 1183–1190 (2011).Article 
PubMed 
PubMed Central 

Google Scholar 
Bauer, S., McNamara, J. M. & Barta, Z. Environmental variability, reliability of information and the timing of migration. Proc. R. Soc. B Biol. Sci. 287, 20200622 (2020).Article 

Google Scholar 
Abrahms, B. et al. Emerging perspectives on resource tracking and animal movement ecology. Trends Ecol. Evol. 36, 308–320 (2020).Article 
PubMed 

Google Scholar 
Visser, M. E., Holleman, L. J. M. & Gienapp, P. Shifts in caterpillar biomass phenology due to climate change and its impact on the breeding biology of an insectivorous bird. Oecologia 147, 164–172 (2006).Article 
PubMed 

Google Scholar 
Aikens, E. O. et al. Wave-like patterns of plant phenology determine ungulate movement tactics. Curr. Biol. 30, 3444–3449 (2020).Article 
CAS 
PubMed 

Google Scholar 
Abrahms, B. et al. Memory and resource tracking drive blue whale migrations. Proc. Natl Acad. Sci. USA 116, 5582–5587 (2019).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Lank, D. B., Butler, R. W., Ireland, J. & Ydenberg, R. C. Effects of predation danger on migration strategies of sandpipers. Oikos 103, 303–319 (2003).Article 

Google Scholar 
Sabal, M. C. et al. Predation landscapes influence migratory prey ecology and evolution. Trends Ecol. Evol. 36, 737–749 (2021).Article 
PubMed 

Google Scholar 
Furey, N. B., Armstrong, J. B., Beauchamp, D. A. & Hinch, S. G. Migratory coupling between predators and prey. Nat. Ecol. Evol. 2, 1846–1853 (2018).Article 
PubMed 

Google Scholar 
Altizer, S., Bartel, R. & Han, B. A. Animal migration and infectious disease risk. Science 331, 296–302 (2011).Article 
CAS 
PubMed 

Google Scholar 
Gunnarsson, T., Gill, J., Sigurbjörnsson, T. & Sutherland, W. Arrival synchrony in migratory birds. Nature 431, 646 (2004).Article 
CAS 
PubMed 

Google Scholar 
Beltran, R. S. et al. Elephant seals time their long-distance migration using a map sense. Curr. Biol. 32, R156–R157 (2022).Article 
CAS 
PubMed 

Google Scholar 
Yang, L. H. & Rudolf, V. H. W. Phenology, ontogeny and the effects of climate change on the timing of species interactions. Ecol. Lett. 13, 1–10 (2010).Article 
CAS 
PubMed 

Google Scholar 
Visser, M. E. & Gienapp, P. Evolutionary and demographic consequences of phenological mismatches. Nat. Ecol. Evol. 3, 879–885 (2019).Article 
PubMed 
PubMed Central 

Google Scholar 
Furey, N. B. et al. Predator swamping reduces predation risk during nocturnal migration of juvenile salmon in a high-mortality landscape. J. Anim. Ecol. 85, 948–959 (2016).Article 
PubMed 

Google Scholar 
Rickbeil, G. J. M. et al. Plasticity in elk migration timing is a response to changing environmental conditions. Glob. Change Biol. 25, 2368–2381 (2019).Article 

Google Scholar 
Schmaljohann, H. & Both, C. The limits of modifying migration speed to adjust to climate change. Nat. Clim. Change 7, 573–576 (2017).Article 

Google Scholar 
Gwinner, E. Circadian and circannual programmes in avian migration. J. Exp. Biol. 199, 39–48 (1996).Article 
CAS 
PubMed 

Google Scholar 
Liedvogel, M., Åkesson, S. & Bensch, S. The genetics of migration on the move. Trends Ecol. Evol. 26, 561–569 (2011).Article 
PubMed 

Google Scholar 
Hauser, D. D. W. et al. Decadal shifts in autumn migration timing by Pacific Arctic beluga whales are related to delayed annual sea ice formation. Glob. Change Biol. 23, 2206–2217 (2017).Article 

Google Scholar 
Palacín, C., Alonso, J. C., Alonso, J. A., Magaña, M. & Martín, C. A. Cultural transmission and flexibility of partial migration patterns in a long-lived bird, the great bustard Otis tarda. J. Avian Biol. 42, 301–308 (2011).Article 

Google Scholar 
Couzin, I. D. Collective animal migration. Curr. Biol. 28, R976–R980 (2018).Article 
CAS 
PubMed 

Google Scholar 
Guttal, V. & Couzin, I. D. Social interactions, information use, and the evolution of collective migration. Proc. Natl Acad. Sci. USA 107, 16172–16177 (2010).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Berdahl, A. M. et al. Collective animal navigation and migratory culture: from theoretical models to empirical evidence. Phil. Trans. R. Soc. B Biol. Sci. 373, 20170009 (2018).Article 

Google Scholar 
Cohen, E. B. & Satterfield, D. A. ‘Chancing on a spectacle:’ co-occurring animal migrations and interspecific interactions. Ecography 43, 1657–1671 (2020).Article 

Google Scholar 
Berdahl, A., Torney, C. J., Ioannou, C. C., Faria, J. J. & Couzin, I. D. Emergent sensing of complex environments by mobile animal groups. Science 339, 574–576 (2013).Article 
CAS 
PubMed 

Google Scholar 
Abrahms, B., Teitelbaum, C. S., Mueller, T. & Converse, S. J. Ontogenetic shifts from social to experiential learning drive avian migration timing. Nat. Commun. 12, 7326 (2021).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Sasaki, T. & Biro, D. Cumulative culture can emerge from collective intelligence in animal groups. Nat. Commun. 8, 15049 (2017).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Helm, B., Piersma, T. & van der Jeugd, H. Sociable schedules: interplay between avian seasonal and social behaviour. Anim. Behav. 72, 245–262 (2006).Article 

Google Scholar 
Piersma, T., Zwarts, L. & Bruggemann, J. H. Behavioural aspects of the departure of waders before long-distance flights: flocking, vocalizations, flight paths and diurnal timing. Ardea 78, 157–184 (1990).
Google Scholar 
Dingle, H. & Drake, V. A. What is migration? BioScience 57, 113–121 (2007).Article 

Google Scholar 
Oestreich, W. K. & Aiu, K. M. Code and data from: The influence of social cues on timing of animal migrations. Zenodo https://zenodo.org/record/6574762 (2022).Furey, N. B., Martins, E. G. & Hinch, S. G. Migratory salmon smolts exhibit consistent interannual depensatory predator swamping: effects on telemetry-based survival estimates. Ecol. Freshw. Fish 30, 18–30 (2021).Article 

Google Scholar 
Berdahl, A., Westley, P. A. H. & Quinn, T. P. Social interactions shape the timing of spawning migrations in an anadromous fish. Anim. Behav. 126, 221–229 (2017).Article 

Google Scholar 
Louca, V., Lindsay, S. W. & Lucas, M. C. Factors triggering floodplain fish emigration: importance of fish density and food availability. Ecol. Freshw. Fish 18, 60–64 (2009).Article 
PubMed 
PubMed Central 

Google Scholar 
Bastille-Rousseau, G. et al. Migration triggers in a large herbivore: Galápagos giant tortoises navigating resource gradients on volcanoes. Ecology 100, e02658 (2019).Article 
PubMed 

Google Scholar 
Bracis, C. & Mueller, T. Memory, not just perception, plays an important role in terrestrial mammalian migration. Proc. R. Soc. B Biol. Sci. 284, 20170449 (2017).Article 

Google Scholar 
Barrett, B., Zepeda, E., Pollack, L., Munson, A. & Sih, A. Counter-culture: does social learning help or hinder adaptive response to human-induced rapid environmental change? Front. Ecol. Evol. 7, 183 (2019).Article 

Google Scholar 
Merkle, J. A. et al. Site fidelity as a maladaptive behavior in the Anthropocene. Front. Ecol. Environ. 20, 187–194 (2022).Article 

Google Scholar 
Teske, P. R. et al. The sardine run in southeastern Africa is a mass migration into an ecological trap. Sci. Adv. 7, eabf4514 (2021).Article 
PubMed 
PubMed Central 

Google Scholar 
Corten, A. The role of ‘conservatism’ in herring migrations. Rev. Fish Biol. Fish. 11, 339–361 (2002).Article 

Google Scholar 
Mukhin, A., Chernetsov, N. & Kishkinev, D. Acoustic information as a distant cue for habitat recognition by nocturnally migrating passerines during landfall. Behav. Ecol. 19, 716–723 (2008).Article 

Google Scholar 
Barker, K. J. et al. Toward a new framework for restoring lost wildlife migrations. Conserv. Lett. 15, e12850 (2022).Article 

Google Scholar 
Teitelbaum, C. S., Converse, S. J. & Mueller, T. The importance of early life experience and animal cultures in reintroductions. Conserv. Lett. 12, e12599 (2019).Article 

Google Scholar 
Hughey, L. F., Hein, A. M., Strandburg-Peshkin, A., Jensen, F. H. & Hughey, L. F. Challenges and solutions for studying collective animal behaviour in the wild. Phil. Trans. R. Soc. B 373, 20170005 (2018).Article 
PubMed 
PubMed Central 

Google Scholar 
Calabrese, J. M. et al. Disentangling social interactions and environmental drivers in multi-individual wildlife tracking data. Phil. Trans. R. Soc. B 373, 20170007 (2018).Article 
PubMed 
PubMed Central 

Google Scholar 
Jesmer, B. R. et al. Is ungulate migration culturally transmitted? Evidence of social learning from translocated animals. Science 361, 1023–1025 (2018).Article 
CAS 
PubMed 

Google Scholar 
Bousquet, C. A. H., Sumpter, D. J. T. & Manser, M. B. Moving calls: a vocal mechanism underlying quorum decisions in cohesive groups. Proc. R. Soc. B Biol. Sci. 278, 1482–1488 (2011).Article 

Google Scholar 
Dibnah, A. J. et al. Vocally mediated consensus decisions govern mass departures from jackdaw roosts. Curr. Biol. 32, R455–R456 (2022).Article 
CAS 
PubMed 

Google Scholar 
Robart, A. R., Zuñiga, H. X., Navarro, G. & Watts, H. E. Social environment influences termination of nomadic migration. Biol. Lett. 18, 20220006 (2022).Article 
PubMed 

Google Scholar 
Dodson, S., Abrahms, B., Bograd, S. J., Fiechter, J. & Hazen, E. L. Disentangling the biotic and abiotic drivers of emergent migratory behavior using individual-based models. Ecol. Modell. 432, 109225 (2020).Article 

Google Scholar 
Kays, R., Crofoot, M. C., Jetz, W. & Wikelski, M. Terrestrial animal tracking as an eye on life and planet. Science 348, aaa2478 (2015).Article 
PubMed 

Google Scholar 
Hussey, N. E. et al. Aquatic animal telemetry: a panoramic window into the underwater world. Science 348, 1255642 (2015).Article 
PubMed 

Google Scholar 
Oestreich, W. K. et al. Acoustic signature reveals blue whale tune life history transitions to oceanographic conditions. Funct. Ecol. 36, 882–895 (2022).Article 
CAS 

Google Scholar 
Chapman, J. W., Reynolds, D. R. & Smith, A. D. Vertical-looking radar: a new tool for monitoring high-altitude insect migration. BioScience 53, 503–511 (2003).Article 

Google Scholar 
Oestreich, W. K. et al. Animal-borne metrics enable acoustic detection of blue whale migration. Curr. Biol. 30, 4773–4779 (2020).Article 
CAS 
PubMed 

Google Scholar 
Fraser, K. C., Shave, A., de Greef, E., Siegrist, J. & Garroway, C. J. Individual variability in migration timing can explain long-term, population-level advances in a songbird. Front. Ecol. Evol. 7, 324 (2019).Article 

Google Scholar 
Byholm, P., Beal, M., Isaksson, N., Lötberg, U. & Åkesson, S. Paternal transmission of migration knowledge in a long-distance bird migrant. Nat. Commun. 13, 1566 (2022).Article 
CAS 
PubMed 
PubMed Central 

Google Scholar 
Schneider, S. S. & McNally, L. C. Waggle dance behavior associated with seasonal absconding in colonies of the African honey bee, Apis mellifera scutellata. Insectes Soc. 41, 115–127 (1994).Article 

Google Scholar 
Raveling, D. G. Preflight and flight behavior of Canada geese. Auk 86, 671–681 (1969).Article 

Google Scholar 
Tennessen, J. B., Parks, S. E. & Langkilde, T. Traffic noise causes physiological stress and impairs breeding migration behaviour in frogs. Conserv. Physiol. 2, cou032 (2014).Article 
PubMed 
PubMed Central 

Google Scholar 
Lagarde, A., Lagarde, F. & Piersma, T. Vocal signalling by Eurasian spoonbills Platalea leucorodia in flocks before migratory departure. Ardea 109, 243–250 (2021).Article 

Google Scholar 
Rees, E. C. Conflict of choice within pairs of Bewick’s swans regarding their migratory movement to and from the wintering grounds. Anim. Behav. 35, 1685–1693 (1987).Article 

Google Scholar 
Mazeroll, A. I. & Montgomery, W. L. Daily migrations of a coral reef fish in the Red Sea (Gulf of Aqaba, Israel). Copiea 1998, 893–905 (1998).Article 

Google Scholar 
Méndez, V. et al. Paternal effects in the initiation of migratory behaviour in birds. Sci. Rep. 11, 2782 (2021).Article 
PubMed 
PubMed Central 

Google Scholar 
Nelson, M. E. Development of migratory behavior in northern white-tailed deer. Can. J. Zool. 76, 426–432 (1998).Article 

Google Scholar 
Sweanor, P. Y. & Sandgren, F. Winter-range philopatry of seasonally migratory moose. J. Appl. Ecol. 26, 25–33 (1989).Article 

Google Scholar 
Rees, E. C. Consistency in the timing of migration for individual Bewick’s swans. Anim. Behav. 38, 384–393 (1989).Article 

Google Scholar 
Corten, A. A possible adaptation of herring feeding migrations to a change in timing of the Calanus finmarchicus season in the eastern North Sea. ICES J. Mar. Sci. 57, 1261–1270 (2000).Article 

Google Scholar 
Loonstra, A. J. et al. Individual black-tailed godwits do not stick to single routes: a hypothesis on how low population densities might decrease social conformity. Ardea 107, 251–261 (2020).Article 

Google Scholar 
Hake, M., Kjellén, N. & Alerstam, T. Age‐dependent migration strategy in honey buzzards Pernis apivorus tracked by satellite. Oikos 103, 385–396 (2003).Article 

Google Scholar 
Gupte, P. R., Koffijberg, K., Müskens, G. J. D. M., Wikelski, M. & Kölzsch, A. Family size dynamics in wintering geese. J. Ornithol. 160, 363–375 (2019).Article 

Google Scholar 
Gonçalves, M. I. C. et al. Movement patterns of humpback whales (Megaptera novaeangliae) reoccupying a Brazilian breeding ground. Biota Neotrop. 18, e20180567 (2018).Article 

Google Scholar 
Trudelle, L. et al. First insights on spatial and temporal distribution patterns of humpback whales in the breeding ground at Sainte Marie Channel, Madagascar. Afr. J. Mar. Sci. 40, 75–86 (2018).Article 

Google Scholar 
De La Gala-Hernández, S. R., Heckel, G. & Sumich, J. L. Comparative swimming effort of migrating gray whales (Eschrichtius robustus) and calf cost of transport along Costa Azul, Baja California, Mexico. Can. J. Zool. 86, 307–313 (2008).Article 

Google Scholar 
Sword, G. A. Local population density and the activation of movement in migratory band-forming Mormon crickets. Anim. Behav. 69, 437–444 (2005).Article 

Google Scholar 
Buhl, J. et al. From disorder to order in marching locusts. Science 312, 1402–1406 (2006).Article 
CAS 
PubMed 

Google Scholar 
Mysterud, A., Loe, L. E., Zimmermann, B., Bischof, R. & Meisingset, E. Partial migration in expanding red deer populations at northern latitudes—a role for density dependence? Oikos 120, 1817–1825 (2011).Article 

Google Scholar 
Bukreeva, O. M. & Lidzhi-garyaeva, G. V. Mass migration of social voles (Microtus socialis Pallas, 1773) in the Northwestern Caspian region. Arid Ecosyst. 8, 147–151 (2018).Article 

Google Scholar 
Eggeman, S. L., Hebblewhite, M., Bohm, H., Whittington, J. & Merrill, E. H. Behavioural flexibility in migratory behaviour in a long-lived large herbivore. J. Anim. Ecol. 85, 785–797 (2016).Article 
PubMed 

Google Scholar 
Weithman, C. et al. Senescence and carryover effects of reproductive performance influence migration, condition, and breeding propensity in a small shorebird. Ecol. Evol. 7, 11044–11056 (2017).Article 
PubMed 
PubMed Central 

Google Scholar 
Rappole, J. H. & Warner, D. W. Relationships between behavior, physiology and weather in avian transients at a migration stopover site. Oecologia 212, 193–212 (1976).Article 

Google Scholar 
Fauchald, P., Mauritzen, M. & Gjøsæter, H. Density‐dependent migratory waves in the marine pelagic ecosystem. Ecology 87, 2915–2924 (2006).Article 
PubMed 

Google Scholar 
Makris, N. C. et al. Critical population density triggers rapid formation of vast oceanic fish shoals. Science 323, 1734–1737 (2009).Article 
CAS 
PubMed 

Google Scholar 
Tøttrup, A. P. & Thorup, K. Sex-differentiated migration patterns, protandry and phenology in North European songbird populations. J. Ornithol. 149, 161–167 (2008).Article 

Google Scholar 
Francis, C. M. & Cooke, C. F. Differential timing of spring migration in rose-breasted grosbeaks. J. Field Ornithol. 61, 404–412 (1990).
Google Scholar 
Corgos, A., Verísimo, P. & Freire, J. Timing and seasonality of the terminal molt and mating migration in the spider crab, Maja brachydactyla: evidence of alternative mating strategies. J. Shellfish Res. 25, 577–587 (2006).Article 

Google Scholar 
Gordo, O., Sanz, J. J. & Lobo, J. M. Spatial patterns of white stork (Ciconia ciconia) migratory phenology in the Iberian Peninsula. J. Ornithol. 148, 293–308 (2007).Article 

Google Scholar 
Sergio, F. et al. Individual improvements and selective mortality shape lifelong migratory performance. Nature 515, 410–413 (2014).Article 
CAS 
PubMed 

Google Scholar 
Manica, L. T., Graves, J. A., Podos, J. & Macedo, R. H. Hidden leks in a migratory songbird: mating advantages for earlier and more attractive males. Behav. Ecol. 31, 1180–1191 (2020).Article 

Google Scholar 
Cade, D. E. et al. Social exploitation of extensive, ephemeral, environmentally controlled prey patches by supergroups of rorqual whales. Anim. Behav. 182, 251–266 (2021).Article 

Google Scholar 
Urbanek, R. P., Fondow, L. E. A., Zimorski, S. E., Wellington, M. A. & Nipper, M. A. Winter release and management of reintroduced migratory whooping cranes Grus americana. Bird Conserv. Int. 20, 43–54 (2010).Article 

Google Scholar 
Németh, Z. & Moore, F. R. Information acquisition during migration: a social perspective. Auk 131, 186–194 (2014).Article 

Google Scholar  More

Pascual, U. et al. Biodiversity and the challenge of pluralism. Nat. Sustain. 4, 567–572 (2021).Article 

Google Scholar 
Cumming, G. S. The relevance and resilience of protected areas in the Anthropocene. Anthropocene 13, 46–56 (2016).Article 

Google Scholar 
Cumming, G. S. et al. Understanding protected area resilience: a multi-scale, social-ecological approach. Ecol. Appl. 25, 299–319 (2015).Article 

Google Scholar 
Ellis, E. C. & Mehrabi, Z. Half Earth: promises, pitfalls, and prospects of dedicating half of Earth’s land to conservation. Curr. Opin. Environ. Sustain. 38, 22–30 (2019).Article 

Google Scholar 
Golden Kroner, R. E. et al. The uncertain future of protected lands and waters. Science 364, 881–886 (2019).Article 
CAS 

Google Scholar 
Palfrey, R., Oldekop, J. & Holmes, G. Conservation and social outcomes of private protected areas. Conserv. Biol. 35, 1098–1110 (2021).Article 

Google Scholar 
Gurney, G. G. et al. Biodiversity needs every tool in the box: use OECMs. Nature 595, 646–649 (2021).Article 
CAS 

Google Scholar 
Kremen, C. & Merenlender, A. M. Landscapes that work for biodiversity and people. Science 362, eaau6020 (2018).Article 

Google Scholar 
Taylor, W. A. et al. South Africa’s private wildlife ranches protect globally significant populations of wild ungulates. Biodivers. Conserv. 30, 4111–4135 (2021).Article 

Google Scholar 
Child, B. A., Musengezi, J., Parent, G. D. & Child, G. F. T. The economics and institutional economics of wildlife on private land in Africa. Pastoralism 2, 18 (2012).Article 

Google Scholar 
Kiffner, C. et al. Community-based wildlife management area supports similar mammal species richness and densities compared to a national park. Ecol. Evol. 10, 480–492 (2020).Article 

Google Scholar 
Naidoo, R. et al. Complementary benefits of tourism and hunting to communal conservancies in Namibia. Conserv. Biol. 30, 628–638 (2016).Article 

Google Scholar 
Cousins, J., Sadler, J. & Evans, J. Exploring the role of private wildlife ranching as a conservation tool in South Africa: stakeholder perspectives. Ecol. Soc. 13, 43 (2008).Article 

Google Scholar 
Kamal, S., Grodzińska-Jurczak, M. & Brown, G. Conservation on private land: a review of global strategies with a proposed classification system. J. Environ. Plann. Manage. 58, 576–597 (2015).Article 

Google Scholar 
Ogar, E., Pecl, G. & Mustonen, T. Science must embrace traditional and indigenous knowledge to solve our biodiversity crisis. One Earth 3, 162–165 (2020).Article 

Google Scholar 
De Vos, A. & Cumming, G. S. The contribution of land tenure diversity to the spatial resilience of protected area networks. People Nat. 1, 331–346 (2019).Article 

Google Scholar 
Biggs, R. et al. Toward principles for enhancing the resilience of ecosystem services. Annu. Rev. Environ. Resour. 37, 421–448 (2012).Article 

Google Scholar 
Cumming, G. & Collier, J. Change and identity in complex systems. Ecol. Soc. 10, 29 (2005).Article 

Google Scholar 
Oliver, T. H. et al. Biodiversity and resilience of ecosystem functions. Trends Ecol. Evol. 30, 673–684 (2015).Article 

Google Scholar 
Leslie, P. & McCabe, J. T. Response diversity and resilience in social–ecological systems. Curr. Anthropol. 54, 114–143 (2013).Article 

Google Scholar 
Clements, H. S., Knight, M., Jones, P. & Balfour, D. Private rhino conservation: diverse strategies adopted in response to the poaching crisis. Conserv. Lett. 13, e12741 (2020).Article 

Google Scholar 
Carpenter, S., Walker, B., Anderies, J. M. & Abel, N. From metaphor to measurement: resilience of what to what? Ecosystems 4, 765–781 (2001).Article 

Google Scholar 
Parker, K., De Vos, A., Clements, H. S., Biggs, D. & Biggs, R. Impacts of a trophy hunting ban on private land conservation in South African biodiversity hotspots. Conserv. Sci. Pract. 2, e214 (2020).
Google Scholar 
World Travel & Tourism Council. The Economic Impact of Global Wildlife Tourism (WTTC, 2019).Lindsey, P. et al. Conserving Africa’s wildlife and wildlands through the COVID-19 crisis and beyond. Nat. Ecol. Evol. 4, 1300–1310 (2020).Article 

Google Scholar 
Hambira, W. L., Stone, L. S. & Pagiwa, V. Botswana nature-based tourism and COVID-19: transformational implications for the future. Dev. South. Afr. 39, 51–67 (2021).Article 

Google Scholar 
Mudzengi, B. K., Gandiwa, E., Muboko, N. & Mutanga, C. N. Innovative community ecotourism coping and recovery strategies to COVID-19 pandemic shocks: the case of Mahenye. Dev. South. Afr. 39, 68–83 (2021).Article 

Google Scholar 
Waithaka, J. et al. Impacts of COVID-19 on protected and conserved areas: a global overview and regional perspectives. Parks 27, 41–56 (2021).Article 

Google Scholar 
Smith, M. K. S. et al. Sustainability of protected areas: vulnerabilities and opportunities as revealed by COVID-19 in a national park management agency. Biol. Conserv. 255, 108985 (2021).Article 

Google Scholar 
Miller-Rushing, A. J. et al. COVID-19 pandemic impacts on conservation research, management, and public engagement in US national parks. Biol. Conserv. 257, 109038 (2021).Article 

Google Scholar 
Thurstan, R. H. et al. Envisioning a resilient future for biodiversity conservation in the wake of the COVID‐19 pandemic. People Nat. 3, 990–1013 (2021).Article 

Google Scholar 
Taylor, W. A., Lindsey, P. A., Nicholson, S. K., Relton, C. & Davies-Mostert, H. T. Jobs, game meat and profits: the benefits of wildlife ranching on marginal lands in South Africa. Biol. Conserv. 245, 108561 (2020).Article 

Google Scholar 
Chidakel, A., Eb, C. & Child, B. The comparative financial and economic performance of protected areas in the Greater Kruger National Park, South Africa: functional diversity and resilience in the socio-economics of a landscape-scale reserve network.J. Sustain. Tour. 28, 1100–1119 (2020).Article 

Google Scholar 
Saayman, M., van der Merwe, P. & Saayman, A. The economic impact of trophy hunting in the South African wildlife industry. Glob. Ecol. Conserv. 16, e00510 (2018).Article 

Google Scholar 
Hall, R. A political economy of land reform in South Africa. Rev. Afr. Polit. Econ. 31, 213–227 (2004).Article 

Google Scholar 
Mkhize, N. Game farm conversions and the land question: unpacking present contradictions and historical continuities in farm dwellers’ tenure insecurity in Cradock. J. Contemp. Afr. Stud. 32, 207–219 (2014).Article 

Google Scholar 
Thakholi, L. Conservation labour geographies: subsuming regional labour into private conservation spaces in South Africa. Geoforum 123, 1–11 (2021).Article 

Google Scholar 
Brandt, F. Power battles on South African trophy-hunting farms: farm workers, resistance and mobility in the Karoo. J. Contemp. Afr. Stud. 34, 165–181 (2016).Article 

Google Scholar 
Child, B. & Barnes, G. The conceptual evolution and practice of community-based natural resource management in southern Africa: past, present and future. Environ. Conserv. 37, 283–295 (2010).Article 

Google Scholar 
Clements, H. S., Baum, J. & Cumming, G. S. Money and motives: an organizational ecology perspective on private land conservation. Biol. Conserv. 197, 108–115 (2016).Article 

Google Scholar 
van der Merwe, P., Saayman, A. & Jacobs, C. Assessing the economic impact of COVID-19 on the private wildlife industry of South Africa. Glob. Ecol. Conserv. 28, e01633 (2021).Article 

Google Scholar 
Clements, H. S. & Cumming, G. S. Traps and transformations influencing the financial viability of tourism on private-land conservation areas. Conserv. Biol. 32, 424–436 (2018).Article 

Google Scholar 
Winterbach, C. W., Whitesell, C. & Somers, M. J. Wildlife abundance and diversity as indicators of tourism potential in northern Botswana. PLoS ONE 10, e0135595 (2015).Article 

Google Scholar 
Di Minin, E., Fraser, I., Slotow, R. & MacMillan, D. C. Understanding heterogeneous preference of tourists for big game species: implications for conservation and management. Anim. Conserv. 16, 249–258 (2013).Article 

Google Scholar 
Clements, H. S., Biggs, R. & Cumming, G. S. Cross-scale and social–ecological changes constitute main threats to private land conservation in South Africa. J. Environ. Manag. 274, 111235 (2020).Article 

Google Scholar 
Breen, C. et al. Integrating cultural and natural heritage approaches to marine protected areas in the MENA region. Mar. Policy 132, 104676 (2021).Article 

Google Scholar 
Munasinghe, H. The politics of the past: constructing a national identity through heritage conservation. Int. J. Herit. Stud. 11, 251–260 (2005).Article 

Google Scholar 
MacKinnon, K. et al. Strengthening the global system of protected areas post-2020: a perspective from the IUCN World Commission on Protected Areas. Parks 36, 281–296 (2020).
Google Scholar 
van Kerkhoff, L. et al. Towards future-oriented conservation: managing protected areas in an era of climate change. Ambio 48, 699–713 (2019).Article 

Google Scholar 
De Vos, A. et al. Pathogens, disease, and the social–ecological resilience of protected areas. Ecol. Soc. 21, 20 (2016).Article 

Google Scholar 
Bengtsson, J. et al. Reserves, resilience and dynamic landscapes. Ambio 32, 389–396 (2003).Article 

Google Scholar 
Broom, D. M., Galindo, F. A. & Murgueitio, E. Sustainable, efficient livestock production with high biodiversity and good welfare for animals. Proc. R. Soc. B 280, 20132025 (2013).Article 
CAS 

Google Scholar 
IPBES Global Assessment Report on Biodiversity and Ecosystem Services of the Intergovernmental Science-Policy Platform on Biodiversity and Ecosystem Services (IPBES Secretariat, 2019).Marnewick, D., Jonas, H. & Stevens, C. Site-level Methodology for Identifying Other Effective Area-based Conservation Measures (OECMs) Draft Version 1.0 (IUCN: Gland, Switzerland; 2020).Nalau, J., Becken, S. & Mackey, B. Ecosystem-based adaptation: a review of the constraints. Environ. Sci. Policy 89, 357–364 (2018).Article 

Google Scholar 
Hartung, C. et al. Open Data Kit: tools to build information services for developing regions. In Proc. 4th ACM/IEEE International Conference on Information and Communication Technologies and Development (ed Unwin, T.) pp 1–12 (Association for Computing Machinery, New York, NY, United States; 2010).Oksanen, A. J. et al. Vegan: Community Ecology Package. R version 2.6–2 (2022). https://cran.r-project.org/web/packages/vegan/vegan.pdfWard, J. Hierarchical grouping to optimize an objective function. J. Am. Stat. Assoc. 58, 236–244 (1963).Article 

Google Scholar 
Maechler, M. et al. Cluster: Finding Groups in Data. R version 2.0.3 (2015). https://cran.microsoft.com/snapshot/2015-11-17/web/packages/cluster/cluster.pdfR Core Team R: A Language and Environment for Statistical Computing (R Foundation for Statistical Computing, 2021). More

Tilman, D. & Clark, M. Global diets link environmental sustainability and human health. Nature 515, 518–522 (2014).Article 
CAS 

Google Scholar 
Godfray, H. C. J. et al. Meat consumption, health, and the environment. Science 361, eaam5324 (2018).Article 

Google Scholar 
Hicks, C. C. et al. Harnessing global fisheries to tackle micronutrient deficiencies. Nature 574, 95–98 (2019).Article 
CAS 

Google Scholar 
Willett, W. et al. Food in the Anthropocene: the EAT–Lancet Commission on healthy diets from sustainable food systems. Lancet 393, 447–492 (2019).Article 

Google Scholar 
Maxwell, S. L., Fuller, R. A., Brooks, T. M. & Watson, J. E. M. Biodiversity: the ravages of guns, nets and bulldozers. Nature 536, 143–145 (2016).Article 
CAS 

Google Scholar 
Tilman, D. et al. Future threats to biodiversity and pathways to their prevention. Nature 546, 73–81 (2017).Article 
CAS 

Google Scholar 
Ellis, E. C., Goldewikj, K. K., Siebert, S., Lightman, D. & Ramankutty, N. Anthropogenic transformation of the biomes, 1700 to 2000. Glob. Ecol. Biogeogr. 19, 589–606 (2010).
Google Scholar 
Crippa, M. et al. Food systems are responsible for a third of global anthropogenic GHG emissions. Nat. Food 2, 198–209 (2021).Article 
CAS 

Google Scholar 
Rosegrant, M. W., Ringler, C. & Zhu, T. Water for agriculture: maintaining food security under growing scarcity. Annu. Rev. Environ. Resour. 34, 205–222 (2009).Article 

Google Scholar 
Tubiello, F. N. et al. The contribution of agriculture, forestry and other land use activities to global warming, 1990–2012. Glob. Change Biol. 21, 2655–2660 (2015).Article 

Google Scholar 
Lee, R. Y., Seitzinger, S. & Mayorga, E. Land-based nutrient loading to LMEs: a global watershed perspective on magnitudes and sources. Environ. Dev. 17, 220–229 (2016).Article 

Google Scholar 
Kroodsma, D. A. et al. Tracking the global footprint of fisheries. Science 359, 904–908 (2018).Article 
CAS 

Google Scholar 
McIntyre, P. B., Liermann, C. A. R. & Revenga, C. Linking freshwater fishery management to global food security and biodiversity conservation. Proc. Natl Acad. Sci. USA 113, 12880–12885 (2016).Article 
CAS 

Google Scholar 
Poore, J. & Nemecek, T. Reducing food’s environmental impacts through producers and consumers. Science 360, 987–992 (2018).Article 
CAS 

Google Scholar 
Hilborn, R., Banobi, J., Hall, S. J., Pucylowski, T. & Walsworth, T. E. The environmental cost of animal source foods. Front. Ecol. Environ. 16, 329–335 (2018).Article 

Google Scholar 
Parker, R. W. R. et al. Fuel use and greenhouse gas emissions of world fisheries. Nat. Clim. Change 8, 333–337 (2018).Article 
CAS 

Google Scholar 
Davis, K. F. et al. Meeting future food demand with current agricultural resources. Glob. Environ. Change 39, 125–132 (2016).Article 

Google Scholar 
Gephart, J. A. et al. The environmental cost of subsistence: optimizing diets to minimize footprints. Sci. Total Environ. 553, 120–127 (2016).Article 
CAS 

Google Scholar 
Gephart, J. A. et al. Environmental performance of blue foods. Nature 597, 360–365 (2021).Article 
CAS 

Google Scholar 
Halpern, B. S. et al. Putting all foods on the same table: achieving sustainable food systems requires full accounting. Proc. Natl Acad. Sci. USA 116, 18152–18156 (2019).Article 
CAS 

Google Scholar 
Béné, C. et al. Feeding 9 billion by 2050—putting fish back on the menu. Food Secur. 7, 261–274 (2015).Article 

Google Scholar 
Tacon, A. G. J. & Metian, M. Fish matters: importance of aquatic foods in human nutrition and global food supply. Rev. Fish. Sci. 21, 22–38 (2013).Article 
CAS 

Google Scholar 
Verones, F., Moran, D., Stadler, K., Kanemoto, K. & Wood, R. Resource footprints and their ecosystem consequences. Sci. Rep. 7, 40743 (2017).Article 
CAS 

Google Scholar 
Mekonnen, M. M. & Hoekstra, A. Y. The green, blue and grey water footprint of crops and derived crop products. Hydrol. Earth Syst. Sci. 15, 1577–1600 (2011).Article 

Google Scholar 
Mekonnen, M. M. & Hoekstra, A. Y. The Green, Blue and Grey Water Footprint of Farm Animals and Animal Products (UNESCO-IHE, 2010).Carlson, K. M. et al. Greenhouse gas emissions intensity of global croplands. Nat. Clim. Change 7, 63–68 (2017).Article 
CAS 

Google Scholar 
Hong, C. et al. Global and regional drivers of land-use emissions in 1961–2017. Nature 589, 554–561 (2021).Article 
CAS 

Google Scholar 
Amoroso, R. O. et al. Bottom trawl fishing footprints on the world’s continental shelves. Proc. Natl Acad. Sci. USA 115, E10275–E10282 (2018).Article 
CAS 

Google Scholar 
Kuempel, C. D. et al. Integrating life cycle and impact assessments to map food’s cumulative environmental footprint. One Earth 3, 65–78 (2020).Article 

Google Scholar 
Halpern, B. S. et al. A global map of human impact on marine ecosystems. Science 319, 948–952 (2008).Article 
CAS 

Google Scholar 
Crain, C. M., Kroeker, K. & Halpern, B. S. Interactive and cumulative effects of multiple human stressors in marine systems. Ecol. Lett. 11, 1304–1315 (2008).Article 

Google Scholar 
Birk, S. et al. Impacts of multiple stressors on freshwater biota across spatial scales and ecosystems. Nat. Ecol. Evol. 4, 1060–1068 (2020).Article 

Google Scholar 
Judd, A. D., Backhaus, T. & Goodsir, F. An effective set of principles for practical implementation of marine cumulative effects assessment. Environ. Sci. Policy 54, 254–262 (2015).Article 

Google Scholar 
IPBES Global Assessment Report on Biodiversity and Ecosystem Services of the Intergovernmental Science-Policy Platform on Biodiversity and Ecosystem Services (IPBES, 2019).Froehlich, H. E., Jacobsen, N. S., Essington, T. E., Clavelle, T. & Halpern, B. S. Avoiding the ecological limits of forage fish for fed aquaculture. Nat. Sustain. 1, 298–303 (2018).Article 

Google Scholar 
FAO The State of World Fisheries and Aquaculture 2020 (FAO, 2020).Froehlich, H. E., Runge, C. A., Gentry, R. R., Gaines, S. D. & Halpern, B. S. Comparative terrestrial feed and land use of an aquaculture-dominant world. Proc. Natl Acad. Sci. USA 115, 5295–5300 (2018).Article 
CAS 

Google Scholar 
FAOSTAT Database: New Food Balances (FAO, 2020); http://www.fao.org/faostat/en/#data/FBSFAOSTAT Database: Production, Crops (FAO, 2020); http://www.fao.org/faostat/en/#data/QCDong, F. et al. Assessing sustainability and improvements in US Midwestern soybean production systems using a PCA–DEA approach. Renew. Agric. Food Syst. 31, 524–539 (2016).Article 

Google Scholar 
Watson, R. A. & Tidd, A. Mapping nearly a century and a half of global marine fishing: 1869–2015. Mar. Policy 93, 171–177 (2018).Article 

Google Scholar 
Robinson, T. P. et al. Mapping the global distribution of livestock. PLoS ONE 9, e96084 (2014).Article 

Google Scholar 
Clark, M. & Tilman, D. Comparative analysis of environmental impacts of agricultural production systems, agricultural input efficiency, and food choice. Environ. Res. Lett. 12, 064016 (2017).Article 

Google Scholar 
Balmford, B., Green, R. E., Onial, M., Phalan, B. & Balmford, A. How imperfect can land sparing be before land sharing is more favourable for wild species? J. Appl. Ecol. 56, 73–84 (2019).Article 

Google Scholar 
Luskin, M. S., Lee, J. S. H., Edwards, D. P., Gibson, L. & Potts, M. D. Study context shapes recommendations of land-sparing and sharing; a quantitative review. Glob. Food Secur. 16, 29–35 (2018).Article 

Google Scholar 
Williams, D. R., Phalan, B., Feniuk, C., Green, R. E. & Balmford, A. Carbon storage and land-use strategies in agricultural landscapes across three continents. Curr. Biol. 28, 2500–2505.e4 (2018).Article 
CAS 

Google Scholar 
Paul, B. G. & Vogl, C. R. Impacts of shrimp farming in Bangladesh: challenges and alternatives. Ocean Coastal Manage. 54, 201–211 (2011).Article 

Google Scholar 
Ahmed, N., Cheung, W. W. L., Thompson, S. & Glaser, M. Solutions to blue carbon emissions: shrimp cultivation, mangrove deforestation and climate change in coastal Bangladesh. Mar. Policy 82, 68–75 (2017).Article 

Google Scholar 
FAOSTAT Database: Livestock Primary (FAO, 2020); http://www.fao.org/faostat/en/#data/QLRamankutty, N., Ricciardi, V., Mehrabi, Z. & Seufert, V. Trade-offs in the performance of alternative farming systems. Agric. Econ. 50, 97–105 (2019).Article 

Google Scholar 
FAOSTAT Database: Detailed Trade Matrix (FAO, 2020); http://www.fao.org/faostat/en/#data/TMFisheries & Aquaculture—Fishery Statistical Collections—Fishery Commodities and Trade (FAO, 2019); http://www.fao.org/fishery/statistics/global-commodities-production/enInternational Food Policy Research Institute. Global spatially-disaggregated crop production statistics data for 2010, version 2.0. Harvard Dataverse https://doi.org/10.7910/DVN/PRFF8V (2019).Clawson, G. et al. Mapping the spatial distribution of global mariculture production. Aquaculture 553, 738066 (2022).Article 

Google Scholar 
Petz, K. et al. Mapping and modelling trade-offs and synergies between grazing intensity and ecosystem services in rangelands using global-scale datasets and models. Glob. Environ. Change 29, 223–234 (2014).Article 

Google Scholar 
Global Fishing Watch. Fishing effort. Fleet daily, v2 100th degree. (2021). https://globalfishingwatch.org/dataset-and-code-fishing-effort/Verdegem, M. C. J., Bosma, R. H. & Verreth, J. A. J. Reducing water use for animal production through aquaculture. Int. J. Water Resour. Dev. 22, 101–113 (2006).Article 

Google Scholar 
Bouwman, A. F., Beusen, A. H. W. & Billen, G. Human alteration of the global nitrogen and phosphorus soil balances for the period 1970–2050. Glob. Biogeochem. Cycles 23, GB0A04 (2009).Article 

Google Scholar 
Bouwman, A. F., Van Drecht, G. & Van der Hoek, K. W. Nitrogen surface balances in intensive agricultural production systems in different world regions for the period 1970–2030. Pedosphere 15, 137–155 (2005).
Google Scholar 
Bouwman, A., Boumans, L. J. M. & Batjes, N. Estimation of global NH3 volatilization loss from synthetic fertilizers and animal manure applied to arable lands and grasslands. Glob. Biogeochem. Cycles 16, 8-1–8-14 (2002).Article 

Google Scholar 
FAOSTAT Database: Inputs, Fertilizers by Nutrient (FAO, 2020); http://www.fao.org/faostat/en/#data/RFNHeffer, P., Gruere, A. & Roberts, T. Assessment of fertilizer use by crop at the global level 2014–2014/15, International Fertilizer Association (2017).Fertilizer Use by Crop 5th edn (FAO, IFA & IFDC, 2002).Islam, Md. S. Nitrogen and phosphorus budget in coastal and marine cage aquaculture and impacts of effluent loading on ecosystem: review and analysis towards model development. Mar. Pollut. Bull. 50, 48–61 (2005).Article 
CAS 

Google Scholar 
Wang, J., Beusen, A. H. W., Liu, X. & Bouwman, A. F. Aquaculture production is a large, spatially concentrated source of nutrients in Chinese freshwater and coastal seas. Environ. Sci. Technol. 54, 1464–1474 (2020).Article 

Google Scholar 
Bouwman, A. F. et al. Hindcasts and future projections of global inland and coastal nitrogen and phosphorus loads due to finfish aquaculture. Rev. Fish. Sci. 21, 112–156 (2013).Article 
CAS 

Google Scholar 
Gavrilova, O. et al. in 2019 Refinement to the 2006 IPCC Guidelines for National Greenhouse Gas Inventories Ch. 10, Intergovernmental Panel on Climate Change (IPCC); Review Editors on Overview: Dario Gómez (Argentina) and William Irving (USA) (2019).Seafood Carbon Emissions Tool, Lisa Max, Robert Parker, Peter Tyedmers, editors; (2020); http://seafoodco2.dal.ca/Hu, Z., Lee, J. W., Chandran, K., Kim, S. & Khanal, S. K. Nitrous oxide (N2O) emission from aquaculture: a review. Environ. Sci. Technol. 46, 6470–6480 (2012).Article 
CAS 

Google Scholar 
IPCC Climate Change 2007: The Physical Science Basis (eds Solomon, S. et al.) (Cambridge Univ. Press, 2007).Lynch, J., Cain, M., Pierrehumbert, R. & Allen, M. Demonstrating GWP*: a means of reporting warming-equivalent emissions that captures the contrasting impacts of short- and long-lived climate pollutants. Environ. Res. Lett. 15, 044023 (2020).Article 
CAS 

Google Scholar 
Global Livestock Environmental Assessment Model, GLEAM, v.2.0.121 (FAO, 2018).Aas, T. S., Ytrestøyl, T. & Åsgård, T. Utilization of feed resources in the production of Atlantic salmon (Salmo salar) in Norway: an update for 2016. Aquacult. Rep. 15, 100216 (2019).
Google Scholar 
Jackson, A. Fish in-fish out (FIFO) explained. Aquacult. Eur. 34, 5–10 (2009).
Google Scholar 
Halpern, B. S. et al. Spatial and temporal changes in cumulative human impacts on the world’s ocean. Nat. Commun. 6, 7615 (2015).Article 
CAS 

Google Scholar 
Frazier, M. et al. Global food system pressure data. https://knb.ecoinformatics.org/view/doi:10.5063/F1V69H1B More

Housley, R. A., Gamble, C. S., Street, M. & Pettitt, P. Radiocarbon evidence for the lateglacial human recolonisation of Northern Europe. Proc. Prehist. Soc. 63, 25–54 (1997).
Google Scholar 
Blockley, S. P. E., Donahue, R. E. & Pollard, A. M. Radiocarbon calibration and Late Glacial occupation in northwest Europe. Antiquity 74, 112–119 (2000).
Google Scholar 
Terberger, T., Barton, N. & Street, M. in Humans, Environment and Chronology of the Late Glacial of the North European Plain (eds Street, M. et al.) 189–207 (Romisch-Germanisches Zentralmuseum, 2009).Miller, R. Mapping the expansion of the Northwest Magdalenian. Quat. Int. 272–273, 209–230 (2012).
Google Scholar 
Riede, F. & Pedersen, J. B. Late Glacial human dispersals in Northern Europe and disequilibrium dynamics. Hum. Ecol. 46, 621–632 (2018).
Google Scholar 
Lazaridis, I. et al. Ancient human genomes suggest three ancestral populations for present-day Europeans. Nature 513, 409–413 (2014).CAS 
PubMed 
PubMed Central 

Google Scholar 
Jones, E. R. et al. Upper Palaeolithic genomes reveal deep roots of modern Eurasians. Nat. Commun. 6, 8912 (2015).CAS 
PubMed 

Google Scholar 
Fu, Q. et al. The genetic history of Ice Age Europe. Nature 534, 200–205 (2016).CAS 
PubMed 
PubMed Central 

Google Scholar 
Villalba-Mouco, V. et al. Survival of Late Pleistocene hunter-gatherer ancestry in the Iberian Peninsula. Curr. Biol. 29, 1169–1177 (2019).Willis, K. J. & Whittaker, R. J. Perspectives: paleoecology. The refugial debate. Science 287, 1406–1407 (2000).CAS 
PubMed 

Google Scholar 
Sommer, R. S. & Nadachowski, A. Glacial refugia of mammals in Europe: evidence from fossil records. Mamm. Rev. 36, 251–265 (2006).
Google Scholar 
Bennett, K. D. & Provan, J. What do we mean by ‘refugia’? Quat. Sci. Rev. 27, 2449–2455 (2008).
Google Scholar 
Terberger, T. & Street, M. Hiatus or continuity? New results for the question of pleniglacial settlement in Central Europe. Antiquity 76, 691–698 (2002).
Google Scholar 
Maier, A. in The Central European Magdalenian. Vertebrate Paleobiology and Paleoanthropology (ed. Maier, A.) 231–241 (Springer, 2015).Reade, H. et al. Radiocarbon chronology and environmental context of Last Glacial Maximum human occupation in Switzerland. Sci. Rep. 10, 4694 (2020).CAS 
PubMed 
PubMed Central 

Google Scholar 
Stevens, R. E., Hermoso-Buxán, X. L., Marín-Arroyo, A. B., González-Morales, M. R. & Straus, L. G. Investigation of Late Pleistocene and Early Holocene palaeoenvironmental change at El Mirón cave (Cantabria, Spain): insights from carbon and nitrogen isotope analyses of red deer. Palaeogeogr. Palaeoclimatol. Palaeoecol. 414, 46–60 (2014).
Google Scholar 
Clark, C. D., Hughes, A. L. C., Greenwood, S. L., Jordan, C. & Sejrup, H. P. Pattern and timing of retreat of the last British–Irish Ice Sheet. Quat. Sci. Rev. 44, 112–146 (2012).
Google Scholar 
Currant, A. P. & Jacobi, R. in The Ancient Human Occupation of Britain Vol. 14 (eds Ashton, N. et al.) 165–180 (Elsevier, 2011).Walker, M. J. C. et al. Devensian lateglacial environmental changes in Britain: a multi-proxy environmental record from Llanilid, South Wales, UK. Quat. Sci. Rev. 22, 475–520 (2003).
Google Scholar 
Hill, T. C. B. et al. Devensian late-glacial environmental change in the Gordano Valley, North Somerset, England: a rare archive for southwest Britain. J. Paleolimnol. 40, 431–444 (2008).
Google Scholar 
Jacobi, R. M. & Higham, T. F. G. The early Lateglacial re-colonization of Britain: new radiocarbon evidence from Gough’s Cave, southwest England. Quat. Sci. Rev. 28, 1895–1913 (2009).
Google Scholar 
Jacobi, R. & Higham, T. in The Ancient Human Occupation of Britain Vol. 14 (eds Ashton, N. M. et al.) 223–247 (Elsevier, 2011).Grimm, S. B. & Weber, M.-J. The chronological framework of the Hamburgian in the light of old and new 14C dates. Quartär. 55, 17–40 (2008).
Google Scholar 
Olalde, I. et al. The Beaker phenomenon and the genomic transformation of northwest Europe. Nature 555, 190–196 (2018).CAS 
PubMed 
PubMed Central 

Google Scholar 
Brace, S. et al. Ancient genomes indicate population replacement in Early Neolithic Britain. Nat. Ecol. Evol. 3, 765–771 (2019).PubMed 
PubMed Central 

Google Scholar 
Jacobi, R. M. & Higham, T. F. G. The ‘Red Lady’ ages gracefully: new ultrafiltration AMS determinations from Paviland. J. Hum. Evol. 55, 898–907 (2008).CAS 
PubMed 

Google Scholar 
Schulting, R. J. et al. A mid-upper Palaeolithic human humerus from Eel Point, South Wales, UK. J. Hum. Evol. 48, 493–505 (2005).PubMed 

Google Scholar 
Richards, M. P., Hedges, R. E. M., Jacobi, R., Current, A. & Stringer, C. FOCUS: Gough’s Cave and Sun Hole Cave human stable isotope values indicate a high animal protein diet in the British Upper Palaeolithic. J. Archaeol. Sci. 27, 1–3 (2000).
Google Scholar 
Proctor, C., Douka, K., Proctor, J. W. & Higham, T. The age and context of the KC4 Maxilla, Kent’s Cavern, UK. Eur. J. Archaeol. 20, 74–97 (2017).
Google Scholar 
Richards, M. P., Jacobi, R., Cook, J., Pettitt, P. B. & Stringer, C. B. Isotope evidence for the intensive use of marine foods by Late Upper Palaeolithic humans. J. Hum. Evol. 49, 390–394 (2005).CAS 
PubMed 

Google Scholar 
Bello, S. M., Saladié, P., Cáceres, I., Rodríguez-Hidalgo, A. & Parfitt, S. A. Upper Palaeolithic ritualistic cannibalism at Gough’s Cave (Somerset, UK): the human remains from head to toe. J. Hum. Evol. 82, 170–189 (2015).PubMed 

Google Scholar 
Andrews, P. & Fernández-Jalvo, Y. Cannibalism in Britain: taphonomy of the Creswellian (Pleistocene) faunal and human remains from Gough’s Cave (Somerset, England). Bull. Nat. Hist. Mus. Geol. 58, 59–81 (2003).
Google Scholar 
Bello, S. M., Parfitt, S. A. & Stringer, C. B. Earliest directly-dated human skull-cups. PLoS ONE 6, e17026 (2011).CAS 
PubMed 
PubMed Central 

Google Scholar 
Currant, A. P., Jacobi, R. M. & Stringer, C. B. Excavations at Gough’s Cave, Somerset 1986–7. Antiquity 63, 131–136 (1989).
Google Scholar 
Davies, M. in Limestones and Caves of Wales (ed. Ford, T. D.) 92–101 (Cambridge Univ. Press, 1989).Dawkins, W. B. Memorandum on the remains from the cave at the Great Ormes Head. Proc. Liverp. Geol. Soc. 4, 156–159 (1880).
Google Scholar 
Sieveking, G. & de, G. The Kendrick’s Cave mandible. Br. Mus. Q. 35, 230–250 (1971).
Google Scholar 
Pettitt, P. B. Discovery, nature and preliminary thoughts about Britain’s first cave art.Capra 5,1–12 (2003).
Google Scholar 
Bello, S. M., Wallduck, R., Parfitt, S. A. & Stringer, C. B. An Upper Palaeolithic engraved human bone associated with ritualistic cannibalism. PLoS ONE 12, e0182127 (2017).PubMed 
PubMed Central 

Google Scholar 
Bocherens, H. & Drucker, D. Isotope evidence for paleodiet of late Upper Paleolithic humans in Great Britain: a response to Richards et al. 2005. J. Hum. Evol. 51, 440–442 (2006).PubMed 

Google Scholar 
Fernandes, R., Millard, A. R., Brabec, M., Nadeau, M.-J. & Grootes, P. Food reconstruction using isotopic transferred signals (FRUITS): a Bayesian model for diet reconstruction. PLoS ONE 9, e87436 (2014).PubMed 
PubMed Central 

Google Scholar 
Rasmussen, S. O. et al. A stratigraphic framework for abrupt climatic changes during the Last Glacial period based on three synchronized Greenland ice-core records: refining and extending the INTIMATE event stratigraphy. Quat. Sci. Rev. 106, 14–28 (2014).
Google Scholar 
Kloss-Brandstätter, A. et al. HaploGrep: a fast and reliable algorithm for automatic classification of mitochondrial DNA haplogroups. Hum. Mutat. 32, 25–32 (2011).PubMed 

Google Scholar 
Skoglund, P., Storå, J., Götherström, A. & Jakobsson, M. Accurate sex identification of ancient human remains using DNA shotgun sequencing. J. Archaeol. Sci. 40, 4477–4482 (2013).CAS 

Google Scholar 
Haak, W. et al. Massive migration from the steppe was a source for Indo-European languages in Europe. Nature 522, 207–211 (2015).CAS 
PubMed 
PubMed Central 

Google Scholar 
Fu, Q. et al. An early modern human from Romania with a recent Neanderthal ancestor. Nature 524, 216–219 (2015).CAS 
PubMed 
PubMed Central 

Google Scholar 
Patterson, N., Price, A. L. & Reich, D. Population structure and eigenanalysis. PLoS Genet. 2, e190 (2006).PubMed 
PubMed Central 

Google Scholar 
Price, A. L. et al. Principal components analysis corrects for stratification in genome-wide association studies. Nat. Genet. 38, 904–909 (2006).CAS 
PubMed 

Google Scholar 
Mallick, S. et al. The Simons Genome Diversity Project: 300 genomes from 142 diverse populations. Nature 538, 201–206 (2016).CAS 
PubMed 
PubMed Central 

Google Scholar 
Patterson, N. et al. Ancient admixture in human history. Genetics 192, 1065–1093 (2012).PubMed 
PubMed Central 

Google Scholar 
Harney, É., Patterson, N., Reich, D. & Wakeley, J. Assessing the performance of qpAdm: a statistical tool for studying population admixture. Genetics 217, iyaa045 (2021).PubMed 
PubMed Central 

Google Scholar 
Currant, A. & Jacobi, R. A formal mammalian biostratigraphy for the Late Pleistocene of Britain. Quat. Sci. Rev. 20, 1707–1716 (2001).
Google Scholar 
Pickard, C. & Bonsall, C. Post-glacial hunter-gatherer subsistence patterns in Britain: dietary reconstruction using FRUITS. Archaeol. Anthropol. Sci. 12, 142 (2020).
Google Scholar 
Stevens, R. E., Jacobi, R. M. & Higham, T. F. G. Reassessing the diet of Upper Palaeolithic humans from Gough’s Cave and Sun Hole, Cheddar Gorge, Somerset, UK. J. Archaeol. Sci. 37, 52–61 (2010).
Google Scholar 
Sala, N. & Conard, N. Taphonomic analysis of the hominin remains from Swabian Jura and their implications for the mortuary practices during the Upper Paleolithic. Quat. Sci. Rev. 150, 278–300 (2016).
Google Scholar 
Saladié, P. & Rodríguez-Hidalgo, A. Archaeological evidence for cannibalism in prehistoric Western Europe: from Homo antecessor to the Bronze Age. J. Archaeol. Method Theory 24, 1034–1071 (2017).
Google Scholar 
Cook, J. Ice Age Art: Arrival of the Modern Mind (British Museum Press, 2013).Gupta, S., Collier, J. S., Palmer-Felgate, A. & Potter, G. Catastrophic flooding origin of shelf valley systems in the English Channel. Nature 448, 342–345 (2007).CAS 
PubMed 

Google Scholar 
Mills, W. in From the Atlantic to Beyond the Bug River. Finding and Defining the Federmesser-Gruppen/Azilian (eds Grimm, S. B. et al.) 1–24 (Propylaeum, 2020).Amkreutz, L. et al. What lies beneath … Late Glacial human occupation of the submerged North Sea landscape. Antiquity 92, 22–37 (2018).
Google Scholar 
Ward, I., Larcombe, P. & Lillie, M. The dating of Doggerland—post-glacial geochronology of the southern North Sea. Environ. Archaeol. 11, 207–218 (2006).
Google Scholar 
Brock, F., Higham, T., Ditchfield, P. & Ramsey, C. B. Current pretreatment methods for AMS radiocarbon dating at the Oxford Radiocarbon Accelerator Unit (ORAU). Radiocarbon 52, 103–112 (2010).CAS 

Google Scholar 
Dabney, J. et al. Complete mitochondrial genome sequence of a Middle Pleistocene cave bear reconstructed from ultrashort DNA fragments. Proc. Natl Acad. Sci. USA 110, 15758–15763 (2013).CAS 
PubMed 
PubMed Central 

Google Scholar 
Meyer, M. & Kircher, M. Illumina sequencing library preparation for highly multiplexed target capture and sequencing. Cold Spring Harb. Protoc. 2010, pdb.prot5448 (2010).Rohland, N., Harney, E., Mallick, S., Nordenfelt, S. & Reich, D. Partial uracil–DNA–glycosylase treatment for screening of ancient DNA. Philos. Trans. R. Soc. Lond. B 370, 20130624 (2015).
Google Scholar 
Kircher, M., Sawyer, S. & Meyer, M. Double indexing overcomes inaccuracies in multiplex sequencing on the Illumina platform. Nucleic Acids Res. 40, e3 (2012).CAS 
PubMed 

Google Scholar 
Briggs, A. W. et al. Patterns of damage in genomic DNA sequences from a Neandertal. Proc. Natl Acad. Sci. USA 104, 14616–14621 (2007).CAS 
PubMed 
PubMed Central 

Google Scholar 
Skoglund, P. et al. Separating endogenous ancient DNA from modern day contamination in a Siberian Neandertal. Proc. Natl Acad. Sci. USA 111, 2229–2234 (2014).CAS 
PubMed 
PubMed Central 

Google Scholar 
Quinlan, A. R. & Hall, I. M. BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–842 (2010).CAS 
PubMed 
PubMed Central 

Google Scholar 
Petr, M., Vernot, B. & Kelso, J. admixr—R package for reproducible analyses using ADMIXTOOLS. Bioinformatics 35, 3194–3195 (2019).CAS 
PubMed 
PubMed Central 

Google Scholar 
Busing, F. M., Meijer, E. & Van Der Leeden, R. Delete-m jackknife for unequal m. Stat. Comput. 9, 3–8 (1999).
Google Scholar 
Fu, Q. et al. Genome sequence of a 45,000-year-old modern human from western Siberia. Nature 514, 445–449 (2014).CAS 
PubMed 
PubMed Central 

Google Scholar 
Raghavan, M. et al. Upper Palaeolithic Siberian genome reveals dual ancestry of Native Americans. Nature 505, 87–91 (2014).PubMed 

Google Scholar 
Lazaridis, I. et al. Genomic insights into the origin of farming in the ancient Near East. Nature 536, 419–424 (2016).CAS 
PubMed 
PubMed Central 

Google Scholar 
Lipson, M. et al. Parallel palaeogenomic transects reveal complex genetic history of early European farmers. Nature 551, 368–372 (2017).CAS 
PubMed 
PubMed Central 

Google Scholar 
Gallego Llorente, M. et al. Ancient Ethiopian genome reveals extensive Eurasian admixture throughout the African continent. Science 350, 820–822 (2015).CAS 
PubMed 

Google Scholar 
Villalba-Mouco, V. et al. Survival of Late Pleistocene hunter-gatherer ancestry in the Iberian Peninsula. Curr. Biol. 29, 1169–1177 (2019).CAS 
PubMed 

Google Scholar  More

Animal experimentsTeratoma formation experiments were performed at Iwate University. All surgical procedures and animal husbandry were performed in accordance with the international guidelines of the Animal Experiments of Iwate University and were approved by the university’s Animal Research Committee (approval number A201734).Chicken embryonic fibroblasts (Rhode Island Red) were obtained from a primary culture of chicken embryonic tissue provided by Prof. Atsushi Tajima, Tsukuba University. Chicken culture cells were obtained from chicken embryos, and the acquisition of these cells did not require approval. Mouse embryonic fibroblasts (CF-1 strain) were purchased from a manufacturer (CMPMEFCFL; DS Pharma Biomedical, Osaka, Japan). Approval was not required to obtain these cells.Somatic cells were obtained from wild animals (ex., Okinawa rail). The sampling details described below do not include the exact location of sampling to protect against poaching.Fibroblast cells from Okinawa rail and Japanese ptarmigan were obtained from dead animals, such as those killed by vehicles (Fig. 1A and Supplementary Fig. 1). Approval was not required to obtain these samples.Dead Okinawa rail were found on May 21, 2008, by the Okinawa Wildlife Federation, a nonprofit organization that focuses on the conservation of wild animals in the Okinawa area in the southwest region of Japan. The organization has permission from the Japanese Ministry of the Environment (MOE) to handle and perform first aid activities on endangered animals. The dead birds were transferred the following day to the National Institute for Environmental Studies (NIES). Primary cell culture was carried out from muscle tissue and skin of the dead birds (NIES ID: 715A).On July 8, 2004, tissues recovered from dead Japanese ptarmigan (e.g., skin and retina tissues) were also transferred to NIES from Gifu University Department of Veterinary Medicine. Primary cell culture from this tissue was performed (NIES ID: 22A).Somatic cells from Blakiston’s fish owl and Japanese golden eagle were obtained from emerging pinfeathers. Concerning the Blakiston’s fish owl, the MOE carries out bird banding, of wild birds with identification tags. The emerging pinfeathers we used had been accidentally release during banding. The banding had been performed by a veterinarian at the Institute for Raptor Biomedicine Japan (IRBJ) in the Hokkaido area on June 2, 2006. IRBJ is a private organization that primarily focuses on emergency medicine first aid and care for wild avians in Hokkaido region of Japan. IRBJ is contracted to MOE to handle and administer first aid for endangered animals. The MOE banding ring was 14C0242. Since banding was carried out with the permission of MOE for capturing wildlife, we did not require the approval to obtain these avian somatic cells. On July 8, 2006, Blakiston’s fish owl pinfeathers were transferred to from IRBJ to NIES, where primary cell culture was performed (NIES ID: 215A).Concerning the Japanese golden eagle, an emerging pinfeather accidentally fell off a bird during blood collection at the Yagiyama Zoo in Sendai, Japan on July 11, 2018. Dr. Yukiko Watanabe, an IRBJ veterinarian, collected the emerging pinfeather. The sample was shipped the following day to NIES where primary cell culture was performed (NIES ID: 5228).In addition to these birds, we obtained somatic cells emerging avian pinfeathers of Steller’s sea eagle, white-tail eagle, mountain hawk-eagle, northern goshawk, Taiga bean goose, and Latham’s snipe. These samples were provided by IRBJ.Concerning the Steller’s sea eagle, an injured individual was found in Hokkaido on July 11, 2006 (ID: 06-NE-SSE-1). The eagle was transferred to IRBJ. On December 4, 2006, IRBJ veterinarian Dr. Keisuke Saito collected fallen pinfeathers. Primary cell culture was performed at NIES on December 8, 2006 (NIES ID: 369A).Concerning the white-tailed eagle, an injured individual was found in Hokkaido, Japan, on July 12, 2007 (ID: 07-NE-WTE-4). The bird was transferred to IRBJ the same day for emergency treatment. On January 15, 2008, Dr. Saito collected fallen pinfeathers. Primary cell culture was performed on January 18, 2008 at NIES (NIES ID: 492A).Concerning the mountain hawk-eagle, an injured individual was found in the Hokkaido area on August 10, 2008 (ID: 08-Tokachi-HHE-2). The bird was transferred to IRBJ the same day. The bird was treated by an IRBJ veterinarian, but died on September 8, 2008. Emerging pinfeathers were collected from the dead bird by Dr. Saito. Primary cell culture was performed on September 11, 2008 at NIES (NIES ID: 847A).Concerning the Northern Goshawk, IRBJ accepted an injured bird for treatment on June 12, 2006. Following treatment and recovery, the bird was released into the wild in the Hokkaido area on August 1, 2006. During the treatment (July 4, 2006), Dr. Saito collected fallen pinfeathers. The primary cell culture was performed at NIES on July 6, 2006 (NIES ID: 222A).Concerning the Taiga bean geese, an injured individual was found in Hokkaido on September 15, 2016 (ID: 13B8005). The injured bird was transferred to IRBJ the same day for emergency treatment. On September 16, 2016, IRBJ veterinarian Dr. Yukiko Watanabe collected fallen emerging pinfeathers. Primary cell culture was performed on September 20, 2016 (NIES ID: 4420A).Finally, concerning the Latham’s snipe, fallen pinfeathers were collected during MOE approved bird banding performed on September 17, 2006, by Dr. Saito. Dr. Saito also collected fallen emerging pinfeathers (ID: 6A22598). The samples were transferred to NIES on September 20, 2006, for primary cell culture (NIES ID: 338A).All records are available at NIES.Cell culture and preservationOkinawa rail, Japanese ptarmigan, and Blakiston’s fish owl-derived fibroblasts were preserved in liquid nitrogen for 8–12 years (Fig. 1f). The preservation solution contained 90% fetal bovine serum (FBS) and 10% dimethyl sulfoxide. Cells were preserved at a cell density of 1 × 106–4 × 106 cell/mL. During the freezing period, the cells were maintained at minus The cells were frozen at a temperature of −135 °C. Japanese golden eagle fibroblasts were used without freezing.Avian-derived fibroblasts were cultured with Kuwana’s modified avian culture medium-1 (KAv-1), which is based on alpha-MEM containing 5% FBS and 5% chicken serum23. Mouse embryonic fibroblasts were cultured in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% FBS and 1% antibiotic–antimycotic mixed stock solution (161–23181; Wako Pure Chemical Industries, Osaka, Japan). All avian and mouse cells were cultured at 37 °C under 5% CO2.Reprogramming vectorWe chemically synthesized an expression cassette that included seven reprogramming factors (MyoD-derived transactivation domain-linked Oct3/4, Sox2, Klf4, c-Myc, Klf2, Lin28, and Nanog; all genes derived from mice). The self-cleaving 2A peptide was inserted at the junction of the coding region (Fig. 1g). We transferred the complementary DNA (cDNA) insert from the shuttle vector to the PiggyBac transposon vector containing green fluorescent protein (PB-CAG-GFP). Although the original transposon vector drive the expression of cDNA with the elongation factor-1 (EF1) promoter (PJ547-17; DNA 2.0, Menlo Park, CA, USA), we replaced the EF1 promoter to CAG promoter in our previous study22,24. The reprogramming vector was designated PB-TAD-7F (Fig. 1g).In addition to the PB-TAD-7F reprogramming vector, we used the PB-DDR-8F reprogramming vector to establish Japanese golden eagle iPSCs. The complete coding sequence of DDR-8F (DDR-Oct3/4, Sox2, Klf4, c-Myc, Klf2, Nanog, Lin28, and Yap) was chemically synthesized. The expression cassettes containing the eight reprogramming factors were excised from the shuttle vector using restriction enzymes. The cDNA fragments were transferred to the PB-CAG-GFP PiggyBac transposon-based vector22,24. Detailed information regarding the PB-DDR-8F reprogramming vectors is shown in Fig. 10a.Establishment of iPSCsWe transfected PB-R6F or PB-TAD-7F reprogramming vectors into mouse, chicken, Okinawa rail, Japanese ptarmigan, and Blakiston’s fish owl-derived fibroblasts using Lipofectamine 2000 transfection reagent (Thermo Fisher Scientific, Waltham, MA, USA). After hygromycin selection (Wako Pure Chemical Industries), the cells were reseeded onto a mouse embryonic fibroblast (MEF) feeder layer. On days 14–32, we picked primary iPSC-like colonies and seeded them on new MEF feeder cell plates. The detailed protocol is shown in Fig. 1h.To establish Japanese golden eagle-derived iPSCs, we transduced PB-TAD-7F or PB-DDR-8F reprogramming vectors into Japanese golden eagle pinfeather-derived somatic cells. Transfection was performed using Lipofectamine 2000 transduction reagent (11668019; Thermo Fisher Scientific) according to the manufacturer’s instructions. After hygromycin selection (Wako Pure Chemical Industries), cells were seeded onto feeder culture plates. The golden eagle iPSCs were cultured in KAv-1-based medium5.The medium used to establish avian iPSCs was supplemented with 1000 × human Leukemia Inhibitory Factor (LIF) (125–05603; Wako Pure Chemical Industries), 4.0 ng/ml basic FGF (064–04541; Wako Pure Chemical Industries), 0.75 μM CHIR99021 glycogen synthase kinase-3 inhibitor (034–23103; Wako Pure Chemical Industries), 0.25 μM PD0325901 mitogen-activated protein kinase inhibitor (163–24001; Wako Pure Chemical Industries). In addition to those supplements, 0.25 μM thiazovivin (202–18011; Wako Pure Chemical Industries) was added in the media used to generate Okinawa rail, Japanese ptarmigan, Blakiston’s fish owl, and chicken iPSCs. In the medium used to generate mouse iPSCs, we added 1000 × LIF, 0.75 μM CHIR99021, and 0.25 μM PD0325901.iPSC culture conditionsTwo types of cell culture media were used: KAv-1 for avian iPSCs and DMEM for mouse iPSCs. The composition of KAv-1 for avian iPSCs was as follows: alpha-MEM containing 5% FBS and 5% chicken serum 1% antibiotic–antimycotic mixed solution, 1% nonessential amino acids (Wako Pure Chemical Industries), and 2 mM glutamic acid was added (Nacalai Tesque, Kyoto, Japan). The composition of DMEM for mouse was follows: DMEM supplemented with 15% SSR, 0.22 mM 2-mercaptoethanol (21438–82, Nacalai Tesque), 1% antibiotic–antimycotic mixed solution, 1% nonessential amino acids5,22. As a supplement to the iPSC medium, we used 1000 × human LIF (125–05603; Wako Pure Chemical Industries), 4.0 ng/ml basic FGF (064–04541; Wako Pure Chemical Industries), 0.75 μM CHIR99021 (034–23103; Wako Pure Chemical Industries), 0.25 μM PD0325901 (163–24001; Wako Pure Chemical Industries) for the media used to culture Okinawa rail, Japanese ptarmigan, Blakiston’s fish owl, Japanese golden eagle, and chicken-derived iPSCs. The supplements for media used to culture Okinawa rail and Japanese ptarmigan-derived iPSCs included 2.5 μM Gö6983 (074–06443, Wako Pure Chemical Industries). To analyze the cellular characteristics, we focused on the Janus kinase (JAK), FGF, ROCK, and glycolytic pathways, since the dependency of these pathways can indicate differences in cellular characteristics. We used 1–10 μM JAK inhibitor I (4200099; MERCK, Darmstadt, Germany), 0.5–4 μM of PD173074, which inhibits FGF receptor (FGFR) inhibitor (160–26831; Wako Pure Chemical Industries), 10 μM of Y27632, which inhibits ROCK (036–24023; Wako Pure Chemical Industries), and 2 or 4 mM 2-deoxyglucose (2DG, D0051; Tokyo Chemical Industry, Tokyo, Japan).AP and immunological staining of fibroblasts and iPSCsA red-color AP staining kit (AP100 R-1; System Bioscience, Palo Alto, CA, USA) was used to detect AP activity of iPSCs. iPSCs were stained for SSEA-1, SSEA-3, and SSEA-4 antibodies (Supplementary Table 2). To stain the iPSCs with the SSEA antibodies, the cells were fixed in 4% paraformaldehyde in phosphate buffered saline (PBS) for 3 min. Cells were permeabilized by 0.5% Triton X-100 (35501-15; Nacalai Tesque, Kyoto, Japan) for 60 min. After three washes with PBS, the iPSCs were blocked with 1% bovine serum albumin (BSA, 01863-06; Nacalai Tesque) for 45 min. iPSCs were incubated with a primary antibody overnight and then exposed to the corresponding fluorescent-labeled secondary antibodies for 60 min. Counterstaining was performed with a 4′,6-diamidino-2-phenylindole (DAPI) solution (Cellstain-DAPI solution, DOJINDO, Kumamoto, Japan).Japanese golden eagle and chicken-derived fibroblasts were seeded in 12-well cell culture plates for immunological staining. After 48 h of incubation, F-actin staining was performed using Alexa Fluor 568 phalloidin (A12380; Thermo Fisher Scientific) according to the manufacturer’s protocol. Double staining was performed with an anti-vimentin antibody (MA5-11883; Thermo Fisher Scientific) and Alexa Fluor 488-labeled secondary antibody (A-11001; Thermo Fisher Scientific) (Supplementary Table 2). The samples were counterstained with Cellstain-DAPI solution (DOJINDO) as described above.Detection of reprogramming vectors and internal control genes from iPSCsDNA was isolated using the EZ1 DNA Tissue Kit (953034; QIAGEN, Hilden, Germany). PCR was performed with 100 ng of template DNA. Primer sequences are listed in Supplementary Tables 3 and 4. We performed PCR assays using KOD FX Neo (KFX-201; TOYOBO, Osaka, Japan). PCR was conducted by predenaturation at 94 °C for 2 min, denaturation at 98 °C for 10 s, and extension at 68 °C for 30 s, with 40 cycles of denaturation and extension. PCR products were analyzed by electrophoresis on 2.0% agarose/Tris-acetate–ethylenediaminetetraacetic acid (EDTA) gels.Sequential passagingMouse, Okinawa rail, and Japanese ptarmigan-derived primary cells and iPSCs were seeded in six-well plates with feeder cells for analysis. When cell growth became confluent, all cells and the number of cells per dish was enumerated using a Countess cell counter (Thermo Fisher Scientific). The harvested and seeded cell numbers were used to calculate the PD time as an indicator of the speed of cell growth, using the formula PD = log2 (A/B), where A is the number of harvested cells at the end of each passage, and B is the number of seeded cells at the start25.Detection of mRNA expressionTotal RNA was isolated from iPSCs using an EZ1 RNA Tissue Mini Kit (959034; QIAGEN). cDNA was synthesized from total RNA using the PrimeScript reverse transcription (RT) reagent kit (Perfect Real Time, RR047A; TaKaRa Bio, Ohtsu, Japan). Real-time PCR was performed in a 12.5 μl volume containing 2 × KOD SYBR qPCR Mix (QKD-201; Toyobo), 10 ng of cDNA solution, and 0.3 μM of each primer. The primer sequences are listed in Supplementary Tables 5–10. The reaction was performed in duplicate. The cycling program was as follows: 98 °C for 120 s (initial denaturation), 98 °C for 10 s (denaturation), 58 °C for 10 s (annealing), and 68 °C for 32 s (extension) for 40 cycles. We normalized the expression levels of the target genes to that of glyceraldehyde-3-phosphate dehydrogenase (GAPDH).Mitochondria stainingMitochondria were stained by incubation with 50 nM MitoTracker Orange (M7510; Thermo Fisher Scientific) or 20 nM tetramethyl rhodamine ethyl ester perchlorate (TMRE, T669; Thermo Fisher Scientific) for 10 min. After staining, the solution was removed, and fresh medium was added for observation.EB formation and in vitro differentiationIn vitro differentiation of Okinawa rail, Japanese ptarmigan, Blakiston’s fish owl, and Japanese golden eagle iPSCs was performed. To generate EBs, iPSCs were seeded in low-binding dishes in KAv-1 medium. After 7–14 days, floating EBs were selected and seeded in 0.1% gelatin-coated 6-well plates with KAv-1 medium. To induce differentiation into neural cells, the floating EBs were cultured in 0.1% gelatin-coated plates containing KAv-1 supplemented with 10 μM ATRA and 4.0 ng/ml FGF for 7 days.Cells were immunochemically stained after in vitro differentiation using antibody to TUJ1, alpha-smooth muscle, or Gata4 (Supplementary Table 2). Differentiated cells were stained based on the immunological staining procedure of iPSCs described above.Teratoma formation and tissue sectioningThe Animal Committee of Iwate University approved the experimental protocol for teratoma formation (approval numbers A201734, A201737). For teratoma formation, 1 × 106 iPSCs were injected into the testes of SCID mice (C.B-17/Icr-scid/scidJcl; CLEA Japan, Tokyo, Japan). After 4–34 weeks post-injection, tumor tissues were excised from the mice. Each tumor tissue was fixed with 10% formaldehyde in PBS. Fixed tissue sections were stained with hematoxylin-eosin (HE) and observed by microscopy.Immunological staining was performed in addition to HE staining. For immunological staining, antibody to TUJ1, alpha-smooth muscle, or Gata4 was used (Supplementary Table 2). The paraffin block of each teratoma was sliced to produce a section 5 μm thick. After deparaffinization, the antigen was activated with citric acid buffer (SignalStain Citrate Unmasking Solution (10×), 14746; Cell Signaling Technology, Beverly, MA, USA) by microwaving for 10 min. To block endogenous peroxidase, tissue sections were incubated with 3% hydrogen peroxide (081–04215; Wako Pure Chemical). After washing with purified water, the tissue sections were incubated with 5% goat serum (555–76251; Wako Pure Chemical) in PBS. Next, the section were incubated in a solution containing a 1:100 dilution of primary antibody overnight at 4 °C. After washing with PBS, the tissue sections were incubated with horseradish peroxidase (HRP) conjugated secondary antibody (anti-IgG (H+L chain), mouse, pAb-HRP, code no. 330; MBL Co., Ltd., Nagoya, Japan) or anti-IgG (H+L chain, rabbit, pAb-HRP, code no. 458; MBL) for 1 h (Supplementary Table 2). After washing with PBS, the tissue sections were incubated with 3,3′-diaminobenzidine substrate solution (Histostar, code no. 8469; MBL) for 5–20 min. After washing with purified water, tissue sections were counterstained with hematoxylin for 1–2 min.DNA component analysisCultured cells fixed with 70% ethanol at least 4 h under −20 °C condition. The fixed cells stained with the Muse Cell Cycle Assay Kit (Merck Millipore Corporation, Darmstadt, Germany). The stained cells analyzed with Muse Cell Analyzer (Merck Millipore Corporation) were used for DNA content analysis.Karyotype analysisOur iPSCs were treated with 0.02 mg/ml colcemid. Those iPSCs exposed to a hypotonic solution and fixed with Carnoy’s fluid. We counted the chromosomal number in 50 cells and performed a G-banding analysis in 20 cells22.Production of interspecific chimeras and their immunological stainingTo evaluate whether iPSCs derived from Japanese ptarmigan could contribute to the generation of interspecific chimeras in chick embryos, iPSCs were stained with 10 μM CellTracker Green CMFDA (5-chloromethylfluorescein diacetate, C7025; Thermo Fisher Scientific) for 30 min. Eggs of white leghorn chicken were purchased from a local farm (Goto-furanjyo, Gifu, Japan). We injected the labeled Japanese ptarmigan iPSCs into stage X chick blastoderms and cultured the embryos26. To confirm the contribution of chimera, fluorescence was observed after 72 h. To analyze the tissue-level contribution of chimera, embryos on day 5. The embryos were embedded in optimal cutting temperature compound (Sakura Finetek Japan, Tokyo, Japan), frozen in liquid nitrogen, and stored at −80 °C until use. Cryosections 20 μm in thickness were prepared using a cryostat, air-dried for 30 min at room temperature, and fixed with 4% paraformaldehyde for 2 min at room temperature. After washing three times with PBS, sections were incubated with PBS containing 5% FBS for 1 h. After blocking with FBS, the sections were incubated with an anti-hygromicin resistance gene antibody (anti-HPT2; Supplementary Table 2) overnight. After washing three times with PBS, the sections were incubated with secondary antibody (goat anti-mouse IgG, Alexa Fluor 568; Supplementary Table 2) and Cellstain- DAPI solution (DOJINDO) for 1 h.Detection of contribution of chimera from genomeWe injected Japanese ptarmigan iPSCs (without CellTracker Green CMFDA label) into a stage X chicken blastoderms. On day 5, the entire chicken embryos were collected. The genome of each embryo was collected using NucleoSpin Tissue (U0952S; MACHEREY-NAGEL, Düren, Germany). After collecting the chimeric genome, we detected the reprogramming vector cassette using genomic PCR analysis using 50 ng of template genome. To extend the target sequence, we used the KOD FX Neo (KFX-201; TOYOBO). Primer information is provided in Supplementary Table 11. This analysis was performed according to the manufacturer’s protocol. The cycling program comprised 45 cycles of 94 °C for 120 s (initial denaturation), 98 °C for 10 s (denaturation), and 68 °C for 50 s (annealing and extension). After PCR, 2% agarose gel electrophoresis was performed. Gels were stained with GelGreen (517–53333; Biotium, Inc., Fremont, CA, USA).Real-time PCR was also performed to detect the contribution of chimera. The fluorescence probe and primers designed to detect chimeric contributions are summarized in Supplementary Table 12. The template was a 30 ng genome. The analysis was performed using 1 × THUNDERBIRD Probe qPCR Mix (QPS-101; TOYOBO), 0.3 μM of each primer, 0.2 μM of probe, and 1 × Rox. Fifty cycle of 95 °C for 60 s (initial denaturation), 95 °C for 15 s (denaturation), and 60 °C for 60 s (annealing and extension) were used. The expression levels of the target genes were normalized to that of chicken Tsc-2.RNA preparation and sequencing for RNA-seq analysisTotal RNA from iPSCs, fibroblasts, and chicken embryo stage X was collected using NucleoSpin Tissue (740952.50; MACHEREY-NAGEL). Triplicate samples of all iPSCs, fibroblasts, and chicken embryo stage X were prepared. To prepare the library, we used the TruSeq Stranded mRNA LT Sample Prep Kit (RS-122-2101; Illumina, San Diego, CA, USA). The quality of the library was evaluated using the Qubit DNA Assay (Thermo Fisher Scientific) on a TapeStation with a D1000 screen tape (Agilent Technologies, Santa Clara, CA, USA). The cDNA samples were used for the sequencing reaction on an Illumina HiSeq X sequencing machine, resulting in more than 40 M reads with 150 bp ends for each sample, except chicken fibroblast No. 3, which displayed more than 40 M reads with 75 bp ends. To analyze the RNA-seq data, we used the CLC Genomic Workbench (CLC Bio, Aarhus, Denmark). In the trim read step, low-quality sequence with the quality score of the CLC workbench, 5′ end, 3′ end, and short sequences (shorter than 15 sequences) were removed. The trimmed sequence data were mapped onto the chicken reference genome. Gene expression data were obtained in this step. PCA was performed and a heat map created with CLC Genomic Workbench using gene expression data. In this step, normalization was automatically performed using TMM methods. To compare chicken cells, RNA-seq data from SRA (SRP115012 (GEO: GSE102353) and SRP087639 (GSE86592) were used. The RNA-seq data has been submitted to the DNA DataBank of Japan under accession number DRA013522 (Submission), PRJDB13093(BioProject), SAMD00444261–SAMD00444287 (BioSample).Statistics and reproducibilityNonparametric multiple comparison analysis used the Steal–Dwass test (Figs. 2e, 3 [Okinawa rail, Japanese ptarmigan, and Blakiston’s fish owl], 4d, 4f, 5b, 5d, 5f, 5h, 10i). For nonparametric independent two-group analysis, we used the Mann–Whitney U test (Fig. 3, for mouse and chicken, and 4b). Statistically significant differences are indicated by *(p  More

Deforested areas rim a highway running through the state of Amazonas, in Brazil.Credit: Michael Dantas/AFP/Getty

Countries are failing to meet international targets to stop global forest loss and degradation by 2030, according to a report. It is the first to measure progress since world leaders set the targets last year at the 26th United Nations Climate Change Conference of the Parties (COP26) in Glasgow, UK. Preserving forests, which can store carbon and, in some cases, provide local cooling, is a crucial part of a larger strategy to curb global warming.
Tropical forests have big climate benefits beyond carbon storage
The analysis, called the Forest Declaration Assessment, shows that the rate of global deforestation slowed by 6.3% in 2021, compared with the baseline average for 2018–20. But this “modest” progress falls short of the annual 10% cut needed to end deforestation by 2030, says Erin Matson, a consultant at Climate Focus, an advisory company headquartered in Amsterdam, and author of the assessment, published on 24 October.“It’s a good start, but we are not on track,” Matson said at a press briefing, although she cautioned that the assessment looks at only one year’s worth of data. A clearer picture of deforestation trends will emerge in successive years, she added.The assessment, which was carried out by a number of civil-society and research groups, including the World Resources Institute, an environmental think tank in Washington DC, comes as nations gear up for the next big climate summit (COP27), to be held in November in Sharm El-Sheikh, Egypt. Scientists agree that in order to limit global warming to 1.5–2 °C above preindustrial levels — a threshold beyond which Earth’s climate will become profoundly disrupted — deforestation must end.Tropical forests are keyTo track deforestation over the past year, the groups analysed indicators such as changes in forest canopy, as measured by satellite data, and the forest landscape integrity index, which is a measure of the ecological health of forests. The slow progress they found is mainly attributable to a few tropical countries where deforestation is highest (see ‘Progress report’). Among them is Brazil — the world’s largest contributor to tree loss — which saw a 3% rise in the rate of deforestation in 2021, compared with the baseline years. Rates also rose in heavy deforesters Bolivia and the Democratic Republic of the Congo, by 6% and 3%, respectively, over the same period.

Adapted from the 2022 Forest Declaration Assessment

The loss of tropical forests, in particular, is worrisome because a growing body of research shows that besides sequestering carbon, these forests can physically cool nearby areas by creating clouds, humidifying the air and releasing certain cooling molecules. Keeping tropical forests standing provides a massive boost to global cooling that current policies ignore, says a report, “Not Just Carbon”, released alongside the Forest Declaration Assessment.A region made up of tropical countries in Asia is the only one on track to halt deforestation by 2030, according to the assessment (see ‘Movement towards goal’). The region cut the rate at which it lost humid, old-growth forests last year by 20% from the 2018–20 baseline, mostly thanks to large strides made by Indonesia — normally one of the world’s largest contributors to deforestation — where the loss of old-growth forests fell by 25% in 2021 compared with the previous year.

Adapted from the 2022 Forest Declaration Assessment

“The progress we see is driven by exceptional results in some countries,” Matson said.Efforts by the government and corporations in Indonesia to address the environmental harms of palm-oil production were key to progress, the assessment says. For example, as of 2020, more than 80% of palm-oil refiners had promised not to cut down or degrade any more forests. And in 2018, the Indonesian government imposed a moratorium on new palm-oil plantations. But the ban expired last year, raising concerns that progress might eventually be reversed.Finance laggingGlobal demand for commodities such as beef, fossil fuels and timber drive much of the forest loss that occurs today, as industry seeks to clear trees for new pastures and resource extraction. Matson said that many governments haven’t introduced reforms, such as protected-area regulations or fiscal incentives to encourage the private sector to safeguard forests, and that this is stalling progress.“Stronger mandatory action is needed,” she said.
How much can forests fight climate change?
In particular, nations are lagging behind in terms of fiscal support for forest protection and restoration. On the basis of previous assessments, the report estimates that forest conservation efforts require somewhere between US$45 billion and $460 billion per year if nations are to meet the 2030 goal. At present, commitments average less than 1% of what is needed per year, it concludes.Matson said that nations need to improve transparency on financing by setting interim milestones and publicly reporting progress. Michael Wolosin, a climate-solutions adviser at Conservation International, a non-profit environmental organization headquartered in Arlington, Virginia, would like to see donor countries recommit to their forest finance pledges at COP27 this year.However, Constance McDermott, an environmental-change researcher at the University of Oxford, UK, cautions against focusing too much on “estimates of forest cover change and dollars spent”. Social equity for Indigenous people and those in local communities should be part of discussions relating to deforestation, but is mostly missing, she says. These communities are the best forest stewards, and more effort is needed to support them by strengthening land rights and addressing land-use challenges that they identify, she says.Otherwise, McDermott warns that “global efforts to stop deforestation are more than likely to reinforce global, national and local inequalities”. More