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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

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A roaring trans-European herring trade that began in the Viking Age might have depleted stocks1.

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doi: https://doi.org/10.1038/d41586-022-03431-y

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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

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