Ecology

We hand-collected adult toads (n = 8 individuals per site) from four sites across the toads’ tropical range within Australia, from Townsville, Qld in the east (GPS coordinates: − 19.26, 146.82, 14 m altitude) to Richmond, Qld (− 20.73, 143.14, 218 m altitude), Middle Point, NT (− 12.56, 131.33, 12 m altitude) and Kununurra, WA (− 15.78, 128.74, 49 m altitude) in the west. That transect spans the toads’ 80-year invasion history. Although both temperatures and precipitation exhibit a general east–west cline, the greatest disparities in the duration of hot dry conditions per year lie between the easternmost site (Townsville) and the three other sites (Fig. 1). We recorded toad mass (after gently squeezing the animal in a standardized manner to induce it to empty its bladder) and snout-vent length (SVL) immediately before conducting the trials.
Figure 1

Data from Australian Bureau of Meteorology7.

Mean climatic conditions in the four sites from which we collected cane toads (Rhinella marina) for use in laboratory trials. The red line connects mean monthly maximum air temperatures, the green line shows mean monthly air temperatures, and the blue line shows mean monthly minimum air temperatures. Histograms show mean monthly rainfall.

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Toads were not fed for three days prior to experiments, to ensure they would not defecate during the experiment and minimize variability in mass due to stomach contents. Toads from all four populations were housed in a room kept at 18 °C, then moved concurrently to a temperature-controlled room set at 37 °C. All toads were in separate containers (ventilated plastic boxes of 1-L capacity), half of which had dry paper towel as substrate whereas the other half had 40 mL of water, enough to keep the ventral portion of the body moist but not the rest of the body.
We measured toad body temperatures at the beginning of the trial, and after 20 min and 40 min, using an infrared thermometer (Digitech QM7215) held ~ 10 cm from the toad’s dorsal surface. At the beginning and end of the experiment we measured internal temperatures with a cloacal probe (Digitech QM7215 with probe attachment), to check that our measurements of external body temperature offer robust estimates of internal temperature also. Cloacal temperatures were taken within 10 s of each toad’s removal from the container. After a trial, toads were kept at a temperature of 25 °C, allowed to fully hydrate and monitored for wellbeing during recovery. No adverse effects of the trials were evident.
We used mixed model repeated measures analysis to identify factors affecting body temperatures of cane toads during the 40-min heating trials. Sex and body mass were used as covariates in the analysis with climate at each collection site (# consecutive months per year with average maximum temperature  > 30 °C and with  More

When first described, it was hypothesized that the circular traces preserved in basal birds from the Jehol Biota represented remnants of the PFM of mature or nearly mature follicles within the left ovary9. The absence of calcified eggshell in the oviduct suggested that ovulation had not begun in any specimen. In one enantiornithine (STM29-8), the unusual surface texture in the purported follicles was interpreted as the imprints of a well-developed network of blood vessels within a highly vascularized PFM9 (Supplementary Fig. 3).
The results of our analyses support identification of the remains preserved in enantiornithine STM10–12 as remnants of the ovary and a vascularized PFM. The tissues in STM10–12 present the same morphological and histochemical characteristics as those of an avian chordae, a contractile structure made of intertwined collagen fibers with smooth muscle fibers that expel the oocyte during ovulation in extant birds (Fig. 2). STM10–12 also preserves structures morphologically consistent with extant blood vessels (Fig. 3). Given that the analyzed fragments of purported follicles in STM10–12 present virtually all the tissue characteristics (i.e., appropriate size, morphology, and histochemistry) of the three main components found in extant PFMs (i.e., smooth muscle fibers, collagen fibers, and blood vessels) the most parsimonious and plausible conclusion is that the circular structures in STM10–12 are indeed the fossilized remnants of pre-ovulatory ovarian follicles, also consistent with their preserved anatomical location. Fossilized structures morphologically consistent with collagen fibers and muscle fibers have been previously identified in numerous other Mesozoic specimens (e.g., refs. 26,27,28,29,30,31). This study contributes to the mounting evidence that such tissue components can preserve in deep-time.
The fossilized follicles in STM10–12 are by no means completely preserved, but rather represent fragments of these structures. We found no histological evidence of other tissues that are found in the PFM of extant pre-ovulatory follicles, such as non-collagenous fibers of the inner perivitelline membrane, granulosa cells, nerve fibers32, and the ovarian surface epithelium (Supplementary Fig. 1).
Fossilized blood vessels have been previously reported in some specimens of Mesozoic dinosaurs and Cenozoic turtles (e.g., refs. 33,34,35,36). The vessels in STM10–12 are not consistent in morphology with fungal hyphae in that they lack septae and fungal hyphae are much smaller (see refs. 33,34). Given their size and morphology, the most logical interpretation is that these structures are remnants of original blood vessels belonging to an originally highly vascularized PFM.
The fossilization of blood vessels is apparently more common than generally recognized in material from the Mesozoic and these structures have already been thoroughly documented both morphologically and chemically (e.g., refs. 26,29,33,34,37,38,39). In previous studies, fossil blood vessels were mostly observed in three dimensions (3D), photographed as ‘floating’ material in demineralizing solutions, whereas in this study the blood vessels were observed in 2D sections exposing the blood vessels through longitudinal cuts (Fig. 3). In blood vessels analyzed in 3D, branching is also commonly observed (e.g., see refs. 33,38,40) but in contrast, in sectioned vessels branching is much less common (e.g., see Fig. 3f–h), which may explain why very few branching patterns were observed in our sample (only in Fig. 3c).
Noteworthy, blood vessels in STM10–12 were only visible in the demineralized paraffin slides (Fig. 3) but not in the ground-sections nor the SEM images (Fig. 2). A possible logical explanation is that this is due to differences in tissue compaction and distortion between the samples in the ground-sections and the paraffin slides. In the ground sections, the tissues were embedded in resin without being demineralized. Resin-embedded tissues are tightly compacted (Fig. 2d) and do not undergo any significant distortion during preparation. On the other hand, demineralized tissues that get embedded in paraffin go through multiple distortion and tearing events (i.e., during demineralization, processing through different solutions, but especially after being cut on a microtome and placed on top of warm water in the water bath prior to mounting on glass slides, see Supplementary Methods). These distortions create an artificial ‘decompaction’ of the fossil tissues in the paraffin slides (which occurs commonly even while making slides of extant tissues), and this is most likely what enabled the visualization of blood vessels in STM10–12 (Figs. 2g–j and 3) that were not directly visible in ground-sections nor through SEM (Fig. 2d). These results suggest that the three methods employed here (ground-sectioning, SEM, and paraffin histology) yield complementary information and, when used together, can help to provide more rigorous identifications and clarify our understanding of fossilized soft-tissues.
EDS showed that the fossilized soft-tissues preserved in STM10–12 underwent alumino-silicification (Fig. 4). This same process has been reported in fossilized branchiopod (clam shrimp) eggs also from the Jehol Biota, where all the envelopes were made of calcium phosphate but some of the eggs had their internal contents replaced by alumino-silicates41. An explanation for the mechanism of alumino-silicate replacement was not provided for the Jehol clam shrimp eggs41, but we suggest that clay minerals from sediments were involved. Clay minerals have been determined to be important agents in the fossilization of soft-tissues in other settings (such as the Ordovician Soom Shale of South Africa and the Cambrian Burgess Shale of Canada)42,43. In these cases, the soft-tissues were replaced rapidly after death by authigenic clay minerals (via direct precipitation of authigenic clays onto the tissues). It is the most logical explanation for the process of alumino-silicification seen in the tissues of STM10–12, and this process may have happened rapidly after death before extensive tissue decay.
EDS also revealed an enrichment in iron, indicating that the soft-tissues may also have experienced some limited mineralization with iron oxides (Fig. 4, see the low iron levels). The mineralization of soft-tissues via iron oxides (such as goethite and biogenic iron oxyhydroxide) has been reported in Mesozoic dinosaurs38 and a similar process may have occurred as well in STM10–12. A potential source for this iron may be the pyroclastic flows that intermittently interrupted the deposition of lacustrine sediments, and/or microbial mats, as proposed for Jehol invertebrates44,45. Moreover, based on our new data, we also suggest that another source of this iron may be the hemoglobin (a protein found in red blood cells) coming directly from the blood vessels of the organism itself. An experimental study on ostrich blood vessels demonstrated that iron and oxygen from hemoglobin play a key role in tissue stability and in the exceptional preservation of soft-tissues in deep-time38. This hypothesis could be tested in the future with immunohistochemistry and antibodies raised against avian hemoglobin.
Although pyritization (via iron sulfides) has been demonstrated in the soft-body parts of Jehol insects and hypothesized to play an important role in the preservation of soft-tissues in Jehol fossils44,45, the EDS data here do not show any sulfur in the sample, meaning STM10–12 underwent a different fossilization process and that pyritization is simply one of the many processes involved in soft-tissue preservation in the Jehol paleolakes.
All of the histological and histochemical data collected here (Figs. 1–4) demonstrate the exceptional preservation of the soft-tissues in STM10–12 (i.e., fossilized chordae and blood vessels from the PFM), but more precise and more specific chemical analyses (such as synchrotron-FTIR, or immunohistochemistry) are needed to fully characterize the preservation of these tissues at a deeper molecular level.
Until this study, interpretations of the purported follicles as ingested seeds (e.g., see Supplementary Fig. 4) indeed represented a viable alternative hypothesis13,14. Testing these competing hypotheses was imperative to our understanding of both the evolution of the paravian reproductive system (i.e., to confirm whether or not early birds indeed had only one functional ovary like extant birds and lacked strong follicular hierarchy) and digestive system (as yet there is no direct evidence regarding the diet of enantiornithines in the Jehol46,47).
Many morphological arguments have previously been raised against the interpretation of these remains as ingested seeds5. For example, the preserved structures also lack the surficial ornamentation observed in most fossilized seeds (Supplementary Fig. 4). Furthermore, the tissues here identified in STM10–12 (smooth muscle fibers, collagen fibers, and blood vessels; Figs. 2 and 3) are strictly animal tissues and are non-existent in plants. None of the histological slides of STM10–12 reveal tissues reminiscent of fossilized plant material with their characteristic cell walls22,48, from either gymnosperms or angiosperm seed or fruit tissues (e.g., cuticle, seed coat or testa, internal integument layers, and embryonic tissues; e.g., refs. 22,49,50). Although our original intent was to directly compare the tissues in STM10–12 with a fossil seed preserved in the stomach of the holotype of Jeholornis prima (Supplementary Fig. 4), close examination showed that the seeds in this specimen are only impressions, and thus cannot be used for proper comparison with seed tissues. Lastly, the STM10–12 samples were also checked for the presence of phytoliths, which are microscopic structures made of silica that are found in plant tissues and can persist for millions of years after the decay of the plant51, but none were found.
Although the preservation of ovarian follicles in STM10–12 is confirmed, this hypothesis needs to be independently tested in each individual specimen (Supplementary Table 1); as yet, it is still possible some of the purported follicles in other specimens may represent ingested seeds. Confirmation of follicle preservation in one specimen does nonetheless put an end to the controversy regarding whether or not such remains can preserve and confirms that only a single functional ovary was present in at least some non-neornithine avians8,9. Additionally, our results support paleobiological hypotheses based upon the original identification, such as observations regarding the relative sizes of these ovarian follicles. Compared to modern birds only subtle size variations are observed in the fossilized follicles of Jehol birds (meaning follicular hierarchy was absent), leading to the inference that in basal birds yolk deposition occurred much more slowly than in extant bird due to the lower metabolic rates of non-ornithuromorph birds8,9. This study reveals that this was true at least for enantiornithine STM10–12, which suggests this may be similarly true about other non-ornithuromorph birds in which slow growth rates are observed through osteohistology. It is likely that a strong follicular hierarchy is a derived feature of a subset of the Ornithuromorpha (because some of them still retain plesiomorphically slower growth rates), but only direct evidence with preserved follicles in Cretaceous ornithuromorphs could confirm this hypothesis.
The Early Cretaceous Jehol Biota of China preserves one of most extraordinary extinct fauna and flora ever discovered, revealed through a taphonomic environment that was extremely conducive to the fossilization of both hard and soft-tissues1,4. Few histological analyses on preserved soft-tissues exist due to the destructive nature of most of these methods. However, our results demonstrate that these types of analyses can eliminate doubt and help to further understand preservation. Previous taphonomic studies conducted on Jehol invertebrates showed that alumino-silicification and pyritization help preserve soft-tissues in the Jehol41,45. In STM10–12, no pyritization was found, but the tissues apparently underwent alumino-silicification and a slight mineralization with iron, potentially iron oxides. This suggests that soft-tissue fossilization in the Jehol is case-specific and that varied mechanisms were involved. In the case of animal tissues, it is possible that preservation of soft tissues was facilitated by endogenous iron and oxygen from the hemoglobin in blood26,38. The abundant blood vessels in the PFM may explain the high preservation potential of ovarian follicles, although this hypothesis cannot explain why other tissues and organs that are also highly vascularized are not preserved in the same specimens.
In STM10–12 and all other specimens with purported follicles, the overall spherical shape of the follicles is preserved in 2D. Therefore, it is also possible that organic remnants from the original spherical yolk and/or the inner perivitelline membrane are preserved, but extensive further analyses (using different methods such as immunohistochemistry or spectroscopy) are required to confirm this. Additionally, the most external tissue covering preovulatory follicles in extant birds is made of pancytokeratin18, a hydrophobic protein with a high potential for fossilization52. We propose that this hydrophobic molecule may have acted as a barrier and also facilitated the exceptional preservation of follicles. Much more research is necessary to confirm this hypothesis, and more studies on preserved soft-tissues in the Jehol are necessary to further shed light on the modes of tissue preservation through deep-time. More

Does manual control reduce COTS densities?
We first asked whether manual control could control COTS densities at a site. To do this, we examined COTS control data from 52 sites with permanently marked coral monitoring (RHIS) sampling points, where repeated manual control of COTS took place from July 2013 to December 2017. These 52 sites were located at 21 reefs and were distributed across three different management zones (Fig. 1).
Over the 4.5 year period, individual sites were visited on average 15 ± 6.2 (s.d.) times (range 5–36), with the number of voyages to a site in a year ranging from 0 to 11 (mean 3.2 ± 2.3 s.d. voyages yr−1). At the start of the Control Program, COTS densities at the 52 sites averaged 40 ± 54 s.d. individuals ha−1 (range 0–237) and were above an ecologically sustainable density threshold of 3 ha−1 at 45 sites. We use this threshold as a benchmark since coral growth is outpaced by predation by an ‘average’ COTS population at sites with low coral cover (estimated as 5 ha−1 for just the three largest size categories)27, and COTS fertilization (and thus reproductive) success increases substantially at densities of ≥ 3 ha−1 due to Allee effects 28. Manual control was effective in rapidly reducing COTS densities with the median density of COTS encountered being significantly lower on the second and subsequent voyages to a site than on the first voyage (Fig. 2; Friedman’s chi-squared = 9.31, df = 1, P = 0.0023; Table S1). This decline was initially rapid with one voyage sufficient to bring the median COTS density to below the ecologically sustainable threshold. The 75th percentile of sites reached this threshold following five culling voyages and fluctuated around the threshold until the number of sites in the analysis dropped to below 5 sites at 23 voyages and two and one site at voyages 27 and 29 respectively. Over the period of the study an average of 126 COTS ha−1 (range 5–723) were removed from each site.
Figure 2

The effect of number of control voyages on the density of Pacific Crown-of-Thorns Starfish (COTS), Acanthaster cf. solaris. Manual control of COTS took place at a total of 52 sites at 21 reefs in three different types of spatial zoning in the Cairns sector of the Great Barrier Reef from July 2013 to December 2017. The density of COTS encountered during a voyage at a site declined as a function of the number of voyages that had previously visited that site. This decline was initially rapid and after roughly five voyages COTS densities fluctuated while remaining low. Note, beyond 22 visits, sample sizes decline substantially with just six sites visited 23 or more times; variation in COTS densities increases dramatically as a result. Dashed line = the ecologically sustainable threshold for COTS outbreaks27, i.e. the density of COTS that can be sustained before coral cover is lost, Solid bar = median, box = quartiles, whiskers = extremes, circles = outliers. Sample sizes given above the x-axis.

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The second and subsequent voyages to a site resulted in additional COTS being culled indicating the need for repeated visits. This appears to be largely due to the fact that not all COTS present at a site are visible and available to be culled at any one time29 and, to a lesser extent, to immigration into controlled sites from adjacent areas (see below). This result emphasizes the need for repeat voyages to a site in order to achieve sustained, reliable reductions to below the ecological threshold.
The impact of manual control was not consistent across the four COTS size categories (Fig. 3). The median densities of the largest size classes ( > 15–25 cm,  > 25–40 cm, and  > 40 cm diameter) were significantly lower on the second and subsequent voyages than on the first voyage (Friedman’s chi-squared = 27, df = 1, P  40 cm diameter) show a sharp decline during the first four voyages, while densities of the smallest size class (35) in the same period, with these declines being at least in part attributed to the current COTS population outbreaks36. Interestingly, in reporting on inshore coral reef surveys, Thompson, et al.37 noted that ongoing manual control of COTS at the Frankland Islands between January 2017 and March 2018 had contributed to mitigating their impacts on coral loss.
Not only was the final absolute hard coral cover related to the effort invested in manual control at a site but the proportional change in hard coral cover at a site relative to its initial cover across the 52 sites over the 4.5 years period was significantly and positively related to the number of voyages that visited these sites (linear regression: R2 = 0.19, F1, 50 = 11.99, P  More

Materials
Sediment samples used in this study were collected by IODP Expedition 329 in the South Pacific Gyre (SPG; Supplementary Fig. 1)7. The samples collected from Site U1365 (23° 51.0377′S 165° 38.6502′W) are zeolitic and/or metalliferous pelagic clays, those from Site U1368 (27° 54.9920′S, 123° 09.6561′W) are calcareous nannofossil oozes, and those from Site U1370 (41° 51.1267′S, 153° 06.3674′W) are black metalliferous clay, containing light yellowish brown clay-bearing nannofossil ooze. The water depths of these sites are respectively 5697, 3739, and 5075 meters below sea level (mbsl) at U1365, U1368, and U1370. Whole-round core samples of U1365C 8H-2 (obtained from 68.9 meters below seafloor [mbsf]), 95.4 Ma), U1365C 9H-3 (74.5 mbsf), U1368D 1H-2 (1.6 mbsf), 1368D 2H-5 (14.7 mbsf), and U1370F 7H-6 (62.9 mbsf) were used for incubation experiments as described in the following section. Those samples were from horizons with no observable coring disturbance by visual core description35. Drilling fluid contamination assessment by chemical tracer revealed minimal drilling contamination of the samples (≤1 cell/g-sediment for U1365C 8H-2 and 1368D 2H-5, below detection to 0 cell/g-sediment for U1365C 9H-3, U1368D 1H-2, and U1370F 7H-6)35. All Expedition 329 data are archived and available online in the IODP database (http://iodp.tamu.edu/tasapps) and archived online in the IODP Expedition 329 Proceedings7. Additional sediment age estimation is available in reports by Dunlea et al.13 and Alvarez Zarikian et al.36.
Incubation experiments
To identify autotrophic and heterotrophic microbial populations, as well as their potential rates for growth and substrate uptake, sediment samples were incubated with stable isotope-labeled substrates (13C6-glucose, 13C2-acetate, 13C3-pyruvate, 13C-bicarbonate, 13C-15N-amino acids mix [mixture of 20 Amino Acids], and 15N-ammonium). The incubation experiments were initiated onboard during expedition. Avoiding outer edges of the sediment cores (which are likelier to be contaminated by drill fluid), each 15 cm3 sample was taken from an interior portion of a core with a sterile tip-cut 30 mL syringe, and placed in a 50 cm3 sterile glass vial (Nichidenrika-Glass Co. Ltd.) sealed with a sterile rubber stopper and a screw cap, followed by flushing with 0.22 μm filter-sterilized nitrogen gas and storage at 10 °C. All of the sterilization was done by autoclaving of the materials at 121 °C for 20 min. Given O2 concentrations in interstitial water during Expedition 329 (~70 μM for U1365C 8H-2 and 9H-3, ~150 μM for U1368D 1H-2, ~130 μM for 1368D 2H-5, and ~1.5 μM for U1370F 7H-67), we set O2 concentration in the headspace of vials (other than U1370F 7H-6) at ~3.3% (v/v), which corresponds to the aqueous concentration of oxygen at ~43 μM (assuming salinity of sea water), by adding 0.22 μm filter-sterilized air. The labeled substrates were injected (15 μM of 13C-labeled substrates, and 1.5 μM of 15N-labeled ammonium, dissolved in 50–100 μL of sterile water) onto each subcore sample by syringe and needle and incubated at 10 °C (Supplementary Fig. 3a). All reagents and gas components, including air used for sample preparation, were filtered through 0.22 μm syringe top filter. After setting up the incubations, one of the vials from each set of incubation had no substrate added, and a sediment split from the same sample was fixed by adding equal volume (15 mL) of 4% paraformaldehyde (PFA) in PBS solution for 5 h at 4 °C (time point T0). At each of three time points (T1: ~3 weeks [21 days], T2: 6 weeks [68 days], T3: 18 months [557 days] after starting incubation), vials were opened and sediment samples were fixed with equal volume (15 mL) of 4% PFA in PBS solution for 5 h at 4 °C. Fixed samples were frozen at −80 °C. After storage, they were washed twice with PBS and preserved in PBS/ethanol (1:1 [v/v]) at −20 °C until analysis.
Cell enumeration and selective sorting onto the membrane
To efficiently analyze substrate incorporation into microbial cells with NanoSIMS, cells were separated from their sediment matrix and fluorescence-activated cell sorting (FACS) was conducted to concentrate and purify cells in a small area for analysis (~0.5 mm2, Supplementary Fig. 3b)14,37. Cell separation, enumeration, and FACS were all conducted in the clean-booth and clean-room facilities at the Kochi Institute for Core Sample Research, Japan Agency for Marine-Earth Science and Technology (JAMSTEC).
Cell separation, microscopy, and sorting procedures followed the method of Morono et al.37 with modifications. Eight milliliters of fixed slurry (1/3 [v/v] sediment in ethanol-PBS solution) was mixed with the same volume of 2.5% NaCl solution, followed by centrifugation at 4500 × g for 15 min, after which the supernatant was discarded and the pellet resuspended by adding 2.5% NaCl solution to be 20 mL of sediment slurry. The sediment slurry was added with 2.5 mL each of detergent mix38 (100 mM EDTA, 100 mM sodium pyrophosphate, 1% [v/v] Tween 80) and methanol, then vigorously shaken for 60 min at 500 rpm using a Shake Master (Bio Medical Science, Tokyo, Japan). After shaking, the sediment slurry was sonicated (Bioruptor UCD-250; COSMO BIO) in an ice bath for 20 cycles of 30 s at 200 W on and 30 s off. The processed slurry was then carefully layered onto a manually layered high-density cushion solution consist of (from top) 4 mL of 30% [v/v] Nycodenz, 4 mL of 50% [v/v] nycodentz, 4 mL of 80% [v/v] Nycodentz, and 4 mL of 67% [w/v] of sodium polytungstate. Samples were centrifuged at 4000 × g for 120 min, after which the supernatant, including the high-density layer(s), was carefully removed and transferred to a separate vial. Cells in the supernatant were trapped onto an Anopore Inorganic Membrane (Anodisc, Whatmann, Kent, UK), washed with TE buffer, and then stained with 100 μL of SYBR Green I staining solution. After staining for 5 min, the SYBR-stained cells were washed with 2 mL of TE buffer, and then the membrane was placed into a 50 mL centrifuge tube containing 5 mL of TE buffer. Cells were detached from the membrane by sonication at 20 W for 10–30 s using a Model UH-50 Ultrasonic Homogenizer (SMT Co. Ltd., Tokyo, Japan) and concentrated to be 1.5 mL by centrifugation at 7000 × g for 10 min and discarding 3.5 mL of the supernatant. Part (0.5 mL) of the stained cell suspension was filtered onto 0.22-μm pore size black polycarbonate membrane (Isopore GTBP02500; Millipore) and used for counting microbial cells by the fluorescence color-based discriminative cell enumeration method39,40. Cells were sorted following the flow cytometry protocol of Morono et al.37 directly from the sorter onto NanoSIMS-compatible 0.2-μm polycarbonate filters coated with indium tin oxide (ITO)14,41 and non-coated membrane (Isopore GTBP02500; Millipore). ITO coating on polycarbonate membranes (Isopore GTBP02500; Millipore) was prepared by a sputtering deposition technique at Astellatech Co. Ltd. The sorted cells on the non-coated membrane were stored at −20 °C until DNA extraction.
NanoSIMS analysis of single cell-image acquisition and data processing
Cell targets were identified by fluorescence of SYBR Green I stain and marked on NanoSIMS membranes with a laser dissection microscope (LMD6000; Leica Microsystems) for ease of rediscovery on the NanoSIMS (an example is shown in Supplementary Fig. 3b). Microbial cells that incorporated stable isotope-labeled substrates were analyzed using NanoSIMS 50L (AMETEK Co. Ltd., CAMECA BU) at the Kochi Institute for Core Sample Research, JAMSTEC. Samples on the ITO-coated polycarbonate membrane were pre-sputtered at high beam currents (30 pA/s/µm2) before measurement. The 12C−, 13C−, 12C14N−, 12C15N− and 32S− secondary ions were collected and measured in parallel at a mass resolution of 8000 that is sufficient to separate 13C− from the 12CH− and 12C15N− from 13C14N−. Samples were measured using a 1–2 pA Cs+ primary beam that was rastered over 25 × 25 µm field of a 256 × 256 pixels with a counting time of 5 ms per pixel. Recorded images and data were processed using CAMECA WinImage software and OpenMIMS plugin42 in ImageJ43 distribution of Fiji44. Different scans of each image were aligned to correct image drift during acquisition. Final images were created by adding the secondary ion counts of each recorded secondary ion from each pixel over all scans. Intracellular carbon and nitrogen uptake from stable isotope-labeled substrates was calculated by drawing regions of interest (ROI) on CN− images (recognizing cells in the images) and calculating 13C/12C and the 15N/14N ratio (calculated from the 12C15N/12C14N ratio). Instrumental mass fractionation of NanoSIMS analysis was calibrated by the conversion factor obtained by comparing carbon and nitrogen isotopic ratios of E. coli cells of varying isotopic enrichments measured at single cells with NanoSIMS and at bulk with an elemental analyzer/isotope ratio mass spectrometer (EA/IRMS, FlashEA 1112/DeltaPlus Advantage, Thermo Fisher Scientific). Concentration of bicarbonate (DIC) in the original sample determined on board7 was used to calculate substrate incorporation ratio (atom %) for bicarbonate in single cells.
Biomass and isotope calculations
Isotope incorporation data analysis and display as violin plots of the kernel density function were done using R45 with the “ggplot2”46, “cowplot”47, “ggsci”48, “scales”49 packages. The “active” cell ROIs, those incorporated 13C- and/or 15N-labeled substrates, were determined by their isotopic ratio exceeding the 99.7% confidence interval of background carbon and nitrogen isotopic abundance of polycarbonate membranes (1.24 atom % for 13C and 0.446 atom % for 15N). As measures for the rate of microbial biomass synthesis, we calculated the biomass-based specific growth rate and the substrate incorporation-based biomass generation rates. The biomass-based specific growth rates (µB, Eq. (1)) were calculated from the abundance of cells in the sediment samples at the start of incubation X0 and the abundance at the incubation period t (Xt). The substrate incorporation-based biomass generation rates14,50 (CµS and NµS, Eqs. (2) and (3), for carbon and nitrogen substrates, respectively) were calculated from the fractional abundance of isotope label in cellular biomass, where μS is the biomass generation rate (encompassing both cell maintenance and generation of new cells), t is the length of the incubation, Flabel is the labeling strength, Ft is the single-cell NanoSIMS measurement, and Fnat is the natural abundance. For the calculation of CµS and NµS, a conservative approach was used by only including ROIs where either 13C or 15N ratio was above the 99.7% confidence interval of backgrounds shown above. These rate calculations assumed that the carbon and nitrogen for the biomass generation were all derived from the substrates.

$$mu_{rm{B}} = frac{{ln, X_{rm{t}} – ln, X_{rm{0}}}}{t}$$
(1)

$${!,}^{C}mu_{rm{S}} = left( – lnleft( 1 – frac{left( {!,}^{C} F_{rm{t}} – ,{!,}^{C} F_{{rm{nat}}} right)}{left({!,}^C F_{{rm{label}}} – ,{!,}^{C} F_{{rm{nat}}} right)} right)right)/t$$
(2)

$${!,}^{N}mu_{rm{S}} = left(- lnleft( 1 – frac{left({!,}^{N} F_{rm{t}} – ,{!,}^{N} F_{{rm{nat}}} right)}{left( {!,}^{N} F_{{rm{label}}} – ,{!,}^{N} F_{{rm{nat}}} right)}right)right){mathrm{/}}t$$
(3)

To document the physiological status of microorganisms in the original sediment samples, the fraction of microbes that originally existed in the sediment samples (f0) was calculated from the observed active ROI fractional ratio (ft), a factor of biomass increase (A) by following equation (Eqs. (4)–(6)).

$$X_{rm{t}} = AX_{rm{0}}$$
(4)

$$X_{rm{0}}left( 1 – f_{rm{0}} right) = X_{rm{t}}left( 1 – f_{rm{t}} right)quad left[ f_{rm{t}} , More

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AGAGE data for marine N2O studies
The network of Advanced Global Atmospheric Gases Experiment (AGAGE) stations has been quantifying greenhouse gas levels including N2O since the late 1970s27. This global network now produces high frequency, high precision measurements; the data improved substantially in the mid-1990s with the incorporation of higher precision methodologies28. N2O data from the last ~20 years are measured every 40 min (Fig. 1b, Supplementary Figs. 1 and 2) and have a precision on any individual measurement of 0.1 ppb but much higher confidence for the ensemble28,29. Combining these measurements with atmospheric back trajectories generated by the HYSPLIT4 model from monitoring stations (Fig. 1a, Supplementary Data 1), we map the relationship between atmospheric N2O concentrations and marine O2 content. Similar analyses have previously been conducted to investigate recent continuing sources of chlorofluorocarbons from land30. The marine environment is dynamic, and time-resolution is difficult to sample for many biologically-active chemical parameters requiring in situ point measurements. The AGAGE dataset, however, does not suffer the same restrictions, and the high-frequency measurements can be utilized to investigate variability in detail, including the important interannual dynamics that arise due to the influence of El Niño–Southern Oscillation (ENSO) on these regions.
Land sources could obscure the attribution of marine N2O, so all back trajectories that intersect continents over a 20-day model run are excluded in our analysis. This rigid protocol limits the choice of stations and marine areas. Multiple stations were considered for analysis, but eliminated as being ill-suited for identifying features in the OMZs. The land and monsoonal influence in the Arabian Sea obscures any marine signal in that OMZ. Similarly, the available high precision North Pacific atmospheric measurement stations are located along the California coast and in Hawaii, and do not communicate with the eastern tropical North Pacific OMZ before mixing homogenizes any advected signal (Supplementary Fig. 3). The station at American Samoa, however, permits excellent monitoring of the ETSP. The easterly trade winds rapidly transport air parcels intersecting the region of low O2 across the Pacific (scale of days to weeks) before turbulent mixing with other air masses is able to fully mask the OMZ signal (Fig. 2). Moreover, along this trajectory, air tends to remain in the troposphere (Supplementary Fig. 4), minimizing stratospheric impacts on surface conditions. The raw N2O data from Samoa, when de-trended for the long-term anthropogenic effect and de-seasonalized to minimize the intra-annual oscillation driven largely by interhemispheric transport from the north as the Intertropical Convergence Zone moves southward during boreal winter (Supplementary Figs. 1 and 2), reveal striking relationships with the ocean biogeochemical state (Fig. 3).
Fig. 2: Back trajectory of Samoa atmospheric N2O.

Locations of air parcels with their de-trended nitrous oxide measurements at Samoa are plotted (a) 5, (b) 10, (c) 15, and (d) 20 days prior to arrival at Samoa. De-trended nitrous oxide measurements are represented by the color of the corresponding dot, as anomalies relative to the station mean. The data maintain a clear spatial delineation for the full 20-day period, with much higher concentrations passing over the eastern tropical Pacific, and much lower concentrations arriving from the west and the Southern Ocean. Overlain on this map are the 10 and 20% oxygen saturation horizons at the 1026.5 kg m−3 potential density level (black contours). The orange circle indicates the location of the Samoa station.

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Fig. 3: Gridded N2O anomalies relative to the Samoa mean.

De-trended and de-seasonalized N2O concentrations from Samoa (color) are gridded based on back-trajectory locations 15 days prior, and a mean value is calculated for air passing over each 5° grid cell at that time (color). Overlain on this map are the 10 and 20% oxygen saturation horizons at the 1026.5 kg m−3 potential density level (black lines). These two data sets come from entirely independent sources, and the coincidence between them is striking; high atmospheric N2O concentrations align with the oxygen minimum zone, extending westward into the tropical Pacific and down the Chilean coast of South America. The orange circle indicates the location of the Samoa station.

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The seasonal modulation in Samoa N2O has been well-established as an effect of northern influence. However, across the Pacific sites, trajectories remain mostly well-confined to the hemispheres in which they originated (Supplementary Fig. 5). The air parcels that reach Samoa from the northern hemisphere however, do not have markedly higher N2O concentrations than their southern counterparts. Many high concentration trajectories do travel across the Isthmus of Panama from the Caribbean (Supplementary Fig. 6), but these are filtered out by the land filter. Notably, 20 days prior there are still a large number of high concentration trajectories in the southern hemisphere, which travel northward off the coast of South America before transiting to Samoa. Indeed, very few trajectories actually cross over the South American continent; most are deflected either north or south along its western edge. As a result, land emissions of N2O from South America are unlikely to have a large impact on the visible spatial signal.
Oxygen minimum zones emerge as N2O hotspots
A divergence in concentrations is visible just one-day prior, with higher concentrations arriving from the north than from the south (Supplementary Fig. 3). This pattern remains moving further back in time; the highest concentrations pass over the eastern tropical Pacific, while the lowest come from the west passing over the Southern Ocean. The pattern becomes slightly less defined between 15 and 20 days-prior, as parcels with high concentrations diverge again along the western coasts of North and South America. Given the velocity of the trade winds and the distance between Samoa and the ETSP, ~8 m s−1 and 104 km, respectively, this time scale is reasonable. Certainly the observed atmospheric concentration anomalies of N2O at Samoa are a combined result of intra-hemispheric biological production with interhemispheric transport, but the analysis presented herein aims to tease apart the biological impact from the tropical Pacific ocean from the additional seasonal supply from the N2O-enriched northern hemisphere (Supplementary Fig. 7).
In defining the OMZ boundary by the 20% O2 saturation contour on the 1026.5 kg m−3 potential density surface, the 15-day back trajectories identify higher than average N2O among the air parcels passing over the OMZ (Fig. 3). This isopycnal layer, generally at depths of ~200–500 m (shallower in the eastern Pacific), tends to co-locate the minimum O2 concentration within the water column31. Whereas water this deep does not tend to equilibrate rapidly with the atmosphere, this density surface reflects broader features in the water column, such as the existence of a shallower N2O maximum driven by microbial processes18. This shallower N2O maximum can be more easily transmitted to the surface waters from the interplay of upwelling and lateral and vertical mixing32. Mesoscale and sub-mesoscale eddies in particular, below the spatial resolution that can be resolved by these data, are likely pathways that promote mixing of deeper N2O into the surface waters33,34.
The air masses reaching Samoa that pass over the OMZ are ~0.4 ppb higher than the remainder of the South Pacific. This intense localization to the lowest O2 waters in the eastern tropical Pacific, extending southward along the Peruvian and Chilean margin, suggests a disproportionate role of OMZs and coastal upwelling waters in generating atmospheric N2O relative to the open ocean. Extension of higher N2O anomalies along the equator outside of the 20% O2 contour also remain visible west to 140° W (Fig. 3), albeit with reduced magnitude, indicative of a separate likely N2O source from equatorial upwelling and subsequent outgassing35,36. Further, the air overlying the coastal Pacific along the South American upwelling region is especially high in its N2O content, indicative of the heterogeneities that exist even within suboxic waters17. The coastal-most waters are the most productive and the resulting organic matter fuels greater microbial generation of N2O. Moreover, the depth of the anoxic onset and coincident N2O maximum is generally shallower toward the coast, increasing the communication between ocean and air. Narrow, shallow shelves within the OMZs in particular are important producers of N2O17,23,37,38, but local sources with such spatial resolution cannot be identified via our methods.
While the absolute difference between N2O mixing ratios in air that has passed over the OMZ versus those that did not is small, the abundances above the OMZs are likely much higher but are diluted to the observed anomalies after 15 days of transport and mixing. Notably, the air parcels will continue to gain N2O as they travel over supersaturated water masses or lose N2O over undersaturated areas. The coherence of the 15-day backtracked Samoan N2O signal and its clear demarcation with the ETSP highlight the significance of the OMZs for the marine N2O efflux. Indeed, the offset between the air passing over the OMZ grid cells and the rest of the South Pacific matches the average seasonal variability in the Samoan data. The seasonal cycle, which includes terrestrial sources and is heavily influenced by seasonal shifts in the intertropical convergence zone, peaks in Austral winter (January/February) with minima 0.50 ± 0.13 ppb (se) lower in summer (July/August). The rapid N2O cycling rates inherent to the highly productive OMZs cause these regions to respond quickly to time-dependent shifts in the underlying biogeochemistry and physical structure of the water column18,24.
Further modulation by El Niño and La Niña
It is known that ETSP biogeochemistry is influenced by the El Niño/La Niña oscillations of the tropical Pacific39,40. La Niña brings enhanced upwelling of nutrients to the surface, thereby increasing primary production, further de-oxygenating the subsurface region, and supplying more organic matter to drive greater denitrification and N2O production13. Conversely, during an El Niño decreased upwelling reduces the surface productivity in the eastern tropical Pacific and deepens the oxycline, thereby contracting the OMZ and decreasing N2O emissions. In this study, the near-continuous data over 21 years permits a more quantitative analysis. The period from 1996 through 2016 included both a number of strong El Niño (1997/98, 2002/03, 2009/10, 2015/16) and La Niña (1998/99, 2007/08, 2010/11) states. Our analysis consists of dividing the N2O dataset among ENSO states in two different ways, both of which produced equivalent results: First, by aligning the N2O oscillation against the Niño 3.4 index, resulting in a time lag of 3 months relative to the ENSO state; second, by producing a running integral of the Niño 3.4 index forward in time, which essentially represents a cumulative effect of prolonged El Niño/La Niña periods (Fig. 1c). The Niño 3.4 index describes a 5-month running mean of sea surface temperature anomalies in the central equatorial Pacific, and was used because it is indicative of the broader Pacific region.
Strikingly, during La Niña, air parcels passing over OMZ grid cells increase on top of their already higher than average concentrations, whereas parcels that do not pass over OMZ grid cells exhibit no systematic change (Fig. 4). This increase with La Niña is not uniform across the OMZ, however, as the lowest O2 waters ( More

Experimental site and test cultivars
A field experiment of cover crop planting in a winter fallow field was conducted in Changsha (28° 11′ N, 113° 04′ E), Hunan Province, China, from 2014–2015. The soil in the experimental field was tidal clay, with 1.16% organic carbon, 0.17% total N, and a pH of 6.15.
Experimental design and field management
A randomized block experiment was established with 3 different treatments, including cover crops (Lolium spp. and Astragalus sinicus) with chicken grazing (+ C), cover crops without chicken grazing (− C), and a bare, fallow field (CK). Each field plot covers 140 m2, and there were three replications. To prevent the movement of water between adjacent plots, ridges were covered with a plastic sheet inserted into the soil to a depth of 0.5 m.
Ryegrass and milk vetch were planted on October 10th, 2014, at seed densities of 23 and 40 kg ha−1, respectively. Thirty-day-old yellow chickens were introduced into the field on November 25th. To ensure the homogeneity of the chicken manure inputs, a 3 m × 3 m cage was used during the process of chicken grazing. There were 30 chickens in each cage. Five kilograms of corn flour was fed to the chickens in each cage daily. The corn flour was 1.8% nitrogen. The cage was moved every 7 days in the chicken-grass plot until February 2, 2015. The quantity of in situ chicken manure input into the system within the symbiotic period (69 days) in these plots was estimated to be 96.3 t ha−1 by collecting the chicken waste in an underground container. The underground container was a square with a side length of 50 cm and a height of 10 cm. There were 3 symbiotic periods in these plots, and the chicken waste samples were collected every 12 h for three days. On March 27th, 2015, the average aboveground biomass of the cover crops was 11.7 t ha−1 in the + C plot and 14.4 t ha−1 in the − C plot. All the procedures used in this experiment were conducted in accordance with the Chinese Guidelines for Animal Welfare. The experimental procedures performed in the current study were approved by the Hunan Agricultural University Institutional Animal Care and Use Committee (Changsha, China). Furthermore, all the experimental protocols, including animal handling, were performed humanly, and animal welfare was specially considered. We further confirmed that no animals were harmed or stressed during the experimental period.
The cover crops were incorporated into the soil on March 27th, and all the plots were used to grow double-season rice. The early rice cultivar ‘Zhongjiazao 17’ and the late rice cultivar ‘Xiangwanxian 12’ were used in the experiment, and their growth durations were 109 days and 115 days, respectively. Rice seedlings were transplanted on May 5th and harvested on July 12th for the early-season rice, followed by the late-season rice, which was transplanted on July 25th and harvested on October 30th. The seedlings were 35 and 25 days old in the early and late seasons, respectively. The transplantation density was 30 hills m−2 for the early rice season and 25 hills m−2 for the late rice season.
We supplied nitrogen (N) in the form of urea, calcium superphosphate for phosphorus pentoxide (P2O5), and potassium chloride for potassium oxide (K2O) in the rice growing season. The quantity of N supplied was 74 kg ha−1 in the early rice season and 102 kg ha−1 in the late rice season. Urea was applied three times during the rice season; the ratio of tillering fertilizer to panicle fertilizer (grain fertilizer) was 70:30 in the early rice season and 50:50 in the late rice season. The quantity of P2O5 and K2O supplied was 60 kg ha−1, and the same quantity was applied in both seasons. Potassium chloride was applied twice during the rice season, 50% as basal fertilizer and 50% as tillering fertilizer. The calcium superphosphate was applied as a basal fertilizer before transplantation. Water management was performed according to the technology used for double rice cropping systems (local high-yield cultivation) (Table 4).
Table 4 Experimental design16.
Full size table

Soil chemical properties
Soil samples from the 0–20 cm soil layer were used to determine the soil chemical properties. The samples were collected during cover crop harvesting, early rice harvesting and late rice harvesting. The soil samples were air dried and the soil organic matter was determined using K2Cr2O7 and concentrated H2SO4 and heating. The soil total N was determined with the Kjeldahl method, which involved two steps: (1) the digestion of the samples to convert organic N into ({text{NH}}_{4}^{ + })–N and (2) the determination of ({text{NH}}_{4}^{ + })–N in the digest. The soil C:N ratio was calculated by dividing the SOC concentration by the TN concentration. Soil ammonium N was analyzed using indophenol blue colorimetry. Soil nitrate–N was analyzed using ultraviolet spectrophotometry.
In situ CH4 and CO2 flux measurements
During the rice growing season, in situ CH4 and CO2 flux were measured with a static chamber by circulating the gas within the chamber and pipes of an ultraportable greenhouse gas analyzer (CH4/CO2/H2O Analyzer; Los Gatos Research Corp., USA). The static chamber was a square with a side length of 50 cm and a height of 120 cm. A fluted base consistent with the static chamber was inserted in the soil in advance. On the sampling dates, daytime samples were collected from 9:00–11:00 a.m. and 15:00–17:00 p.m., and nighttime samples were collected from 19:00–21.00 p.m. The testing time in each plot was 5 min. The sampling dates were 170, 185, 199, 215, 230, 252, 268, 291, 304, 322, and 347 days after the chickens were introduced into the field. The samples were collected at intervals of 14 days, plus or minus one day if the weather forecast for a sampling date was rainy.
The temperature inside the static chamber needs to be accurately recorded at a soil depth of 3 cm. Plants (excluding the border plants) were sampled from a 0.24 m2 area of each plot on the sampling date. The plant samples were manually separated into leaf and straw and/or grains. The volume of the plant samples was measured with drainage. The effective volume in the chamber was reduced to subtract the internal plant volume from the chamber. The leaf area was determined with a leaf area meter (LI-3000A, LICOR, Lincoln, NE, USA). Lastly, the plant samples were oven-dried at 70 °C to constant weight to determine the aboveground biomass.
The CO2 (F, g m−2 day−1) and CH4 (F, mg m−2 day−1) fluxes were calculated using the following formula (Eq. 1):

$$ {text{F}} = frac{{{text{P}} times {text{V}}}}{{{text{R}} times {text{A}} times left( {{text{T}} + 273.15} right)}} times frac{{{text{dc}}}}{{{text{dt}}}}, $$
(1)

where P is the atmospheric pressure under standard conditions (101.2237 × 103 Pa); V is the effective volume in the chamber (m3), the difference between the volume of the static chamber and the volume of the plant, fan and temperature recorder; R is a gas constant (8.3144 J⋅mol−1 K−1); A is the area of the chamber cover (m2); T is the average temperature at testing time inside the chamber (°C); and dc/dt is the rate of change in the concentration of CO2 and CH4.
To accurately calculate the CO2 and CH4 fluxes in the paddy field, the daytime and nighttime CO2 and CH4 fluxes on the sampling dates were calculated using the following formulas (Eq. 2–4):

$$ {text{F}}_{{{text{daytime}}}} = {text{ S}}_{{{text{daytime}}}} times {text{M}} times left( {{text{F}}_{{1}} + {text{F}}_{{2}} } right)/{2,} $$
(2)

$$ {text{F}}_{{{text{night}}}} = {text{ F}}_{{3}} times {text{S}}_{{{text{night}}}} times {text{M,}} $$
(3)

$$ {text{F}}_{{{text{day}}}} = {text{ F}}_{{{text{daytime}}}} + {text{F}}_{{{text{night}}}} , $$
(4)

where F1, F2 and F3 represent the values at 9:00–11:00 a.m. and 15:00–17:00 p.m. on sunny days and 19:00–21:00 p.m., respectively; S is the day length (s day−1) on the sampling date; and M is the relative molecular mass of CO2 or CH4 (g mol−1).
Seasonal emissions in CO2 and CH4 were calculated using the following formula (Eq. 5):

$$ {text{T }} = {text{a}} times {1}0 times left( {mathop sum limits_{{{text{i}} = 1}}^{{text{n}}} [frac{{{text{F}}_{{text{i}}} + {text{F}}_{{{text{i}} + 1}} }}{2}left( {{text{t}}_{{{text{i}} + 1}} – {text{t}}_{{text{i}}} } right)] + frac{{{text{F}}_{{text{i}}} + {text{F}}_{{text{n}}} }}{2}} right), $$
(5)

where T (g m−2) is the total seasonal emissions, Fi and Fi+1 are the measured fluxes on two consecutive sampling days, ti+1 − ti is the number of days between the two sampling dates, 10 is the conversion coefficient from g m−2 to kg ha−1, and a is the conversion coefficient of the rice growth period (86/61 in the early season and 132/96 in the late season).
In addition, the period from early rice harvesting to late rice transplanting is 13 days. The emissions were calculated using the following formula (Eq. 6):

$$ {text{T}}_{{{text{ER}} – {text{LR}}}} = {text{ T}}_{{{text{ER}}}} /{86} times {6}.{5} + {text{T}}_{{{text{LR}}}} /{132} times {6}.{5,} $$
(6)

where TER-LR (g m−2) is the total emissions from early rice harvesting to late rice transplanting, TER and TLR are the total seasonal emissions in the early rice season and late rice season, respectively, and 86 and 132 are the number of days from sowing to harvesting in the early rice season and late rice season, respectively.
Soil microbe and dissolved carbon and nitrogen measurements
In 2014, soil was sampled from the 0–20 cm soil layer, and the sampling dates were 10, 28, 56, 74, 120, 170, 183, 199, 215, 234, 252, 268, 294, 301, 322, and 347 days after chicken grazing. Fresh soil samples were taken to determine the soil microbial carbon and nitrogen contents by chloroform fumigation-incubation and K2SO4 extraction. Soil microbial carbon (SMC, mg kg−1) = EC/0.38 and soil microbial nitrogen (SMN, mg kg−1) = EN × 0.45, where 0.33 and 0.45 are the conversion coefficients of SMC and SMN, respectively. EC and EN are the differences in organic carbon and nitrogen between fumigation and nonfumigation based K2SO4 extraction. In addition, other fresh soil samples were used to determine the soil dissolved carbon and nitrogen by K2SO4 extraction.
Yield and its components
When the rice was mature, 10 hills were sampled randomly from a 5 m2 harvest area to determine the yield components. Panicle number was counted on each hill to determine the panicle number per m2. The panicles were hand-threshed, and the filled spikelets were separated from the unfilled spikelets by submerging them in tap water. Three subsamples of 30 g of filled spikelets and 3 g of unfilled spikelets were taken to count the number of spikelets. Based on the spikelets per panicle, the grain-filling percentage (100 × filled spikelet number/total spikelet number) was determined. The grain yield was determined from a 5 m2 area in each plot and adjusted to the standard moisture content of 0.14 g H2O g−1.
Data analysis
The global warming potential (GWP) was the overall GWP of CH4 and N2O emissions per unit rice field (ha). The 100-year radiative forcing potential coefficients relative to CO2 were 25 and 298 for CH4 and N2O, respectively (IPCC, 2007). The net ecosystem exchange (NEE) was the value of Fdaytime, ecosystem respiration (Reco) was the value of Fnighttime, and gross primary production (GPP) was the sum of the NEE and Reco. The means of the indexes were organized in Excel 2016. The SD (standard deviation) of the indexes were determined by descriptive statistics with a 95% confidence interval. Analysis of variance (ANOVA) and multiple comparisons were performed using Statistix ver. 8.0 (2004) to evaluate the effects of planting cover crops and chicken grazing on the SOC, STN, C:N ratio, DOC, DON, SMN, SMC, and grain yield and its components. More

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