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The following museum acronyms are used (mostly following55):
BMNH, NHM, BM: Natural History Museum, London, formerly British Museum of Natural History (UK);
CAS: California Academy of Sciences, San Francisco (USA);
MHNC: Musée d’histoire naturelle, La Chaux-de-Fonds (CH);
MHNG: Muséum d’Histoire Naturelle, Genève (CH);
MNHN: Muséum national d’Histoire naturelle, Paris (F);
SMNS: Staatliches Museum für Naturkunde, Stuttgart (D);
UWBM The Washington State Museum of Natural History and Culture/Burke Museum, University of Washington (USA);
ZMB: Museum für Naturkunde Berlin, formerly Zoologisches Museum Berlin (D).
Study organism and taxonomic background
Acanthodactylus was first described as a subgenus of Lacerta Cuvier (sic)56, the type species is A. boskianus (Daudin, 1802). The few meristic characters described to be diagnostic for Acanthodactylus56 include: collar connected in the center but free on the sides, tempora squamosal, i.e., temporal region covered by small scales rather than larger shields, ventral scales rectangular and arranged in longitudinal rows, digits acutely fimbriate-denticulate forming “toe fringes”. Fringed toes have evolved in various shapes multiple times in lizards and in Lacertidae, and when triangular, projectional or conical as in Acanthodactylus they are commonly seen as adaptation to windblown sand substrate57.
The enigmatic Acanthodactylus guineensis is among the lesser-known species of the Lacertidae. Only limited information is available regarding the species’ morphology, habitat and distribution (see Meinig & Böhme44 for a review and references therein; and 34, 38, 45, 58). Based on one young specimen, A. guineensis was described as member of the genus Eremias Fitzinger, 1834 by Boulenger40. Boulenger does not mention the slightly projecting third row of scales around the toes and fingers (“fringed toes”) of the type specimen in his quite comprehensive description40, probably because he did not notice it in such a small (SVL 24 mm) specimen and because the projection is indeed rather minor compared to other members of the genus. This is probably also the reason why he did not place guineensis in the genus Acanthodactylus Wiegmann, 1834 but in Eremias Fitzinger, 1834 (in56).
About 30 years later, Boulenger revised the genus Eremias and divided it into five sections he assumed to be natural associations59: (1) Taenieremias Boulenger, 1918—monotypic, type species Eremias guineensis Boulenger, 1887 (currently Acanthodactylus guineensis); (2) Lampreremias Boulenger, 1918—type species Eremias nitida Günther, 1872 (currently Heliobolus nitidus); (3) Pseuderemias Boettger, 1883—type species Eremias mucronata (Blandford, 1870) (currently Pseuderemias mucronata); (4) Mesalina Gray, 1838—type species Eremias rubropunctata (Lichtenstein, 1823) (currently Mesalina rubropunctata); (5) Eremias s. str. Fitzinger, 1834—type Eremias velox (Pallas, 1771) (currently Eremias velox).
Monard47 described Eremias (Taenieremias) benuensis from Cameroon based on a few minor morphological differences compared to E. guineensis, but in 1969 E. benuensis was synonymized with E. guineensis60.
Salvador17 and one year later also Arnold18 in their respective major revisions of the genus Acanthodactylus both found that Eremias (Taenieremias) guineensis agrees with all the characteristic morphological features of Acanthodactylus with the exception of the arrangement of scales around the nostril. However, it was suggested that the E. guineensis condition with an extra suture across the area occupied by the first upper labial scale to produce a smaller first upper labial is easily derived from that found in Acanthodactylus, evidenced by BMNH 1966.430, a juvenile A. erythrurus lineomaculatus (sic) with a similar scale arrangement18. Within Acanthodactylus, both authors placed guineensis in the Western clade and in the Acanthodactylus erythrurus group which was assumed to consist of A. erythrurus, A. savignyi, A. boueti, A. guineensis, and A. blanci which was considered a subspecies of A. savignyi at the time14, 20.
Molecular data and phylogenetic analyses
We aimed at reconstructing a well-supported phylogeny of the genus Acanthodactylus using whole mitochondrial DNA sequences, assembled by means of a shotgun next generation sequencing strategy. Analyses of whole mitogenomes have been shown to resolve many nodes of the lacertid tree with high statistical support (e.g.31). We retrieved muscle tissue from a museum voucher of A. guineensis (ZFMK 59511 from Daroha, near Bobo Dioulasso, Burkina Faso43) that likely was never in contact with formalin for preservation and therefore offered good chances to obtain sufficient amounts of DNA of decent quality for sequencing. We extracted genomic DNA using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, Hilden, Germany) following the protocol provided by the manufacturer. We additionally sequenced a tissue sample of a freshly caught individual of Acanthodactylus schmidti (SB 642) from Abu Dhabi (UAE) to increase sampling of Acanthodactylus spp. in the mitogenomic tree. The lizard was euthanized by injection of an aqueous solution of benzocaine (20%) into the body cavity. Subsequently, a sample of muscle tissue was taken from the thigh, preserved in 98% Ethanol and stored in − 80 °C. Handling, euthanizing and collection of tissue samples of A. schmidti individuals was approved by the NYUAD Institutional Animal Care and Use Committee (IACUC 19-0002) and UAE No Objection Certificate (NOC 8416), and all applied methods were performed in accordance with the relevant guidelines and regulations.
Genomic DNA of SB 642 was extracted from ethanol-preserved muscle tissue using the Qiagen MagAttract HMW DNA Kit (Qiagen, Hilden, Germany) for high molecular weight DNA. We determined DNA yields on a Qubit fluorometer (Qubit, London, UK) with a dsDNA high sensitivity kit. Totals of 52 ng and 80 ng of DNA (diluted in 26 µl 10 mM Tris·Cl, 0.5 mM EDTA, pH 9.0 (AE buffer)) from A. guineensis and A. schmidti, respectively, were used for library preparation.
ZFMK 59511 libraries were prepared with NEB Ultra II FS DNA (New England Biolabs, Ipswich, MA, USA) kit as per protocol instructions with input below 100 ng. For sample SB642, linked reads were generated on a 10X Genomics Chromium following Genome reagent kits v2 instructions. Resulting libraries’ concentration, size distribution and quality were assessed on a Qubit fluorometer (Qubit, London, UK) with a dsDNA high sensitivity kit and on an Agilent 2100 Bioanalyzer (Agilent, Santa Clara, CA, USA) using a High Sensitivity DNA kit. Subsequently, libraries were normalized and pooled, and pools quantified with a KAPA Library quantification kit for Illumina platforms (Roche Sequencing, Pleasanton, CA, USA) on a ABI StepOnePlus qPCR machine (Thermo Fisher Scientific Inc., Waltham, MA, USA), then loaded on a SP flowcell and paired-end sequenced (2 × 150 bp) on an Illumina NovaSeq 6000 next generation sequencer (Illumina, San Diego, CA, USA), and a S2 flowcell for linked read library. Raw reads were deposited in the Sequence Read Archive (SRA) under BioProject ID PRJNA700414 . All mitogenome assemblies and original alignments were deposited in Figshare under  https://doi.org/10.6084/m9.figshare.13754083.v1 .
Raw FASTQ sequenced reads were first assessed for quality using FastQC v0.11.561. The reads where then passed through Trimmomatic v0.3662 (parameters ILLUMINACLIP: trimmomatic_adapter.fa:2:30:10 TRAILING:3 LEADING:3 SLIDINGWINDOW:4:15 MINLEN:36) for quality trimming and adapter sequence removal. Following the quality trimming, the reads were assessed again using FastQC. The executed workflows were performed using BioSAILs63.
For SB 642 (A. schmidti), we received 436,370,000 single-end reads (average read length 139.5 bases (b)) with a raw coverage of 29.29X and a scaffold N50 of 21.12 kilobases (kb). For the MITObim analysis, read number for SB 642 was randomly reduced to 1,000,000 reads (360,463,035 b) using the awk command. For ZFMK 59511 (A. guineensis), we received 57,936,345 paired-end reads (average read length 151 b, raw coverage ~ 10x). Subsequently, the two sets of reads were converted to interleaved format. We used the quality filtered reads to assemble the mitogenomes of A. guineensis and A. schmidti using an iterative mapping strategy in MITObim v. 1.9.164. We used the Acanthodactylus aureus mitogenome (GB accession number xxxxx; assembly method is provided in the next paragraph) as seed for both samples; this rendered an initial mapping of a conserved region from a more distantly related individual unnecessary64. We therefore applied the –quick option in MITObim and iterations were run until no additional reads could be incorporated into the assembly (14 in A. guineensis, eight in A. schmidti).
We also assembled complete or nearly complete mitogenomes for additional twelve species of Gallotiinae and Eremiadini using anchored hybrid enrichment sequence data from Garcia-Porta et al.6 (see Supplementary methods in6 for extraction protocol and sequencing methods and Supplementary Table S1 online in6) with the Podarcis muralis complete mitogenome (GB accession number NC_011607) as reference sequence. The raw data of each individual was quality filtered using Trimmomatic v0.3662 (parameter MINLEN:45) and assembled using MITObim v. 1.9.1. All multiple alignment files generated in the final MITObim iteration were imported to Geneious R11 (https://www.geneious.com) to check for assembly quality and coverage.
The resulting assemblies were annotated with MITOS65 using defaults settings with the vertebrata database as a reference. The annotated assemblies were imported into PhyloSuite66 together with existing mitogenomic sequences of Lacertidae, and Blanus cinereus (Amphisbaenia; outgroup) available in GenBank (as of July 2020). Details of all mitogenomic sequences included in this study and the corresponding GenBank accession numbers are provided in Supplementary Table S1 online. Using PhyloSuite, we exported all protein coding sequences plus the two rRNAs. The resulting files were aligned with MAFFT67 using “auto” settings. Alignments of coding sequences were refined with MACSE68 to account for open reading frame structure during the alignments. The protein coding and rRNAs alignments were finally concatenated into a single alignment delimiting partitions by marker and, for the protein-coding genes, by codons within markers. Mitogenome phylogenies were inferred using a maximum likelihood approach with IQ-TREE v. 1.5.4 software69 and Bayesian inference (BI) analysis with MrBayes 3.270.
For the maximum likelihood approach, best-fitting partitioning schemes and substitution models were selected based on the Akaike Information Criterion (AIC) using the heuristic algorithms implemented in the MFP + MERGE option (Supplementary Table S3 online). After ML inference, branch support was assessed with 1000 ultrafast bootstrap replicates. As the alignments used for phylogenetic inference contained more than one sequence for some species, these terminals in the resulting tree were collapsed a-posteriori for aesthetic reasons using FigTree v1.4 (as depicted in Fig. 1). An uncollapsed version of this tree is presented in Supplementary Fig. S1 online. For the BI analysis, the best-fitting partition/substitution model scheme, as selected by the ModelFinder algorithm in iQTree (Supplementary Table S4 online), was implemented with MrBayes 3.270. Results of two independent runs of 10 million generations, each comprising four Markov Chains (three heated and one cold), were sampled every 1000 generation. Chain mixing and stationery was assessed by examining the standard deviation of split frequencies and by plotting the –lnL per generation using Tracer 1.5 software71. Samples corresponding to the initial phase of the Markov chain (25%) were discarded as burn-in and the remaining results were combined to obtain a majority rule consensus tree and the respective posterior probabilities of nodes.
An additional, more comprehensive, alignment was produced containing all currently known Acanthodactylus species to obtain a higher resolution perspective on the phylogenetic placement of the target species. Since no nuclear data was available for A. guineensis, we extracted from the newly obtained sequences the four best represented mitochondrial gene fragments across Lacertidae, as compiled by Garcia-Porta et al.6, despite the known shortcomings of partial mitochondrial datasets to resolve lacertid relationships9, 72, 73. The corresponding sequences of the ribosomal RNAs (12S, 16S), COB, and NADH-dehydrogenase subunit (ND4) were later added to the alignment of6 using –add and –keeplength options in MAFFT (Supplementary Table S5 online).
Species distribution modeling
As basis for characterizing the species’ bioclimatic envelopes, we compiled an updated distribution map for A. guineensis and A. boueti including new discoveries in museum collections from the current study (see Supplementary Material online), all known records from the literature16, 34, 37, 38, 40, 42, 44, 45, 47, 58, 60, 74,75,76,77 and from the Global Biodiversity Information Facility (GBIF78). We provide all coordinates as latitude (decimal degrees), longitude (decimal degrees). Due to the general scarcity of records for A. guineensis and A. boueti, we added two localities for A. guineensis (12.5°, − 2.5°; 7.5°, 13.5°) and one locality for A. boueti (− 2.5°, 7.5°) published in Trape et al.45 that were not found on GBIF or in other publications, by extracting the center coordinates of the respective 1 × 1 degree grid. These additional localities have coordinates with very low accuracy and the corresponding environmental variables extracted from a 1 × 1 km resolution grid (30 arc-seconds; see below) therefore have high degrees of uncertainty. When examined, the majority of values fell within the total range for the respective environmental variable (the two exceptions are mentioned in the Results section), we consequently decided to keep them for our analysis. We reduced the final locality dataset to one record per km2 (the resolution of the environmental input data, see below) to avoid pseudoreplication using the R package spThin79. We then tested whether the remaining presence points are randomly dispersed or clustered using the Nearest Neighbor Index NNI in the R packages sp80 and spatialEco81. NNI was 1.4 for A. guineensis and 1.2 for A. boueti, indicating that the filtered point dataset consists of randomly distributed points which are not spatially autocorrelated.
As environmental parameters we downloaded spatial layers of the Terrestrial Ecoregions of the World (TEOW)82, global bioclimate and elevation layers39, data on potential evapotranspiration83 and global aridity index84 with a spatial resolution of 30 arc-seconds (~ 1 km2 near the equator). Although the climate datasets are interpolated from data from weather stations much farther apart from one another and are consequently not measurements of actual environmental conditions, they were rigorously cross-validated with observed data (including satellite data) during the development of the dataset and generally showed high correspondence with observations (especially temperature variables)39. Since for many applications such as our species distribution models data at high spatial resolution (i.e., 1 × 1 km) are preferable over lower resolution to capture variation for example across steep climate gradients in mountains39 we chose the 30 arc-seconds dataset over the 5 arc-minutes dataset (~ 9 × 9 km near the equator) for the ecoregions and climate datasets. In addition, we downloaded spatial layers with forest and grassland/scrub/woodland cover from the harmonized soil database (only available in 5 arc-minutes spatial resolution85). We downscaled the forest and grassland/scrub/woodland dataset to 30 arc-seconds resolution despite the resulting slight inaccuracy, which we kept in mind during interpretation but did not consider relevant for the vegetation datasets.
In order to compare the prevailing climate within the distribution range of A. guineensis and A. boueti to all other species of the genus Acanthodactylus we downloaded all available records of the other currently known Acanthodactylus species from GBIF. We cleaned the downloaded datasheet by deleting all entries with missing data for either latitude, longitude, species epithet, or all of those, and removed duplicates or spelling mistakes. We further removed all GBIF entries with imprecise coordinates, i.e. records with coordinates that when plotted landed just outside of coastal areas in the ocean instead of on land (usually coordinates with only two decimals). In addition, we added several curated locality data from Garcia-Porta et al.6 as available from Figshare (https://doi.org/10.6084/m9.figshare.8866271.v1/). Following Tamar et al.14 we treated A. lineomaculatus as a junior synonym of A. erythrurus and changed the respective records in our database accordingly. For species that did not have records published on GBIF we added further locality data from the literature (Supplementary Table S6 online). The final database contained 4286 records from all 44 currently recognized species of Acanthodactylus. We acknowledge that our database is by no means complete, however, we wanted to follow the most conservative approach and use only records that are supported by a voucher specimen and can be traced back using published databases. We trust that our record list covers a nearly complete representation of the ecological conditions inhabited by all currently accepted Acanthodactylus species.
Using the bioclim dataset39 we plotted annual mean temperature (bio1) and annual precipitation (bio12) for all locality records in our Acanthodactylus species database and compared the respective data for A. guineensis and A. boueti with its congeners. We developed species distribution models using Maxent 3.4.146 for A. guineensis and A. boueti. Maxent applies the maximum entropy principle86 for model fitting under the basic premise that the estimated species distribution deviates from a uniform distribution as minimally as required to explain the observations46. We used the cleaned datasets of A. guineensis and A. boueti with only one record per km2 as input presence locality data. We extracted environmental data for all presence sites from the 19 bioclim variables (bio1-19), elevation (alt), aridity index (AI), percentage cover of forest (forest) and grassland, scrub and woodland (grass) per grid cell, as well as four parameters comprising potential evapotranspiration (PET): PET of the wettest, driest, warmest and coldest quarter of the year (PETwet, PETdry, PETwarm, PETcold). To avoid issues resulting from correlated parameters we reduced the number of environmental predictors by performing multicollinearity analyses using the Variance Inflation Factor (VIF) with a threshold  More

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Physicochemical properties of water
Physicochemical properties of water and associated environmental factors according to the flow directions of surface flow wetlands are shown in Table 1. Analysis of variance for indexes with p values of less than 0.05 showed that all indexes exhibited large variations during different months.
Table 1 Physicochemical characteristics of the surface-flow constructed wetlands in each unit.
Full size table

Some indexes exhibited large variations during different months according to the flow directions of surface flow wetlands. For example, DO was first reduced and then increased in May, but increased in August and showed differences compared with that in May and October. The salinity was first reduced and then increased in May but then remained stable from August to October. Some indexes showed similar changes according to the flow directions of surface flow wetlands. For example, ORP showed an initial decrease followed by an increase. At the same time, pH, SpCond, TDSs, and TN showed reduced variability over time. The changes in temperature were minimal, although the temperature was higher in the summer and autumn than in the spring.
Surface flow wetlands are in direct contact with the environment and are greatly influenced by outside environmental factors17. Thus, most physicochemical factors of the water samples showed large variability.
Denitrifying bacteria diversity and abundance
For 10 samples from different seasons showing 97% similarity in clustering analysis, the numbers of OTUs differed in May, August, and October (575, 869, and 741, respectively), and the Fig. 2 showed that the denitrifying bacterial abundance indexes (ACEs) were 686.8, 996.2, and 887.3 in May, August, and October, respectively. Additionally, the Shannon-Weiner indexes (H′) were 3.718, 4.303, and 4.432, respectively, indicating that the abundance tended to increase initially, followed by a decrease, and diversity tended to increase. The different seasons affected both the denitrifying bacteria abundance and diversity. Abundance was the largest in August, but its diversity was lower than that in October. These data suggested that the main species became dominant during August, affecting the structure of the denitrifying bacteria.
Figure 2

Biodiversity and abundance of the surface-flow constructed wetland in each unit.

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For different processing units, the abundance and diversity of denitrifying bacteria varied slightly; both the ACE and H′ index showed low variability. The units F, H, and J showed greater declines than the initial values. In May, the ACE index peaked, with a value of 841 at location I. In August, the ACE index peaked at location E (1251), and that in October peaked at location G (1042). In different months, denitrifying bacteria abundances showed similar changes. Because bacterial diversity in the flowing water and static water were affected by different factors, the surface flow wetlands will be susceptible to various factors, and the bacterial community interactions with internal and external environmental factors will be important for bacterial survival18. Additionally, the number of constructed wetland bacteria decreases as the depth and distance increases19,20, suggesting that denitrifying bacteria may be affected by physical and chemical indicators of changes in water.
Community structure of denitrifying bacteria
Similar OTUs (97% similarity) were identified by sequencing. Database analysis of sequence alignment results revealed that there were many bacteria in the environmental samples that could not be cultivated but that showed high similarity; thus, the denitrifying bacteria were mostly present in the surface flow wetlands and were not cultured. Figure 3 shows statistical analysis of the denitrifying bacterial categories in a histogram format. During the different months, OTUs mainly belonged to seven genera: unclassified bacteria (37.12%), unclassified Proteobacteria (18.16%), Dechloromonas (16.21%), unranked environmental samples (12.51%), unclassified Betaproteobacteria (9.73%), unclassified Rhodocyclaceae (2.14%), Rhodanobacter (1.51%), and other genera (2.62%, representing less than 1% each). Several genera have also been found in surface flow wetlands21 and other types of constructed wetlands22,23,24, albeit with different proportions.
Figure 3

Community structure of the surface-flow constructed wetland in each unit.

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The same processing units showed different denitrifying bacterial community structures during different seasons and were always changing. Unclassified bacteria showed a greater weight during May for the A processing unit, although its weight was lower than that of Dechloromonas in August. In October, unclassified bacteria had become the most dominant group, and the proportion of Dechloromonas was extremely low. For the B processing unit, from May to October, the proportion of Dechloromonas was decreased, and the proportion of unclassified Proteobacteria was increased, overtaking Dechloromonas. For the C and D processing units, unclassified Proteobacteria were dominant in May, and unclassified bacteria were dominant in August and October. For the E, G, H, I, and J processing units, unclassified bacteria were dominant at all time points. For the F processing unit, the bacterial groups were similar to those of the B processing unit, with proportion of Dechloromonas decreasing and the proportion of unclassified bacteria increasing in October.
Figure 4 shows the means and variances of denitrifying bacteria genus proportions among different processing units and seasons. The means and variances of the dominant genus were large at the same time during different seasons. Thus, the dominant genus often determined the changes in denitrifying bacteria community structures during different seasons in the same unit. However, the greatest variance was observed in the genus Dechloromonas, which was the second most dominant genus in May. This suggested that this genus showed greater changes in different processing units than others. In August, the largest variances were observed in the genus Dechloromonas and in unclassified Proteobacteria, which had lower means than unclassified bacteria. Similar results were observed in October. The largest variance was observed in unclassified Proteobacteria, indicating that the denitrifying bacterial community structures were affected by the second dominant genus over time in the different processing units.
Figure 4

Mean and variance of different denitrifying bacterial taxa in the surface-flow constructed wetland (May, August, and October).

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For the denitrifying bacteria community structures in different months, we used nonmetric multidimensional scaling to determine the similarities between different processing units during the same months. As shown in Fig. 5, in May, A, B, C, and E showed high similarity, whereas other samples were more dispersed. The distances between D and H and between G and I were shorter than the other distances. F and J were alone in a group. Sample distributions were concentrated in August; C, D, E, F, G, H, and I were relatively similar, and D and E showed maximum similarity. A and B showed some similarity. In contrast, J was distinct. In October, distributions were more dispersed, and the distances between two points were not relatively similar, whereas the differences between the various processing units were higher.
Figure 5

Nonmetric multidimensional scaling map (May, August, and October).

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Relationships between denitrifying bacteria and environmental factors
Next, we carried out PCA analysis to determine the main factors affecting denitrifying bacteria. After maximum variance orthogonal rotating (p = 0.05), there were two principal component eigenvalues that were greater than the average. The two top principal components contributed to 53.9% and 28.7% of the variance. The first principal component mainly reflected SpCond, TDSs, ORP, and salinity (factor loading was 0.409, 0.403, 0.398, and 0.403, respectively), and the second principal component reflected DO, TN, pH, and temperature (factor loading was 0.449, 0.449, 0.465, and 0.446, respectively). The load distribution characteristics of different environmental factors showed that the surface flow wetlands were affected by the main environmental factors, including temperature, SpCond, DO, pH, ORP, and TN (Fig. 6).
Figure 6

PCA of various environmental factors in the surface-flow constructed wetlands.

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Table 2 shows bacterial abundance indexes and H′ index for different external environmental factors, as analyzed by Pearson correlation analysis. The results showed that the bacterial abundance was strongly correlated with temperature, DO, and pH, and H′ was strongly correlated with all parameters except TN.
Table 2 Relationships between biodiversity and environment factors.
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RDA was performed (Fig. 7) for analysis of community distributions and the relationships among environmental factors. For screening of the physicochemical factors of water and the proportions of denitrifying bacterial genera, standardization to center (Monte Carlo permutation) tests were used, and refinement of the information extracted from the first and second axes showed that the total explained variance rate was 80.94%. The results showed that all denitrifying bacterial genera were greatly affected by environmental factors, including temperature and pH, and that the effects of SpCond and ORP were similar. The predominance of unclassified bacteria and unclassified Proteobacter could be explained by positive correlations with temperature, pH, ORP, and SpCond and negative correlations with TN and DO. Dechloromonas showed the opposite trends. In contrast, unranked environmental samples were similar to unclassified Betaproteobacteria, with positive correlations for temperature and pH but negative correlations for TN and DO.
Figure 7

Relationships between denitrifying bacterial community structures and environment factors.

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Denitrifying bacterial diversity is affected by water nutrient elements and other environmental factors. Most denitrifying bacteria were heterotrophic bacteria. In this study, the autotrophic denitrifying bacteria Dechloromonas accounted for a large proportion in each processing unit25,26,these bacteria can accumulate phosphate and exhibit denitrification activity, partly explaining the lack of TOC removal in association with the observed TN removal. The SpCond of the water reflected its salinity and could be explained by positive correlations with a high proportion of unclassified Proteobacteria. However, SpCond was not generally correlated with denitrifying bacterial abundance. Our results showed that the water SpCond in surface-flow constructed wetlands affected salinity-related denitrifying bacteria but did not affect other denitrifying bacteria. The ORP was positively correlated with denitrifying bacterial genera that were suitable for survival in a strong oxidizing environment, such as unclassified Proteobacteria.
Different physical and chemical properties can influence the structure of the bacterial community owing to the influence of different species on the living environment27,28. In this study, we assessed environmental factors that differed according to season and showed that denitrifying bacteria varied according to some environmental parameters. A comprehensive analysis of the trend of physical and chemical properties of water showed that all parameters except DO and salinity were not highly affected by season and that the trend of the abundances of denitrifying bacteria communities did not change with season along the flow direction of different processing units. However, environmental indicators have a more significant impact on different denitrifying bacteria, which also changes the diversity of denitrifying bacteria community. Accordingly, these results, combined with prediction models of the effects of environmental factors on nitrogen and phosphorus29,30,31, could be used to predict changes in the denitrifying bacterial community structure. More

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Radiation experiments
We experimentally quantified radiolytic hydrogen (H2) production in (i) pure water, (ii) seawater, and (iii) seawater-saturated sediment. We irradiated these materials with α- or γ-radiation for fixed time intervals and then determined the concentrations of H2 produced. Sediment samples were slurried with natural seawater to achieve a slurry porosity (φ) of ~0.83, which is the average porosity of abyssal clay in the South Pacific Gyre34. The seawater source is described below. To avoid microbiological uptake of radiolytic H2 during the course of the experiment, seawater and marine sediment slurries were pre-treated with HgCl2 (0.05% solution) or NaN3 (0.1% wt/vol). To ensure that addition of these chemicals did not impact radiolytic H2 yields, irradiation experiments with pure water plus HgCl2 or NaN3 were also conducted. HgCl2 or NaN3 addition had no statistically significant impact on H2 yields5,6,10.
Experimental samples were irradiated in 250 mL borosilicate vials. A solid-angle 137Cs source (beam energy of 0.67 MeV) was used for the γ-irradiation experiments at the Rhode Island Nuclear Science Center (RINSC). The calculated dose rate for sediment slurries was 2.19E−02 Gy h−1 accounting for the (i) source activity, (ii) distance between the source and the samples, (iii) sample vial geometry, and (iv) attenuation coefficient of γ-radiation through air, borosilicate, and sediment slurry. 210Po (5.3 MeV decay−1)-plated silver strips with total activities of 250 μCi were used for the α-irradiation experiments. For α-irradiation of each sediment slurry, a 210Po-plated strip was placed inside the borosilicate vial and immersed in the slurry. Calculated total absorbed doses were 4 Gy and 3 kGy for γ-irradiation and α-irradiation experiments, respectively.
The settling time of sediment grains in the slurries (1 week) was long compared to the time span of each experiment (tens of minutes to an hour for α-experiments, hours to days for γ-experiments). Therefore, we assumed that the suspension was homogenous during the course of each experiment.
H2 concentrations were measured by quantitative headspace analysis via gas chromatography. For headspace analysis, 30 mL of N2 was first injected into the sample vial. To avoid over-pressurization of the sample during injection, an equivalent amount of water was allowed to escape through a separate needle. The vials were then vigorously shaken for 5 min to concentrate the H2 into the headspace. Finally, a 500-μL-headspace subsample was injected into a reduced gas analyzer (Peak Performer 1, PP1). The reduced gas analyzer was calibrated using a 1077 ppmv H2 primary standard (Scott-Marrin, Inc.). A gas mixer was used to dilute the H2 standard with N2 gas to obtain various H2 concentrations and produce a five-point linear calibration curve (0.7, 2, 5, 20, and 45 ppm). H2 concentrations of procedural blanks consisting of sample vials filled with non-irradiated deionized 18-MΩ water were also determined. The concentration detection limit obtained using this protocol was 0.8–1 nM H2. Relative error was less than 5%. Radiation experiments were performed at a minimum in triplicate.
Sample selection and experimental radiolytic H2 yields, G(H2)
Millipore Milli-Q system water was used for our pure-water experiments. For seawater experiments, we used bottom water collected in the Hudson Canyon (water depth, 2136 m) by RV Endeavor expedition EN534. Salinity of North Atlantic bottom water in the vicinity of the Hudson Canyon (34.96 g kg−1) is similar to that of mean open-ocean bottom water (34.70 g kg−1)44,45.
The 20 sediment samples used for the experiments were collected by scientific coring expeditions in three ocean basins (expedition KN223 to the North Atlantic46, expedition KN195-3 to the Equatorial and North Pacific47, International Ocean Discovery Program (IODP) Expedition 329 to the South Pacific Gyre34, MONA expedition to the Guaymas Basin48, expedition EN32 to the Gulf of Mexico49, and expedition EN20 to the Venezuela Basin50). To capture the dominant sediment types present in the global ocean, we selected samples typical of five common sediment types [abyssal clay (11 samples), nannofossil-bearing clay or calcareous marl (2 samples), clay-bearing diatom ooze (3 samples), calcareous ooze (2 samples), and lithogenous sediment (2 samples)]. The locations, lithological descriptions, and mineral compositions of the samples are given in Supplementary Tables 1,  2,  3, and Supplementary Fig. 1. Additional chemical and physical descriptions of the sediment samples used in the radiation experiments can be found in the expedition reports for the expeditions on which the samples were collected34,46.
Energy-normalized radiolytic H2 yields are commonly expressed as G(H2)-values (molecules H2 per 100 eV absorbed)1. As shown in Supplementary Fig. 2, for all irradiated samples (pure water, seawater, and marine sediment slurries), H2 production increased linearly with absorbed α- and γ-ray-dose. We calculated G(H2)-values for each sample and radiation type (α or γ) as the slope of the least-square regression line of radiolytic H2 concentration versus absorbed dose (Supplementary Fig. 2). The error on the yields is less than 10% for each sample. G(H2)-values for each sample and radiation type (α or γ) are reported in Supplementary Table 3.
Although radiolytic OH• is known to react with dissolved organic matter51, total organic content does not appear to significantly impact radiolytic H2 production, since the most organic-rich sediment (e.g., Guaymas Basin and Gulf of Mexico sediment) did not yield particularly high H2 (Supplementary Table 3).
Calculated radiolytic production rates of H2 and oxidants in the cored sediment of individual sites
We calculated radiolytic H2 production rates (PH2, in molecules H2 cm−3 yr−1) for the cored sediment column at nine sites with oxic subseafloor sediment in the North Pacific, South Pacific, and North Atlantic; and seven sites with anoxic subseafloor sediment in the Bering Sea, South Pacific, Equatorial Pacific, and Peru Margin (see Supplementary Fig. 3 for site locations). For these calculations, we used the following equation from Blair et al.2:

$$P_{{mathrm{H}}_2} = {sum} A _{{mathrm{m}},i}rho left( {1 – {{upvarphi}} } right)E_i{mathrm{G}}({mathrm{H}}_2)_i$$
(1)

where i is alpha, beta, or gamma radiation; Am is radioactivity per mass solid; φ is porosity; ρ is density solid; (E_i) is decay energy; and ({mathrm{G}}({mathrm{H}}_2)_i) is radiolytic yield.
We calculated radiolytic oxidant production rates for these sediment columns from the H2 production rates. Because H2 production and oxidant production are stochiometrically balanced in water radiolysis [2H2O → H2 + H2O2], the calculated radiolytic H2 production rates (in electron equivalents) are equal to radiolytic oxidant production rates (in electron equivalents).
The in situ γ- and α-radiation dosages in marine sediment are, respectively, 13 and 15 orders of magnitude lower than the dosage used in our experiments. Because the measured G(H2) for pure water in our γ-irradiation experiment (dose rate = 2.19E-02 Gy h−1) is statistically indistinguishable from previously published G(H2) values at much higher dose rates (ca. 1.00E+3 Gy h−1)5, we infer that the γ-irradiation G(H2) value is constant with dose rate over five orders of magnitude. Similarly, our experimental pure water H2 yields following α-particle irradiation from a 210Po-source (dose rate of 2.55E+03 Gy h−1) are indistinguishable from the yield obtained by Crumière et al.6 [G(H2) = 1.30 ± 0.13] for air-saturated deionized water exposed to a cyclotron-generated He2+ particle beam at higher dose rate (dose rate 1.62E+05 Gy h−1). The close similarity in H2 yields obtained in both experiments implies that (i) radiolytic H2 yield from α-particle irradiation is identical to that from cyclotron-generated He2+ particle irradiation, and (ii) this yield is constant over a two-orders-of-magnitude range dose rate. Therefore, we use our experimentally determined α- and γ-irradiation G(H2) values for the low radiation dose rate found in the subseafloor. Because the G(H2) of β irradiation has not been experimentally determined for water-saturated materials, we assume that the G(H2) of β-radiation matches the G(H2) of γ-radiation for the same sediment types. In pure water, their G(H2) values differ by only 17%1. Because β radiation, on average, contributes only 11% of the total radiolytic H2 production from the U, Th series and K decay in marine sediment, these estimates of total H2 production differ by only 2–5% relative to estimates where the G(H2) of β radiation is assumed equal to that for pure water or for α radiation of the same sediment types.
To calculate H2 production rates for the entire sediment column at seven South Pacific sites and two North Atlantic sites, we measured downcore sediment profiles of U, Th, and K (i) 187 sediment samples from IODP Expedition 329 Sites U1365, U1366, U1367, U1368, U1369, U1370, and U137134,52, and (ii) 40 samples from KN223 expedition Sites 11 and 12 (ref. 46). Total U and Th (ppm) and K2O (wt%) for these sites are reported in the EarthChem SedDB data repository. We measured U, Th, and K abundances using standard atomic emission and mass spectrometry techniques (i.e. ICP-ES and ICP-MS) in the Analytical Geochemistry Facilities at Boston University. Sample preparation, analytical protocol, and data are reported in Dunlea et al.52. The precision for each element is ~2% of the measured value, based on three separate digestions of a homogenized in-house standard of deep-sea sediment.
To calculate H2 production rates for the sediment columns at North Pacific coring Sites EQP10 and EQP11 (ref. 47), we used radioactive element content data from Kyte et al.53, who measured chemical concentrations at high resolution in bulk sediment in core LL44-GPC3. Because Site EQP11 was cored at the same location as LL44-GPC3 (ref. 53) and the sediment retrieved at all three sites is homogeneous abyssal clay, we assume the radioactive element abundances measured in core LL44-GPC3 to be representative of Sites EQP10 and EQP11 (ref. 47). Calculated radiolytic H2 production rates for South Pacific sites are listed in Supplementary Table 4 and for North Atlantic and North Pacific sites in Supplementary Table 5.
For Bering Sea Sites U1343 and U1345 (ref. 54), sedimentary U, Th, and K content measurements are unavailable. Since sediment recovered at these two sites is primarily siliciclastic with a varying amount of diatom-rich clay, we use U, Th, and K concentration values reported for upper continental crust by Li and Schoonmaker for these Bering Sea sites55. Finally, we calculate downhole radiolytic H2 production rates for ODP Leg 201 Sites 1225, 1226, 1227, and 1230 (ref. 35). Sediment compositions for these sites include nannofossil-rich calcareous ooze (Site 1225), alternation of nannofossil (calcareous) ooze and diatom ooze (Site 1226), and siliciclastic with diatom-rich clay intervals (Sites 1227 and 1230). Because sedimentary U, Th, and K measurements are not available for Leg 201 sites, we used average U, Th, and K concentration values measured in North Atlantic46 and South Pacific Sites34,52 with corresponding lithologies.
We use isotopic abundance values reported in Erlank et al.56 to calculate the abundance of 238U, 235U, 232Th, and 40K from the measured ICP-MS values of total U, Th, and K concentration. We then converted radionuclide concentrations to activities using Avogadro’s number and each isotope’s decay constant2. We refer to Blair et al. for a detailed explanation of activities and radiolytic yield calculations2.
Calculation of global radiolytic H2 and oxidant production rates in marine sediment
We calculated global radiolytic H2 production in ocean sediment by applying Eq. (1) (ref. 2) globally. As with the rates at individual sites, we calculated global radiolytic oxidant production (in electron equivalents) from global H2 production and the stochiometry of water radiolysis [2H2O → H2 + H2O2].
Our global radiolytic H2 production calculation spatially integrates calculations of sedimentary porewater radiolysis rates that are based on (i) our experimentally constrained radiolytic H2 yields for the principal marine sediment types, (ii) measured radioactive element content of sediment cores in three ocean basins (North Atlantic46, North Pacific53, and South Pacific34,52), and (iii) global distributions of sediment lithology57, sediment porosity58, and sediment column length59,60.
To generate the global map of radiolytic H2 production, we created global maps of seafloor U, Th, and K concentrations, density, G(H2)-α values, and G(H2)-γ-and-β by assigning each grid cell in our compiled seafloor lithology map (Supplementary Fig. 4) its lithology-specific set of input variables (Supplementary Table 6). Because our model assumes that lithology is constant with depth, U, Th, and K content, grain density, and G(H2)-values are constant with depth.
The G(H2)-values (α, β, and γ radiation), radioactive element content (sedimentary U, Th, and K concentration), density, porosity, and sediment thickness are determined as follows.
Radiolytic yield [G(H2)] for α,β-&-γ radiation
Radiolytic yields for the main seafloor lithologies are obtained by averaging experimentally derived yields for the respective lithologies (Supplementary Table 6). We assume that G(H2)-β values equal G(H2)-γ values.
Sediment lithology
For these calculations of radiolytic chemical production, we generally used seafloor lithologies and assumed that sediment type is constant with sediment depth. For seafloor lithology, the geographic database of global bottom sediment types57 was compiled into five lithologic categories: abyssal clay, calcareous ooze, siliceous ooze, calcareous marl, and lithogenous (Supplementary Fig. 4). Some areas of the seafloor are not described in the database57. These include (i) high-latitude regions (as the seafloor lithology database extends from 70°N to 50°S)57 and (ii) some discrete areas located along continental margins (e.g., Mediterranean Sea, Timor Sea, South China Sea, Supplementary Fig. 4). We used other data sources to identify seafloor lithologies for these regions. We added an opal belt (siliceous ooze) in the Southern Ocean between 57°S and 66°S61,62. The geographic extent of this opal belt was based on DeMaster62 and Dutkiewicz et al.61. We defined the areas of the seafloor from 50°S to 57°S, from 66°S to 90°S, and in the Arctic Ocean as mostly composed of lithogenous material, based on (i) drillsite lithologies in the Southern Ocean [ODP: Site 695 (ref. 63), Site 694 (ref. 63), Site 1165 (ref. 64), Site 739 (ref. 65)], the Bering Sea and Arctic Ocean [International Ocean Discovery Program (IODP): Sites U1343 and U1345 (ref. 54), Site M0002 (ref. 66), ODP: Site 910 (ref. 67), Site 645 (ref. 68)] and between 50°S and 57°S [Deep Sea Drilling Project (DSDP): Site 326 (ref. 69), Ocean Drilling Program (ODP): Site 1138 (ref. 70), Site 1121 (ref. 71)], and Dutkiewicz et al.61.
In the North and South Atlantic, sediment type can be very different at depth than at the seafloor. For these regions, we departed from our assumption that sediment lithology is the same at depth as at the seafloor. Subseafloor lithologies at ODP Sites [1063 (ref. 72), 951 (ref. 73), 925 (ref. 74), and 662 (ref. 75)] and IODP Sites [U1403 (ref. 76) and U1312 (ref. 77)] indicate that sediment in the Atlantic Ocean basin is generally 30–90% biogenic carbonate content and detrital clay78, even where the seafloor lithology is abyssal clay57. Therefore, regions in the Atlantic Ocean described as abyssal clay in the seafloor lithology database57 were characterized as calcareous marl for our calculations (Supplementary Fig. 4). Because abyssal clay catalyzes radiolytic H2 production at a higher rate than calcareous marl, this characterization may underestimate production of radiolytic H2 and radiolytic oxidants in these Atlantic regions.
Radioactive element content
For four of the five lithologic types in our global maps (abyssal clay, siliceous ooze, calcareous ooze, and calcareous marl), we average U, Th, and K concentrations from sites in the North Atlantic46, North Pacific53, and South Pacific34,52. The average U, Th, and K concentration values are consistent with data reported in Li and Schoonmaker55 for the characteristic U, Th, and K content found in abyssal clay and calcareous ooze. For lithogenous sediment, we use U, Th, and K concentration values reported for upper continental crust by Li and Schoonmaker55. Lithology-specific radioactive element values are given in Supplementary Table 6 and used to calculate Am,i in Eq. (1).
Density
Characteristic density values for calcite, quartz, terrigenous clay, and opal-rich sediment were extracted from the Proceedings of the Integrated Ocean Drilling Program Volume 320/321 and are assigned to calcareous ooze, lithogenous sediment, abyssal clay, and siliceous ooze, respectively79.
Global porosity
For global porosity, we use a seafloor porosity data set by Martin et al.58 and accounted for sediment compaction with depth by using separate sediment compaction length scales for continental-shelf (0–200 m water depth; c0 = 0.5 × 10−3), continental-margin (200–2500 m; c0 = 1.7 × 10−3), and abyssal sediment ( >3500 m; c0 = 0.85 × 10−3)80,81. Once the porosity was 0.1%, the depth integration was halted.
Global sediment thickness
We calculated global depth-integrated radiolytic H2 production by summing the seafloor production rates over sediment depth in one-meter intervals (Fig. 3 in main text). Sediment thickness is from Whittaker et al., supplemented with Laske and Masters where needed82,83.
Ocean depth
For porosity calculations, water depths were determined using the General Bathymetric Chart of the Oceans84, resampled to a 5-arc minute grid, i.e. the resolution of the Naval Oceanographic Office’s Bottom Sediment Type (BTS) database “Enhanced dataset”57.
Dissolved H2 concentration profiles
H2 concentrations from South Pacific Sites U1365, U1369, U1370, and U1371, and the measurement protocol, are described in ref. 1. H2 concentrations from North Atlantic KN223 Sites 11, 12, and 15, and North Pacific Site EQP11 were determined using the same protocol and are posted on SedDB (see “Data availability”). The detection limit for H2 ranged between 1 and 5 nM H2, depending on site, and is displayed as gray vertical lines in Fig. 2 of the main text. H2 concentrations for Equatorial Pacific Site 1225 and Peru Trench Site 1230 were measured by the “headspace equilibration technique”, which measures steady-state H2 levels reached following laboratory incubation of the sediment samples85,86.
For comparison to these measured H2 concentrations, we use diffusion-reaction calculations to quantify what in situ H2 concentrations would be in the absence of H2-consuming reactions. The results of these calculations are represented as solid circles (•) in Fig. 2 of the main text. Temporal changes in H2 concentration due to diffusive processes and radiolytic H2 production in situ are expressed by Eq. (2):

$$frac{{partial {mathrm{H}}_2(x,t)}}{{partial t}} = frac{D}{{varphi F}}frac{{partial ^2{mathrm{H}}_2(x,t)}}{{partial x^2}} + P(x)$$
(2)

with
D: the diffusion coefficient of H2(aq) at in situ temperature
(varphi): porosity
F: formation factor
x: depth
Z: sediment column thickness
({mathrm{H}}_2): hydrogen concentration
P: radiolytic H2 production rate
t : time.
With constant radiolytic H2 production, P(x) = P with depth,
and at steady-state,

$$frac{{partial ^2{mathrm{H}}_2(x)}}{{partial x^2}} = – frac{{Pvarphi F}}{D}.$$
(3)

We integrate Eq. (3) over the length x twice,

$${mathrm{H}}_2(x) = – frac{1}{2}frac{{Pvarphi F}}{D}x^2 + Ax + B$$
(4)

where A and B in Eq. (4) are constants of integration. We use two boundary conditions to derive the value of these constants.
Boundary condition 1: concentration of H2 at the sediment-water interface, x = 0, is zero due to diffusive loss to the overlying water column.
Boundary condition 2: concentration of H2 at the basement–sediment-water interface, x = Z, is zero due to diffusive loss to the underlying basement.
With these boundary conditions, (A = frac{1}{2}frac{{Pvarphi F}}{D}Z) and B = 0
and

$${mathrm{H}}_2(x) = frac{1}{2}frac{{Pvarphi F}}{D}(xZ – x^2).$$
(5)

In cases where we expect radiolytic H2 production rates to significantly vary with depth due to changes in lithology, we adapted the boundary conditions and applied a two-layer diffusion model to account for this variation.
Calculation of Gibbs Energies for the Knallgas reaction
For H2 concentrations above the detection limits at South Pacific IODP Expedition 329 sites (Supplementary Fig. 5)34, we quantified in situ Gibbs energies (ΔGr) of the Knallgas reaction (H2 + ½O2 → H2O). In situ ΔGr values depend on pressure (P), temperature (T), ionic strength, and chemical concentrations, all of which are explicitly accounted for in our calculations:

$$Delta G_{mathrm{r}} = Delta G^circ _{mathrm{r}}left( {T,P} right) + 2.3,RT,{mathrm{log}}_{10}Q$$
(6)

where:
ΔGr: in situ Gibbs energy of reaction (kJ mol H2−1)
ΔG°r(T,P): Gibbs energy of reaction under in situ T and P conditions (kJ mol H2−1)
R: gas constant (8.314 kJ−1 mol K−1)
Q: activity quotient of compounds involved in the reaction.
We use the measured composition of the sedimentary pore fluid to determine values of Q.
For a more complete overview of in situ Gibbs energy-of-reaction calculations in subseafloor sediment, see Wang et al.87.
Calculation of organic oxidation rates (net rates of O2 reduction and DIC production)
We calculated the vertical distribution of net O2 reduction rates at nine sites where the sediment is oxic from seafloor to basement and the vertical distribution of DIC production rates at seven sites where the subseafloor sediment is anoxic (see Supplementary Fig. 3 for site locations). Dissolved O2 concentrations are from Røy et al.47 and D’Hondt et al.88. DIC concentrations are from ODP Leg 201 (ref. 35), and the Proceedings of the IODP Expedition 323 (Sites U12343, U1345)54 and IODP Expedition 329 (Site U1371 (ref. 34)).
The net rates are calculated using the MatLab program and numerical procedures of Wang et al.89, modified by using an Akima spline, rather than a 5-point running mean, to generate a best-fit line to the chemical concentration data. Details of the calculation protocol for O2 production rates and DIC production rates are respectively described in the supplementary information of D’Hondt et al.88 and in Walsh et al.90. The DIC reaction rates and their first standard deviations calculated for the seven sites are given in Supplementary Table 7.
To facilitate comparisons of radiolytic chemical rates to net DIC production rates, rates are converted to electron equivalents (2 electrons per H2, 4 electrons per O2, 4 electrons per organic C oxidized).
Estimation of sediment ages
We estimated sediment ages for Sites U1343 and U1343 using the sediment-age model of Takahashi et al.54, which is based on biostratigraphic and magnetostratigraphic data. Because detailed chronostratigraphic data are not available for the remaining sites (Equatorial Pacific sites (1225 and 1226), Peru Trench Site 1230 and Peru Basin Site 1231, South Pacific sites U1365, U1366, U1367, U1369, U1370, and U1371, North Pacific sites EQP9 and EQP10, and North Atlantic sites KN223-11 and KN223-12), we used the mean sediment accumulation rate for each of these sites (Supplementary Fig. 3) to convert its sediment depth (in meters below seafloor) to sediment age (in millions of years, Ma). Mean sediment accumulation rate was calculated by dividing sediment thickness by basement age91 (Supplementary Table 8). For Sites 1225, 1226, 1230, 1231, U1365, U1366, U1367, U1369, U1370, and U1371, sediment thickness was determined by drilling to basement34,35. For Sites EQP9, EQP10, KN223-11, and KN223-12, sediment thicknesses were determined from acoustic basement reflection data. More

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