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25 August 2021

African tropical montane forests store more carbon than was thought

The inaccessibility of African montane forests has hindered efforts to quantify the carbon stored by these ecosystems. A remarkable survey fills this knowledge gap, and highlights the need to preserve such forests.

Nicolas Barbier

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Nicolas Barbier

Nicolas Barbier is at AMAP, Université de Montpellier, IRD, CNRS, INRAE, CIRAD, Montpellier 34980, France.

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In a paper in Nature, Cuni-Sanchez et al.1 report the assembly of a large database of tree inventories for 226 mature montane-forest plots in 12 African countries. The authors analyse the data to determine the amount of aboveground biomass and carbon stored in these highly diverse and threatened ecosystems. Their results suggest that African montane forests store more carbon than was previously thought, and the findings should help to guide efforts to conserve these ecosystems.Cuni-Sanchez and colleagues measured trunk diameters and heights of the trees in plots, and identified the botanical species to deduce wood density — an approach that constitutes the gold standard for estimating the biomass, and thus the amount of carbon, contained per unit of forest area. This method involves the use of general statistical equations for describing tree form, called allometric models, and considers only the aboveground parts of trees. It therefore disregards several other pools of carbon, notably in the roots and soil. The overall approach might seem crude, but recognizing and measuring the many hundreds of tree species found on steep, cloud-shrouded slopes (Fig. 1), let alone the underground carbon, without visiting the sites, will long remain difficult, even with the best drones and satellite systems.

Figure 1 | Montane forest in Boginda, Ethiopia. Cuni-Sanchez et al.1 use data from a survey of montane tropical forests in Africa to quantify the amount of carbon stored above ground in these ecosystems.Credit: Bruno D’Amicis/Nature Picture Library

Anyone who has conducted field inventories in tropical mountains knows that measuring and identifying 72,336 trees, often just a few steps away from the void, is an amazing feat. For comparison, a previously reported study2 based its estimates of the carbon stored in montane African forests on as few as seven plots. The study also brings together contributions from numerous researchers and institutions, including many in Africa, to greatly increase the size of the data set, which is also a remarkable achievement. Even so, the total area of forest studied is less than 150 hectares, whereas African montane forest covers about 100,000 times that area, inevitably raising questions about how representative the inventory is.Statisticians might raise their eyebrows at the sampling design. As is usually the case in meta-analyses, the data set was neither homogeneous (for example, there is a roughly tenfold variation in the plot sizes), nor were the sites selected at random. However, the authors did their best to rule out possible biases induced by sampling artefacts.
Read the paper: High aboveground carbon stock of African tropical montane forests
Cuni-Sanchez et al. chose not to discuss one tricky aspect of surveys of this sort (extensively discussed elsewhere2): how should the land area of a steep slope be measured? The authors followed standard practice, which is to measure the extent of forest plots and of land-cover types in reference to horizontal, planimetric areas (that is, the areas that would be represented on a 2D map, as if seen from the air). This tends to overestimate aboveground carbon because the sloped surface area is greater than that of the planimetric area — which means that the tree density of the planimetric area is higher than it is on the slope. By contrast, the use of planimetric areas underestimates total montane-forest area (by about 40%; see ref. 2). These two biases should roughly cancel each other out when estimating carbon stocks, or changes to stocks, for a region or country. But care should be taken not to combine data acquired using planimetric and non-planimetric areas in future meta-analyses, because the resulting estimates could end up well off the mark.One might expect that trees in mature African montane forests would be, on average, shorter — and therefore store less carbon — than their lowland counterparts, because of their lower environmental temperatures and shallow soils, frequent landslides and strong winds. However, this is not what Cuni-Sanchez et al. report. Instead, they find that average aboveground carbon stocks are not significantly different from those of mature lowland forests. This contrasts with the situation in the neotropics and southeast Asia, where montane forests store, on average, less carbon than do lowland forests.However, the new results fit with the 2016 discovery that the tallest African trees (81.5 metres) grow on Mount Kilimanjaro3, the highest mountain in Africa. African forests, in general, tend to contain fewer but larger-statured tree stands than does, for example, Amazonia4. The current study confirms that this peculiarity applies even at high altitudes.The authors investigate several possible drivers for the variations in biomass observed at different sites in their study, including topography, climate, landslide hazard, and even the presence of elephants or certain conifers (Podocarpaceae), but were unable to identify any clear pattern. Many environmental, historical and biological effects probably interact, with each of these effects varying greatly in ways that are poorly captured by available data sets. These effects must therefore be disentangled before a predictive model of African montane carbon distribution can be developed.
Tropical carbon sinks are saturating at different times on different continents
Nevertheless, Cuni-Sanchez and colleagues’ study underlines a crucial message: African montane forests are immensely valuable, and not only because they host the source of the River Nile, mountain gorillas and ecosystems such as mysterious lichen-covered forests. They also store vast amounts of carbon, and thereby have a key role in tackling climate change. Of course, this immense intrinsic value does not preclude intense human exploitation of these ecosystems, which can lead to rapid degradation and deforestation. For instance, on the basis of satellite monitoring, Cuni-Sanchez and colleagues report that Mozambique lost nearly one-third of its montane forests between 2000 and 2018.There is, however, the faint hope that putting a financial value on carbon, and the establishment of economic incentives to avoid deforestation in tropical countries, might help to check the flood of damage5. The aim is to reward African countries — for which montane forest sometimes constitutes the last remaining forests — for their conservation endeavours, and for renouncing efforts to access the timber and ore in these ecosystems, even when such resources are otherwise desperately lacking. By gathering the best-available data to provide precise, country-level estimates of average aboveground carbon content in African montane forests, Cuni-Sanchez and colleagues’ study will add weight to such efforts — not least because the new estimates are, on average, two-thirds higher than the values reported by the Intergovernmental Panel on Climate Change6.The next step should be to extend measurements in these forests, particularly by continuing to support national forest-inventory efforts. These inventories target all vegetation types, rather than just the most intact forests, and all carbon pools, using standardized protocols and systematic sampling methods. Remote sensors, both in the sky and in space, should also be used to fully map the detailed spatial variation of forest diversity, structure and dynamics. But there is no excuse for delaying policymaking — we already know enough to justify immediate decisive action to preserve yet another of Earth’s threatened treasures.

Nature 596, 488-490 (2021)
doi: https://doi.org/10.1038/d41586-021-02266-3

References1.Cuni-Sanchez, A. et al. Nature 596, 536–542 (2021).Article 

Google Scholar 
2.Spracklen, D. V. & Righelato, R. Biogeosciences 11, 2741–2754 (2014).Article 

Google Scholar 
3.Hemp, A. et al. Biodivers. Conserv. 26, 103–113 (2017).Article 

Google Scholar 
4.Lewis, S. L. et al. Phil. Trans. R. Soc. B 368, 20120295 (2013).PubMed 
Article 

Google Scholar 
5.Venter, O. et al. Science 326, 1368 (2009).PubMed 
Article 

Google Scholar 
6.Domke, G. et al. in 2019 Refinement to the 2006 IPCC Guidelines for National Greenhouse Gas Inventories Vol. 4 (eds Calvo Buendia, E. et al.) Ch. 4, 4.48 (IPCC, 2019).
Google Scholar 
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In place of its normal load of cars and vans, the repurposed ferry boat sported a mobile science laboratory and a large fan on its deck as it left Townsville, Australia, in March. Researchers dropped anchor in a coral lagoon some 100 kilometres offshore and then fired up the cone-shaped turbine, which blew a mist of seawater off the back of the boat. What happened next came as a welcome surprise: after briefly drifting along the ocean surface, the plume ascended into the sky.Looking a bit like a jet engine, this mist machine is at the centre of an experiment that, if successful, could help to determine the future of the Great Barrier Reef. Three-hundred and twenty nozzles spewed a cloud of nano-sized droplets engineered to brighten clouds and block sunlight — providing a bit of cooling shade for the coral colonies below. Scientists used sensors aboard the ferry, drones and a second boat to monitor the plume as it migrated skyward.The experiment wasn’t big enough to significantly alter the clouds. But preliminary results from the field tests — which were shared exclusively with Nature — suggest that the technology might perform even better than computer models suggested it would, says Daniel Harrison, an oceanographer and engineer at Southern Cross University in Coffs Harbour, Australia, who is heading up the research. “We are now very confident that we can get the particles up into the clouds,” Harrison says. “But we still need to figure out how the clouds will respond.”Harrison’s project is the world’s first field trial of marine cloud brightening, one of several controversial geoengineering technologies that scientists have studied in the laboratory for decades. The research has been driven by fear that humans might one day be forced to deliberately manipulate the Earth’s climate and weather systems to blunt the most severe impacts of global warming.For many Australians, that day arrived in 2017, when a marine heat wave spurred massive coral bleaching and death across much of the 2,300-kilometre Great Barrier Reef. That crisis hit just a year after another bleaching event along the reef, which supports more than 600 species of coral and an estimated 64,000 jobs in industries such as tourism and fishing. Research suggests that the reef lost more than half of its coral between 1995 and 2017, as a result of warming waters, tropical storms and predatory starfish (A. Dietzel et al. Proc. R. Soc. B. 287, 20201432; 2020).
These corals could survive climate change — and help save the world’s reefs
The project has raised concerns among some scientists abroad, in part because the Australian group has published little about its work. Environmentalists outside Australia objected to the project last year after news of the first trial broke, and there could be similar criticism when details of the 2021 trial emerge.Harrison stresses that the cloud-brightening project is about local adaptation to climate change, not global geoengineering, because its application would be limited in both space and time. It’s also just one part of a larger Aus$300 million (US$220 million) Reef Restoration and Adaptation Program (RRAP) launched last year by Australia to investigate and develop techniques and technologies to save the country’s reefs. Many of the proposals, from cloud brightening to breeding heat-tolerant corals, would represent unprecedented human interventions in the natural reef system.Ecological modelling suggests that a large-scale intervention involving multiple strategies — including a fleet of mist machines — could prolong the life of the reef while governments work to eliminate greenhouse-gas emissions. The goal now is to work out what’s achievable in the real world, says Cedric Robillot, executive director of the RRAP.“You need to consider every angle, from the fundamental science to the very pointy end of engineering, if you want to succeed,” Robillot says. “It’s not enough to just prove you could do it. You need to explain how you would do it.”Into the cloudsHarrison conducted his first field test in March 2020: a three-day proof-of-concept expedition on a small car ferry with four scientists, one representative from a local Indigenous group, and two shipping containers for equipment and sleeping quarters. The team had a minimal Aus$400,000 budget and limited scientific instrumentation to monitor the mist, but it was enough to document that the plume flowing out of their mist machine rode a draught of warm air high into the sky.It was the first time they had witnessed this phenomenon. Their models had suggested that evaporation of the brine droplets would cool the plume, which would then float across the surface of the ocean, only slowly mixing upwards into the low-lying marine clouds. The models also indicated a risk that the tiny droplets might merge and drop out of the air. Instead, brine droplets floated along the surface of the ocean for half a kilometre without coalescing, gradually losing water and weight to evaporation along the way. And then they shot upwards.

A marine heat wave in 2017 caused coral bleaching along much of Australia’s Great Barrier Reef.Credit: Juergen Freund/Nature Picture Library

“We didn’t expect that at all,” Harrison says, “but it turned out we were doing this experiment in the middle of a rising air mass.”The scientists feared it was a fluke. Although years of research and development have gone into the nozzles, initially led by a separate American team, this was the first time anybody had ever deployed them in the field with fresh seawater. The team also didn’t know what to expect from clouds and aerosols in that region, because research on the reef has focused almost exclusively on what happens below the water, not the conditions above.For Harrison, the 2020 experiment was more than enough to justify moving forward with another, larger trial in March 2021. But it did raise eyebrows among some scientists and observers abroad, where geoengineering research has met strong opposition and struggled to attract funding.
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Most of the concern has centred on a form of solar geoengineering that involves injecting reflective material into the stratosphere to block sunlight at a global scale. But cloud brightening has also been studied as a potential global intervention, and it has attracted criticism from some environmental groups who argue that it carries inevitable ecological risks and detracts from efforts to limit greenhouse gases.Some scientists, as well as environmental advocates who follow geoengineering research, told Nature that they were surprised to see the experiment move forward without more scrutiny — or without published research to justify such an investment.Critics also worry that Australia is setting the wrong kind of precedent by rebranding a solar-geoengineering experiment that could have regional impacts as a local adaptation project. “One could say that there should have been some level of consultation with the outside world,” says Janos Pasztor, who heads the Carnegie Climate Governance Initiative, an advocacy group in New York City that has been pushing for a global debate over geoengineering governance in the United Nations.Harrison says scientists in the programme have consulted with regulatory authorities, as well as with the general public and Indigenous groups that have historic claims on the reef. He also readily acknowledges trying to avoid getting embroiled in a debate about solar geoengineering, arguing that the project would be more akin to cloud-seeding operations that are designed to promote rain and that are not considered to be geoengineering. One of the next modelling efforts, however, will be to explore any potential regional and global implications, he says.

A plume of seawater droplets rises up into the sky during a field trial in March 2021.Credit: Brendan Kelaher/SCU

Others question the Australian government’s motivations in funding such work. Under the conservative prime minister Scott Morrison, the government has yet to strengthen its climate pledge under the 2015 Paris agreement, as many nations have done in the past year. Morrison has personally ruled out committing to net-zero emissions. Pushing for a technological fix to global warming without moving to aggressively curb greenhouse gases is “sheer lunacy”, says Peter Frumhoff, chief climate scientist for the Union of Concerned Scientists, an advocacy group in Cambridge, Massachusetts.Some researchers, however, are pleased to see marine cloud brightening move from theory to the field, including US scientists working on a similar project that has been struggling to get into the field for nearly a decade. “This is an early example of how climate disruption can drive interest in these things,” says Sarah Doherty, an atmospheric physicist who manages the Marine Cloud Brightening Project at the University of Washington in Seattle. Members of the team provided the initial nozzle design and have been tracking the Australian group’s progress.Coral crisisThe first time that scientists observed a major bleaching event along the Great Barrier Reef was in 1998, and the second event followed four years later. In both cases, corals expelled the algae that live within them and that provide colour and energy through photosynthesis. Most of the corals eventually recovered. But in 2016 and 2017, many corals bleached and then died across two-thirds of the reef.
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“It was absolutely horrifying,” says David Wachenfeld, chief scientist at the Great Barrier Reef Marine Park Authority, which manages the reef. The clear message from those events was that the traditional approach to managing corals and coral reefs would not be enough, he adds. “Our hand was forced.”In 2018, the Australian government allocated Aus$6 million to a consortium of universities and government research institutes for a feasibility study focused on potentially radical strategies that could be applied across the reef. Researchers reviewed some 160 ideas, including putting live corals on ice for long-term preservation and synthetically engineeering new varieties that can tolerate the warmer waters. Many approaches proved too costly and energy intensive, but 43 interventions were singled out for further study. Marine cloud brightening drew support in part because it theoretically provides direct relief precisely when and where corals need it most.Much of the emphasis of the programme is on helping corals to adapt and repopulate the reef, including efforts to improve coral aquaculture operations so that they can produce millions of corals per year rather than thousands. For Madeleine van Oppen, a coral geneticist at the Australian Institute of Marine Science near Townsville, the RRAP programme helps to integrate her team’s work on assisting coral evolution to make them more heat tolerant.Thanks to the RRAP, she says, data from those projects are now being fed directly into models that enable researchers to assess the potential benefits — as well as the risks — of releasing new strains of coral and microalga into the wild. The programme is also raising ecological questions, such as whether the introduction of new coral species can propagate disease, or whether a new variety of more heat-tolerant corals might displace corals struggling to survive.

Researchers are testing specialized nozzles that create jets of seawater mist.Credit: Alejandro Tagliafico/SCU

“It sort of speeds up the whole path from research to implementation in the field,” says van Oppen.In the long run, the models indicate that without interventions, the extent of coral on the reef could shrink by well over 60% by 2070 compared with 2020 levels (S. A. Condie et al. R. Soc. Open Sci. 8, 201296; 2021). But simulations suggest that Australia could cut those losses in half with a three-pronged approach focused on propagating heat-tolerant corals, controlling outbreaks of the predatory crown-of-thorns starfish and brightening clouds to take the edge off of heat waves. Crucially, the latest modelling also suggests that without the cooling provided by Harrison’s cloud brightening project, the other interventions might not amount to much.Testing the windWhen Harrison’s group returned to the field this year, they had more-powerful drones as well as other aerosol sensors on a second boat. As in the previous year’s experiment, each time they created a plume, it rose into the sky after the droplets lost around 90% of their water to evaporation. The likely explanation, Harrison says, is that the reef is creating its own weather as warm water along the shallow corals heats the air above.Many more droplets are making it into the clouds than the scientists had initially calculated, but Harrison says their mist machine might need to be scaled up by a factor of 10 — from 320 to around 3,000 nozzles — to produce enough particles to brighten nearby clouds by around 30%. His team’s modelling suggests that this could in turn reduce the incoming solar radiation on the reef locally by around 6.5%. Even then, the operation would require 800–1,000 stations to cover the length of the Great Barrier Reef.
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But it’s unclear whether that spray of salty droplets will have the desired effect, says Lynn Russell, an atmospheric chemist at the Scripps Institution of Oceanography in La Jolla, California, who has studied cloud brightening. Russell has not seen the latest — and as-yet unpublished — results, but questions whether there are enough of the low layered clouds considered suitable for cloud brightening.Harrison acknowledges such concerns and says that his team sees more of these clouds on the southern part of the reef. His team’s modelling suggests the technology will also work on the clouds that are common across the rest of the reef in summer. Even then, he says, it remains unclear how much coverage a full-scale cloud-brightening operation could provide across the entirety of the reef. More measurements, and detailed modelling, are needed to provide answers.For now, Harrison has secured funding for another two years, and he needs to demonstrate progress. The RRAP is testing all 43 approaches and will redistribute resources to projects that show potential, Robillot says. But he stresses that no amount of science and engineering will preserve the reef in its current form. “Even if we do all of this, the system that you’ll end up with is not going to be the Great Barrier Reef that we know today,” Robillot says. “You might, however, retain a very functional ecosystem.”That’s enough to keep Harrison going, and his team is already preparing for a trip into the field in 2022. The scientists plan to run the mist machine at higher pressure, which should produce a sixfold increase in the number of particles, and they will use new instrumentation to determine how particles alter clouds. They are also investigating an entirely different nozzle technology that could reduce the number of nozzles needed by a factor of 1,000.Harrison is more confident today than he was even a year ago that cloud brightening might work over the reef, but he is also realistic about the future if governments fail to limit carbon emissions. “There are only so many clouds available, and there is only so much you can brighten them,” he says. “Eventually, climate change just overwhelms things.”

Nature 596, 476-478 (2021)
doi: https://doi.org/10.1038/d41586-021-02290-3

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Conceptualization of organismal to tumor biogeography through clone phylogeniesWe first map organismal biogeographic concepts and models to the process of migration and colonization of cancer cells during metastasis. Tumors are populations consisting of a diversity of cancer cells with different genetic profiles that may represent different lineages in the clone phylogeny. We use the example in Fig. 1, which contains a phylogeny of 17 clones found in one primary tumor (P) and four metastases (M1–M4). Events occurring along a branch in a phylogeny are anagenetic events, which include diversification, extinction, and expansion12,14. In organismal evolutionary biology, anagenetic events are not directly observed except through the fossil record. However, one can map the collection of genetic variants that likely arose on individual lineage in a phylogeny. In many cancers, sequencing of temporally sampled biopsies’ can directly reveal anagenetic events similar to the sequencing of ancient DNA in paleogenomics.The other types of evolutionary events in the phylogeny are cladogenetic, including genetic divergences and dispersals (Fig. 1). Genetic differences observed among species and populations are the key to detect cladogenetic events reconstructed in molecular phylogenies of living descendants. In cancer, temporal sampling of biopsies can reveal cladogenetic events that produced extinct descendants.In biogeography, genetic divergence results in the diversification of lineages within an area. Sometimes, the term duplication is used, but we avoid its further use because of the confusion it may cause in evolutionary genomics. Divergence events are also observed in a clone phylogeny, particularly when clone lineages diverge from each other within a tumor or across tumors. The exact opposite of genetic diversification can also be observed when lineages partially or fully disappear from the phylogeny. Extinction can occur due to random chance, selection, or environmental pressures. Even though extinction is rarely discussed in tumor clone phylogenetics, it happens frequently.Phylogenies also reveal movements of lineages between locations (geographic areas or body parts) when the locations of individual cells, species, or populations are known5,6,7,8,9,15. When lineages accumulate genetic differences along a branch in the phylogeny, and the evolved lineages migrate to a new area, we observe an expansion event. Expansions differ from dispersals in such that the growth of a population occurs in the same place. This movement of cells of a clone from one location to another, where they would potentially form a metastasis, results in the dispersal of  these cells of that clone to additional areas, which is modeled by a dispersal rate (d) in organismal biogeography. When a clone genetically diverges following its migration, then a distant dispersal event is said to have occurred. Similarly, when a clone diverges from the rest of the clones within a tumor and disperses to another tumor, we have observed an expansion event. Thus, clone phylogenies can give insights into the origin and trajectory of cancer cells between tumors.When a clone is no longer present at a location, it is extinct at that location. Extinctions are modeled by an extinction rate (e) in biogeographic models. As a result of extinction, the range of descendent clones on a phylogeny can be smaller than the ancestors. Biogeography models also have a parameter (J) to consider founder events that establish new populations from a few individuals. In phylogenies, founder events can be detected if only one or a few cells are found to have moved from one location to another to start diversifying in a new area. Both distant dispersal and founder events may result in forming a new colony of cells, i.e., a new metastasis in the case of cancer cell migrations. The primary distinction between dispersal and founder events is the relative number of migrating cells. Founder events are due to one or a few cells, whereas dispersal events involve a larger number of migrating cells. Founder events are expected to be more common in tumor evolution because metastases are thought to be formed by the spread of only one or a few cancer cells. These biogeographic events have been mathematically modeled and implemented in various approaches to infer species migration events12, which are directly applicable in the inference of cancer cell migrations between tumors.Model fitsWe began by analyzing the statistical fits of six biogeographic models (Table 1) to 80 computer-simulated tumor evolutionary datasets. Simulations enable us to assess the performance of computational approaches and reveal potential caveats associated with their use because the ground truth is known. These datasets were simulated using four main clone migration schemes defined by the different number of migrating clones (1–3), the small and large number of tumor areas (5–7 tumors, m5 datasets; 8–11 tumors, m8 datasets), and the different types of source areas of migration (primary or metastasis). The following seeding scenarios reflect this complexity of the clone migration schemes: monoclonal single-source seeding (mS), polyclonal single-source seeding (pS), polyclonal multisource seeding (pM), and polyclonal reseeding (pR) (see “Methods” section).Table 1 Phylogenetic and biogeographic events considered in seven biogeographic models used for analysis.Full size tableWe considered biogeographic models that weigh genetic divergence, dispersal/expansion, and extinction events differently (Table 1). We also explored the provision of including founder events in our models on the accuracy of detecting clone migrations. The parameterization of the aforementioned events results in models with two free parameters, i.e., dispersal rate (d) and extinction rate (e), and models with three free parameters by adding the founder-event speciation (J); see “Methods” section for more details.Overall, we tested six biogeographic models for their fit to the tumor data, three models with two free parameters and three others with three free parameters. BAYAREALIKE, DEC, and DIVALIKE models have two parameters each. They are nested within their respective models that add the founder effect, resulting in a model with three free parameters (hereinafter +J models). We used the BioGeoBEARS software for all model fit analyses. In data analysis, we first inferred phylogeny of cancer cell populations (clone phylogeny) using an existing method16, followed by the use of BioGeoBEARS to infer the clone migration history in which the clone phylogeny is used along with the location of tumor sites in which each clone is observed (Fig. 2). BioGeoBEARS estimates the probabilities of annotating internal nodes with tumor locations. These annotations are then used to derive cancer cell migration paths when two adjacent nodes are annotated with different tumor locations. In these analyses, we assumed the correct clone phylogeny because our focus was not assessing the impact of errors in a phylogeny on the accuracy of clone migration inferences. We also compared the accuracy of migration histories reconstructed using biogeographic models in BioGeoBEARS with those obtained from the approaches that do not model biogeographic processes (BBM9, MACHINA5, and PathFinder7).Figure 2Data analysis pipeline using BioGeoBEARS in R14 to infer clonal migration histories.Full size imageWe first conducted Likelihood Ratio Tests (LRTs) to examine the improvement offered by considering founder events in modeling tumor migrations. In this case, the fit of the BAYAREALIKE, DEC, and DIVALIKE models was compared to their +J counterparts, respectively. The null hypothesis was rejected for more than 50% of the datasets (BAYAREALIKE: 71.25%, DEC: 60%, and DIVALIKE: 53.75%; P  More

The Fur-regulated T6SS1 plays an important role in iron acquisition in C. necator
To explore the function of T6SS1 (Reut_A1713 to Reut_A1733) in C. necator (Fig. S1A), we analyzed the T6SS1 promoter and identified a Fur binding site (AGAAATA) upstream of gene reut_A1733. This Fur binding site was highly similar to the Fur-box reported in E. coli [38], with a probability score of 2.25 (out of a maximum score = 2.45) (Fig. S1B), which was calculated by applying the position weight matrix to a sequence [39]. Incubation of the T6SS1 promoter probe with purified Fur protein led to decreased mobility of the probe in the electrophoretic mobility shift assay, suggesting a direct interaction between Fur and the T6SS1 promoter (Fig. 1A). To further determine the function of Fur on the expression of T6SS1, a single-copy PT6SS1::lacZ fusion reporter was introduced into the chromosomes of C. necator wild-type (WT), Δfur deletion mutant, and the Δfur(fur) complementary strain. Compared to WT, the PT6SS1::lacZ promoter activity was significantly increased in the Δfur mutant (about 2.2-fold), and this increase could be restored by introducing the complementary plasmid pBBR1MCS-5-fur (Fig. 1B). Similar results were obtained by analyzing the expression of T6SS1 core component genes (hcp1, clpV1, vgrG1, and tssM1) with qRT-PCR (Fig. S1C). These results demonstrate that the expression of T6SS1 in C. necator is directly repressed by Fur, the master regulator of genes involved in iron homeostasis in many prokaryotes [40, 41].Fig. 1: Regulation of T6SS1 expression by Fur.A The interactions between His6-Fur and the T6SS1 promoter examined by EMSA. Increasing amounts of Fur (0, 0.03, 0.06, 0.13, 0.25, and 1.0 μM) and 10 nM DNA fragments were used in the assay. A 500 bp unrelated DNA fragment (Control A) and 1 µM BSA (Control B) were included in the assay as negative controls. B Fur represses the expression of T6SS1. β-galactosidase activities of T6SS1 promoter from chromosomal lacZ fusions in relevant C. necator strains were measured. C Iron uptake requires T6SS1. Stationary-phase C. necator strains were washed twice with M9 medium. Iron associated with indicated bacterial cells were measured with ICP-MS. The vector corresponds to the plasmid pBBR1MCS-5 (B) and pBBR1MCS-2 (C), respectively. Data are represented as mean values ± SD of three biological replicates, each with three technical replicates. **p  More

AffiliationsAnimal Production Systems group, Wageningen University & Research, Wageningen, The NetherlandsAbigail Muscat, Evelien M. de Olde, Raimon Ripoll-Bosch & Imke J. M. de BoerFarming Systems Ecology group, Wageningen University & Research, Wageningen, The NetherlandsHannah H. E. Van ZantenPublic Administration and Policy group, Wageningen University & Research, Wageningen, The NetherlandsTamara A. P. Metze & Catrien J. A. M. TermeerPlant Production Systems group, Wageningen University & Research, Wageningen, The NetherlandsMartin K. van IttersumAuthorsAbigail MuscatEvelien M. de OldeRaimon Ripoll-BoschHannah H. E. Van ZantenTamara A. P. MetzeCatrien J. A. M. TermeerMartin K. van IttersumImke J. M. de BoerCorresponding authorCorrespondence to
Imke J. M. de Boer. More

Department of Environment and Geography, University of York, York, UKAida Cuni-Sanchez, Philip J. Platts, Rob Marchant & Andrew MarshallDepartment of International Environmental and Development Studies (NORAGRIC), Norwegian University of Life Sciences, Ås, NorwayAida Cuni-SanchezDepartment of Natural Sciences, Manchester Metropolitan University, Manchester, UKMartin J. P. SullivanSchool of Geography, University of Leeds, Leeds, UKMartin J. P. Sullivan, Simon L. Lewis, Serge K. Begne, Amy C. Bennett, Martin Gilpin, Jon Lovett & Oliver L. PhillipsLeverhulme Centre for Anthropocene Biodiversity, University of York, York, UKPhilip J. PlattsClimate Change Specialist Group, Species Survival Commission, International Union for Conservation of Nature, Gland, SwitzerlandPhilip J. PlattsDepartment of Geography, University College London, London, UKSimon L. LewisBiology Department, Université Officielle de Bukavu, Bukavu, Democratic Republic of the CongoGérard Imani & Christian AmaniService of Wood Biology, Royal Museum for Central Africa, Tervuren, BelgiumWannes Hubau, Hans Beeckman & John T. MukendiDepartment of Environment, Laboratory of Wood Technology (Woodlab), Ghent University, Ghent, BelgiumWannes HubauUniversity of Jos, Jos, NigeriaIveren AbiemNigerian Montane Forest Project, Yelwa Village, NigeriaIveren Abiem & Hazel ChapmanDepartment of Geosciences and Geography, University of Helsinki, Helsinki, FinlandHari Adhikari, Janne Heiskanen & Petri PellikkaDepartment of Zoology, Faculty of Science, Charles University, Prague, Czech RepublicTomas AlbrechtInstitute of Vertebrate Biology, Czech Academy of Sciences, Brno, Czech RepublicTomas AlbrechtInstitute of Botany of the Czech Academy of Science, Třeboň, Czech RepublicJan Altman & Jiri DolezalCollege of Natural and Computational Science, Addis Ababa University, Addis Ababa, EthiopiaAbreham B. Aneseyee & Teshome SoromessaDepartment of Natural Resource Management, College of Agriculture and Natural Resource, Wolkite University, Wolkite, EthiopiaAbreham B. AneseyeeEuropean Commission, Joint Research Centre, Ispra, ItalyValerio AvitabileUK Centre for Ecology and Hydrology, Edinburgh, UKLindsay BaninUniversité du Cinquantenaire Lwiro, Département de sciences de l’environnement, Kabare, Democratic Republic of the CongoRodrigue BatumikeIsotope Bioscience Laboratory (ISOFYS), Ghent University, Ghent, BelgiumMarijn Bauters, Pascal Boeckx & Joseph OkelloPlant Systematic and Ecology Laboratory, Higher Teachers’ Training College, University of Yaoundé I, Yaoundé, CameroonSerge K. Begne, Vincent Droissart, Marie-Noel Kamdem, Murielle Simo-Droissart & Bonaventure SonkéInstitute of Tropical Forest Conservation, Mbarara University of Science and Technology, Mbarara, UgandaRobert BitarihoBiodiversity and Landscape Unit, Gembloux Agro-Bio Tech, Université de Liege, Liège, BelgiumJan BogaertInstitute for Geography, Friedrich Alexander University, Erlangen–Nuremberg, GermanyAchim Bräuning & Ulrike HiltnerDépartement de Eaux et Forêts, Institut Supérieur d’Agroforesterie et de Gestion de l’Environnement de Kahuzi-Biega (ISAGE-KB), Kalehe, Democratic Republic of the CongoFranklin BulonvuUN Environment World Conservation Monitoring Center (UNEP-WCMC), Cambridge, UKNeil D. BurgessComputational and Applied Vegetation Ecology (CAVElab), Faculty of Bioscience Engineering, Ghent University, Ghent, BelgiumKim Calders & Hans VerbeeckDepartment of Anthropology, George Washington University, Washington DC, USAColin ChapmanSchool of Life Sciences, University of KwaZulu-Natal, Pietermaritzburg, South AfricaColin ChapmanShaanxi Key Laboratory for Animal Conservation, Northwest University, Xi’an, ChinaColin ChapmanInternational Centre of Biodiversity and Primate Conservation, Dali University, Dali, ChinaColin ChapmanUniversity of Canterbury, Canterbury, New ZealandHazel ChapmanInventory and Monitoring Program, National Park Service, Fredericksburg, VA, USAJames ComiskeyUniversity of Ghent, Ghent, BelgiumThales de HaullevilleWorld Agroforestry (ICRAF), Nairobi, KenyaMathieu DecuyperLaboratory of Geo-Information Science and Remote Sensing, Wageningen University, Wageningen, The NetherlandsMathieu Decuyper & Martin HeroldGeography, Environment and Geomatics, University of Guelph, Guelph, Ontario, CanadaBen DeVriesFaculty of Science, University of South Bohemia, České Budějovice, Czech RepublicJiri DolezalAMAP Lab, Université de Montpellier, IRD, CNRS, INRAE, CIRAD, Montpellier, FranceVincent DroissartFaculté de Gestion de Ressources Naturelles Renouvelables, Université de Kisangani, Kisangani, Democratic Republic of the CongoCorneille Ewango & Janvier LisingoCollege of Development Studies, Addis Ababa University, Addis Ababa, EthiopiaSenbeta FeyeraDendrochronology Laboratory, World Agroforestry Centre (ICRAF), Nairobi, KenyaAster GebrekirstosMissouri Botanical Garden, St Louis, MO, USARoy GereauDepartment of Biology, University of Burundi, Bujumbura, BurundiDismas HakizimanaSmithsonian Institution Forest Global Earth Observatory (ForestGEO), Smithsonian Tropical Research Institute, Washington DC, USAJefferson Hall & David KenfackKunming Institute of Botany, Kunming, ChinaAlan HamiltonUniversité Libre de Bruxelles, Brussels, BelgiumOlivier HardyDivision of Vertebrate Zoology, Yale Peabody Museum of Natural History, New Haven, CT, USATerese HartInstitute for Atmospheric and Earth System Research, Faculty of Science, University of Helsinki, Helsinki, FinlandJanne HeiskanenDepartment of Plant Systematics, University of Bayreuth, Bayreuth, GermanyAndreas HempHelmholtz Center Potsdam GFZ German Research Centre for Geosciences, Section 1.4 Remote Sensing and Geoinformatics, Potsdam, GermanyMartin HeroldHelmholtz-Centre for Environmental Research (UFZ), Leipzig, GermanyUlrike HiltnerDepartment of Ecology, Faculty of Science, Charles University, Prague, Czech RepublicDavid Horak & Ondrej SedlacekInternational Gorilla Conservation Programme, Musanze, RwandaCharles Kayijamahe & Eustrate UzabahoDepartment of Natural Resources, Karatina University, Karatina, KenyaMwangi J. KinyanjuiDepartment of Ecosystem Science and Sustainability, Colorado State University, Fort Collins, CO, USAJulia KleinEco2librium LLC, Boise, ID, USAMark LungDepartment of Ecology, Université de Kisangani, Kisangani, Democratic Republic of the CongoJean-Remy MakanaEnvironmental Change Institute, School of Geography and the Environment, University of Oxford, Oxford, UKYadvinder MalhiTropical Forests and People Research Centre, University of the Sunshine Coast, Sippy Downs, Queensland, AustraliaAndrew Marshall & Alain S. K. NguteFlamingo Land Ltd, Malton, UKAndrew MarshallCollege of African Wildlife Management, Mweka, TanzaniaEmanuel H. MartinSchool of GeoSciences, University of Edinburgh, Edinburgh, UKEdward T. A. Mitchard & Charlotte WheelerDepartment of Geography and Environmental Sciences, University of Dundee, Dundee, UKAlexandra MorelIndependent Botanist, Harare, ZimbabweTom MullerDepartment of Horticultural Sciences, Faculty of Applied Sciences, Cape Peninsula University of Technology, Bellville, South AfricaFelix NchuBiology Department, University of Rwanda, Kigali, RwandaBrigitte Nyirambangutse & Etienne ZiberaDepartment of Biological and Environmental Sciences, University of Gothenburg, Gothenburg, SwedenBrigitte Nyirambangutse & Göran WallinMountains of the Moon University, Fort Portal, UgandaJoseph OkelloNational Agricultural Research Organisation, Mbarara Zonal Agricultural Research and Development Institute, Mbarara, UgandaJoseph OkelloSchool of Biological Sciences, University of Southampton, Southampton, UKKelvin S.-H. PehConservation Science Group, Department of Zoology, University of Cambridge, Cambridge, UKKelvin S.-H. PehState Key Laboratory of Information Engineering in Surveying, Mapping and Remote Sensing, Wuhan University, Wuhan, ChinaPetri PellikkaKey Biodiversity Areas Secretariat, BirdLife International, Cambridge, UKAndrew PlumptreSchool of Life Sciences, University of Lincoln, Lincoln, UKLan QieDepartment of Biology, University of Florence, Sesto Fiorentino, ItalyFrancesco RoveroTropical Biodiversity Section, Museo delle Scienze, Trento, ItalyFrancesco RoveroTropical Plant Exploration Group (TroPEG), Mundemba, CameroonMoses N. SaingeCenter for Development Research (ZEF), University of Bonn, Bonn, GermanyChristine B. SchmittConservation and Landscape Ecology, University of Freiburg, Freiburg, GermanyChristine B. SchmittApplied Biology and Ecology Research Unit, University of Dschang, Dschang, CameroonAlain S. K. NguteForest Ecology and Forest Management Group, Wageningen University, Wageningen, The NetherlandsDouglas SheilWater and Land Resources Center, Addis Ababa University, Addis Ababa, EthiopiaDemisse ShelemeAfrican Wildlife Foundation (AWF), Biodiversity Conservation and Landscape Management Program, Simien Mountains National Park, Debark, EthiopiaTibebu Y. SimegnFaculty of Forestry, University of British Columbia, Vancouver, British Columbia, CanadaTerry SunderlandCenter for International Forestry Research (CIFOR), Bogor, IndonesiaTerry SunderlandDepartment of Forest Ecology, Faculty of Forestry and Wood Sciences, Czech University of Life Sciences, Prague, Czech RepublicMiroslav SvobodaDepartment of Plant Biology, Faculty of Sciences, University of Yaoundé I, Yaoundé, CameroonHermann TaedoumgBioversity International, Yaoundé, CameroonHermann TaedoumgUK Research and Innovation, London, UKJames TaplinDepartment of Geography, National University of Singapore, Singapore, SingaporeDavid TaylorInstitute of Forestry and Conservation, University of Toronto, Toronto, Ontario, CanadaSean C. ThomasBiodiversity Foundation for Africa, East Dean, UKJonathan TimberlakeForestry Development Authority of the Government of Liberia (FDA), Monrovia, LiberiaDarlington TuagbenSchool of Forestry and Environmental Studies, Yale University, New Haven, CT, USAPeter UmunayDepartment of Biological Sciences, Florida International University, Miami, FL, USAJason VleminckxSchool of Natural Sciences, University of Bangor, Bangor, UKSimon WillcockRothamsted Research, Harpenden, UKSimon WillcockUniversity of Liberia, Monrovia, LiberiaJohn T. WoodsA.C.-S. conceived the study and assembled the AfriMont dataset. A.C.-S. and M.J.P.S. analysed the plot data (with contributions from S.L.L.) and wrote the manuscript. P.J.P. analysed forest extents and contributed to writing. S.L.L. conceived and managed the AfriTRON forest plot recensus programme. E.T.A.M. and V.A. helped compare plot data with remote sensing carbon maps. All co-authors read and approved the manuscript. More

The results are organized into subsections on in situ temperature profiles, geochemical gradients, and microbial community data. Geochemical data include concentration and isotopic data of dissolved electron acceptors (sulfate, dissolved inorganic carbon (DIC), δ13C-DIC), electron donors (methane, sulfide, SCOAs), and respiration end products (DIC, methane, sulfide), as well as solid-phase organic carbon pools (total organic carbon (TOC), δ13C-TOC, total nitrogen (TN), TOC:TN (C:N)). Microbial community data include bacterial and archaeal 16S rRNA gene copy numbers and bacterial and archaeal community trends. All geochemical and microbiological data are shown in Supplementary Data 1–4.Temperature profilesThe in situ temperatures and temperature gradients differ greatly among sites and hydrothermal areas (Table 1; Fig. 1a, b, 1st column). Certain locations within the SA (Cold Site) and NSA (MUC02, GC13, MUC12) are uniformly cold (~3–5 °C) and thus serve as low-temperature control sites. The fact that Cold Site has no measurable depth-dependent temperature increase suggests that this site, despite being located within the SA, only has minimal hydrothermal fluid seepage. At two sites from the NSA (GC09, GC10), temperatures increase strongly, reaching ~60 °C at 400 cm below the seafloor, with temperature gradients becoming linear below 50 cm. Everest Mound, Orange Mat, and Cathedral Hill in the SA have the steepest temperature gradients ( >165 °C m−1), reaching >80 °C within 25 cm, whereas Yellow Mat from the SA only reaches ~27 °C at 45 cm. Temperature gradients are near-linear at Everest Mound, Cathedral Hill, and Yellow Mat, and in the top ~15 cm of Orange Mat. Below ~15 cm, the temperatures at Orange Mat are nearly constant.Table 1 Overview of all sampling sites.Full size tableFig. 1: Microbial abundance and community structure in relation to temperature and geochemical gradients.Depth profiles of temperature (1st column), porewater dissolved sulfate, methane, and dissolved inorganic carbon (DIC) concentrations (2nd column), bacterial and archaeal 16S rRNA gene abundances (3rd column), bacterial (4th column) and archaeal community structure (5th column) across the 10 study sites. a Sites from the NSA. b Sites from the SA. Bacteria and Archaea community structure is shown at the phylum level, except in Proteobacteria, which are displayed at the class level (see asterisk). To improve visibility, we adjusted the depth axis range for bacterial and archaeal communities at Everest Mound, only showing the top 10 cm, where microbial 16S rRNA genes were above detection. Sulfate and methane data from the NSA, except those from MUC12, were previously published27.Full size imageConcentrations of methane, sulfate, sulfide, and DICPorewater concentration profiles of methane, sulfate and DIC are consistent with higher microbial activity and higher substrate supplies in hydrothermal seep sediments compared to cold control sites or hydrothermal non-seep sediments.Independent of temperature, sediments without fluid seepage, i.e. the hydrothermal NSA sites (GC09, GC10) and low-temperature control sites (MUC02, MUC12, GC13, Cold Site), have similar concentration profiles of sulfate, methane, and DIC (Fig. 1a, b, 2nd column). Methane remains at background concentrations (≤0.02 mM), suggesting minimal methane production. DIC concentrations increase with depth by ~1–2 mM relative to seawater values (~2 mM). Sulfate decreases but remains near seawater values (~28 mM) throughout MUC02, MUC12, and the hydrothermal GC10, but drops more clearly toward the bottom of the hydrothermal GC09 (to 26.4 mM) and the cold GC13 (to 23.8 mM). The only minor deviation is Cold Site from the SA. At this site, sulfate and DIC concentrations change more with depth (sulfate drops to 23.6 mM; DIC increases to 6.2 mM), suggesting higher microbial activity relative to all hydrothermal and control sites within the NSA. Consistent with this interpretation sulfide (HS−) concentrations increase strongly with depth at Cold Site (from 2500 to 6200 µM) but not at the NSA sites, where sulfide concentrations remain much lower (0–52 µM (Supplementary Fig. 1). Furthermore, δ13C-DIC decreases with sediment depth at Cold Site (from −3.3‰ to −10.3‰), suggesting strong input of DIC from organic carbon mineralization (Supplementary Fig. 2). By contrast, δ13C-DIC remains close to seawater values (~0‰) throughout sediments of all NSA sites (−1.7‰ to −0.2‰).Compared to all NSA sites and Cold Site, sulfate, methane, and DIC concentrations are more variable at the seep sites Yellow Mat, Cathedral Hill, Orange Mat, and Everest Mound (Fig. 1b, 2nd column). Methane concentrations at Yellow Mat, Cathedral Hill, and Orange Mat are much higher than at the non-seep sites, reaching 3.3, 5.2, and 2.8 mM, respectively (no data from Everest Mound). These high methane concentrations, which can be mainly attributed to the input of thermogenic methane from below24, almost certainly underestimate in situ concentrations due to outgassing during core retrieval. Sulfate concentrations decrease more strongly with depth than at the NSA sites or Control Site, consistent with previously observed high sulfate-reducing activity6,7 and advection of sulfate-depleted fluid from below29. Nonetheless, sulfate concentrations remain in the millimolar range throughout cores from Yellow and Orange Mat. By contrast, sulfate is below detection (≤0.1 mM) at ≥4.5 cm sediment depth at Everest Mound, and in an intermittent depth interval at Cathedral Hill (~7.5–19.5 cm), below which it increases back to ~6 mM. High, i.e. millimolar, concentrations of sulfide at Orange Mat and Cathedral Hill are consistent with high rates of in situ microbial sulfate reduction and advective input of sulfide from the thermochemical reduction of sulfate in hotter, abiotic layers below (Supplementary Fig. 1). DIC concentrations reach values of >10 mM at Orange Mat, Cathedral Hill, and Yellow Mat (no data from Everest Mound). DIC concentrations fluctuate around 20 mM DIC throughout the core from Cathedral Hill, suggesting high DIC input from deeper layers. C-isotopic values of this DIC are close to those of seawater (~−3‰), suggesting an inorganic source. By contrast, surface sedimentary DIC concentrations at Yellow Mat and Orange Mat are close to seawater values but increase with depth to ~20 and ~14 mM, respectively. Lower δ13C-DIC values in surface sediments, which decrease further to values of ~−20‰ to −24‰ at Yellow Mat and −14‰ to −18‰ at Orange Mat within the top 10–20 cm, suggest that most of this DIC comes from the microbial or thermogenic breakdown of organic matter and/or the microbial anaerobic oxidation of methane.Trends in dissolved SCOAs across locationsPorewater concentration profiles of SCOAs are consistent with higher input of reactive organic carbon substrates to hydrothermal seep sediments compared to cold control sites or hydrothermal non-seep sediments.SCOA concentrations at the cold control sites and hot NSA sites are low, showing no clear depth-related trends, consistent with absence of SCOA input from below and/or biological controlled SCOA concentrations. SCOAs are dominated by acetate (cold MUC02, MUC12, and GC13: 1–3 µM; hydrothermal GCs: 3–6 µM; Cold Site: 1–7 µM), which was detected along with formate, propionate, and lactate (Fig. 2).Fig. 2: Depth profiles of short-chain organic acid (SCOA) concentrations across locations.Note the differences in concentration ranges on the x-axis and depth ranges on the y-axis (Cathedral Hill: 0–50 cm; GC13, GC09, and GC10: 0–500 cm; all others: 0–40 cm).Full size imageBy contrast, SCOA concentrations at all hydrothermal seep sites except Orange Mat, increase with depth and temperature, consistent with a thermogenic source below the cored interval. At Yellow Mat, acetate concentrations are already elevated at the seafloor (32 µM) and increase to >100 µM at 20 cm depth. Cathedral Hill has a similar acetate concentration profile, but reaches even higher concentrations (250 µM). At the hottest site, Everest Mound, acetate concentrations increase from ~150 µM at the seafloor to steady concentrations of ~600 µM below 3 cm. Formate concentrations are also (locally) elevated at Yellow Mat (5-8 µM), Cathedral Hill (to 14 µM), and Everest Mound (94-265 µM), and propionate concentrations reach high values at Cathedral Hill (to 21.8 µM) and Everest Mound (to 125 µM). The only exception among the seep sites is Orange Mat, where acetate is only slightly elevated (10–20 µM), and formate and propionate remain at background concentrations. These concentrations suggest that either thermogenic SCOA input from below is low at this site, or SCOA concentrations are biologically controlled throughout the core. Unlike the other three SCOAs, lactate concentrations remain low at all seep sites, apart from one outlier at Cathedral Hill (34.5 cm: 17.3 µM), suggesting that lactate is not a major product of thermogenic organic matter breakdown.Trends in solid-phase organic matter poolsAll sites have similar δ13C-TOC isotopic compositions, with values ranging from −19‰ to −23‰, consistent with a predominant phytoplankton origin of sedimentary organic carbon (Supplementary Fig. 3). Yet, depth profiles of TOC and TN follow different patterns across the locations (Fig. 3). All cold control sites have similar TOC (~2–4 wt%) and TN contents (~0.3–0.6 wt%), with slight decreases in values from the seafloor downward. Compared to cold controls, GC09 and GC10 have lower TOC and TN contents (TOC: ~0.5–3 wt%; TN: ~0.0–0.3 wt%), in particular in deeper horizons with elevated temperatures. Seep sites within the SA have the widest ranges. Seep sites have higher TOC in surface sediment compared to control sites, suggesting net organic carbon assimilation and synthesis by microbial growth. TOC values are 16 wt% at the seafloor of Orange Mat and 6–7 wt% at the seafloor of the other three locations, and then decrease strongly within the top 10 cm, reaching values similar to those of cold sites or hot NSA sites below 10 cm. TN values in surface sediments of seep sites are generally higher than at control sites (~0.7–0.9 wt%), providing additional evidence of net organic matter synthesis by microbial biomass production, but then decrease steeply to values that are similar to those at hot NSA sites.Fig. 3: Carbon and nitrogen contents of bulk organic matter.Depth profiles of total organic carbon (TOC), total nitrogen (TN), and TOC:TN (C:N) across all sites.Full size imageAs a result of the stable TOC and TN trends, C:N does not change much with depth at the cold locations. Yet, while C:N ranges around 4.4–5.6 at Cold Site, values are considerably higher, around 8.1–10.1, at cold locations within the NSA. By comparison, the hot NSA sites and all seep sites except Orange Mat show increases in C:N with increasing temperature and depth. This increase in C:N is modest, from ~8 to 10 at Yellow Mat, and more pronounced at the hotter GC09 (to 15.9), GC10 (to 13.4), Cathedral Hill (to 14.6), and Everest Mound (to 15.7). Orange Mat has the highest C:N ratios (14.8–26.5), and unlike the other sites does not show an increase in C:N with depth.General trends in bacterial and archaeal 16S rRNA gene copy numbers16S rRNA gene copy numbers indicate distinct trends in bacterial and archaeal abundances that follow temperature increases with sediment depth (Fig. 1a and b, 3rd column).At the four cold locations, bacterial and archaeal gene copy numbers are relatively stable with depth (Bacteria: 108−109 g−1; Archaea: 107−108 g−1). By comparison, gene copy numbers of GC09 and GC10 are in a similar range near the seafloor but decrease strongly with depth. While Archaea are quantifiable throughout both cores to ≤103 gene copies g−1 sediment, bacterial gene copy numbers are not reliably distinguishable from extraction negative controls (~1 × 104 g−1) at temperatures >60 °C. Furthermore, unlike the cold sites, which consistently have higher bacterial gene copy numbers, there is a shift from bacterial to archaeal dominance in gene copy numbers (GC09: at ~50 cm; GC10: at ~150 cm) at both hot NSA sites.Compared to the hot GCs from the NSA, gene copies decrease over much shorter distances at sites with fluid seepage in the SA. This decrease in gene copy numbers appears related to the magnitude of the temperature increase with depth. At Yellow Mat, which only reaches moderately warm temperatures (27 °C), copy numbers of both domains decrease from ~108 g−1 at the seafloor to ~106 g−1 at the bottom of the core. While Orange Mat, Cathedral Hill, and Everest Mound have similar bacterial and archaeal gene copy numbers to Yellow Mat at the seafloor, these values drop off much more steeply with depth, matching the much steeper temperature increases. At Cathedral Hill and Everest Mound, Bacteria could not be reliably detected below 20 and 7.5 cm, respectively. As the only location, the detection limit of archaeal 16S gene sequences was reached at Everest Mound, at a depth of 9.5 cm.Relationships between microbial gene abundances and temperatureWe explored the relationship between 16S rRNA gene copy number and temperature further (Fig. 4a, b). While gene copy numbers of both domains generally decrease with increasing temperature, the shape of this temperature relationship differs between both domains. In bacteria the decrease in gene copy numbers in relation to temperature is nearly linear. By contrast, in Archaea gene copy numbers follow hump-shaped distributions, i.e. they remain stable or only decrease slightly up to a certain temperature threshold, beyond which their copy numbers decrease steeply. This apparent thermal threshold varies between sites, i.e. it is ~85 °C at Orange Mat, ~70 °C at Cathedral Hill, ~50 °C at the NSA sites, and ~20 °C at Everest Mound.Fig. 4: Gene copy trends in relation to temperature.a Bacterial and (b) archaeal 16S rRNA gene copy numbers vs. temperature. c Bacteria-to-Archaea 16S rRNA gene copy ratios vs. temperature (the exponential function and its coefficient of determination (R2), both calculated in Microsoft Excel, are shown in the graph). Symbol sizes indicate the sediment depth of each sample. Cold control sites from both locations are grouped together in the legend for easier viewing.Full size imageThe differences in relationships between bacterial and archaeal gene copy numbers and temperature are reflected in Bacteria-to-Archaea gene copy ratios (Fig. 4c). Bacterial always exceed archaeal gene copies at 45 °C. Between 10 and 45 °C, domain-level gene dominance varies with location. Despite the variability, Bacteria-to-Archaea gene copy ratios follow a highly significant, exponential relationship with temperature (R2 = 0.67, p  More

1.Flombaum P, Gallegos JL, Gordillo RA, Rincón J, Zabala LL, Jiao N, et al. Present and future global distributions of the marine cyanobacteria Prochlorococcus and Synechococcus. Proc Natl Acad Sci USA. 2013;110:9824–9.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
2.Goldin S, Hulata Y, Baran N, Lindell D. Quantification of T4-like and T7-like cyanophages using the polony method show they are significant members of the virioplankton in the North Pacific Subtropical Gyre. Front Microbiol. 2020;11:1210.PubMed 
PubMed Central 
Article 

Google Scholar 
3.Waterbury JB, Valois FW. Resistance to co-occurring phages enables marine Synechococcus communities to coexist with cyanophages abundant in seawater. Appl Environ Microbiol. 1993;59:3393–9.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
4.Marston MF, Sallee JL. Genetic diversity and temporal variation in the cyanophage community infecting marine Synechococcus species in Rhode Island’s coastal waters. Appl Environ Microbiol. 2003;69:4639–47.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
5.Clokie MRJ, Mann NH. Marine cyanophages and light. Environ Microbiol. 2006;8:2074–82.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
6.Avrani S, Wurtzel O, Sharon I, Sorek R, Lindell D. Genomic island variability facilitates Prochlorococcus -virus coexistence. Nature. 2011;474:604–8.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
7.Marston MF, Pierciey FJ, Shepard A, Gearin G, Qi J, Yandava C, et al. Rapid diversification of coevolving marine Synechococcus and a virus. Proc Natl Acad Sci USA. 2012;109:4544–9.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
8.Fuhrman JA. Marine virueses and their biogeochemical and ecological effects. Nature. 1999;399:541–8.9.Suttle CA. Marine viruses – major players in the global ecosystem. Nat Rev Microbiol. 2007;5:801–12.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
10.Breitbart M, Bonnain C, Malki K, Sawaya NA. Phage puppet masters of the marine microbial realm. Nat Microbiol. 2018;3:754–66.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
11.Suttle CA, Chan AM. Marine cyanophages infecting oceanic and coastal strains of Synechococcus: abundance, morphology, cross-infectivity and growth characteristics. Mar Ecol Prog Ser. 1993;92:99–109.Article 

Google Scholar 
12.Sullivan MB, Waterbury JB, Chisholm SW. Cyanophages infecting the oceanic cyanobacterium Prochlorococcus. Nature. 2003;424:1047–51.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
13.Sullivan MB, Coleman ML, Weigele P, Rohwer F, Chisholm SW. Three Prochlorococcus cyanophage genomes: Signature features and ecological interpretations. PLoS Biol. 2005;3:0790–806.CAS 
Article 

Google Scholar 
14.Pope WH, Weigele PR, Chang J, Pedulla ML, Ford ME, Houtz JM, et al. Genome sequence, structural proteins, and capsid organization of the cyanophage Syn5: a “horned” bacteriophage of marine Synechococcus. J Mol Biol. 2007;368:966–81.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
15.Weigele PR, Pope WH, Pedulla ML, Houtz JM, Smith AL, Conway JF, et al. Genomic and structural analysis of Syn9, a cyanophage infecting marine Prochlorococcus and Synechococcus. Environ Microbiol. 2007;9:1675–95.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
16.Labrie SJ, Frois-Moniz K, Osburne MS, Kelly L, Roggensack SE, Sullivan MB, et al. Genomes of marine cyanopodoviruses reveal multiple origins of diversity. Environ Microbiol. 2013;15:1356–76.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
17.Huang S, Wang K, Jiao N, Chen F. Genome sequences of siphoviruses infecting marine Synechococcus unveil a diverse cyanophage group and extensive phage-host genetic exchanges. Environ Microbiol. 2012;14:540–58.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
18.Sullivan MB, Huang KH, Ignacio-Espinoza JC, Berlin AM, Kelly L, Weigele PR, et al. Genomic analysis of oceanic cyanobacterial myoviruses compared with T4-like myoviruses from diverse hosts and environments. Environ Microbiol. 2010;12:3035–56.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
19.Ghai R, Martin-Cuadrado AB, Molto AG, Heredia IG, Cabrera R, Martin J, et al. Metagenome of the Mediterranean deep chlorophyll maximum studied by direct and fosmid library 454 pyrosequencing. ISME J. 2010;4:1154–66.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
20.Ma Y, Allen LZ, Palenik B. Diversity and genome dynamics of marine cyanophages using metagenomic analyses. Environ Microbiol Rep. 2014;6:583–94.PubMed 
Article 
PubMed Central 

Google Scholar 
21.Sabehi G, Shaulov L, Silver DH, Yanai I, Harel A, Lindell D. A novel lineage of myoviruses infecting cyanobacteria is widespread in the oceans. Proc Natl Acad Sci USA. 2012;109:2037–42.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
22.Huang S, Zhang S, Jiao N, Chen F. Comparative genomic and phylogenomic analyses reveal a conserved core genome shared by estuarine and oceanic cyanopodoviruses. PLoS One. 2015;10:1–17.
Google Scholar 
23.Ignacio-espinoza JC, Sullivan MB. Phylogenomics of T4 cyanophages: lateral gene transfer in the ‘core’ and origins of host genes. Environ Microbiol. 2012;14:2113–26.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
24.Crummett LT, Puxty RJ, Weihe C, Marston MF, Martiny JBHH. The genomic content and context of auxiliary metabolic genes in marine cyanomyoviruses. Virology. 2016;499:219–29.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
25.Millard AD, Zwirglmaier K, Downey MJ, Mann NH, Scanlan DJ. Comparative genomics of marine cyanomyoviruses reveals the widespread occurrence of Synechococcus host genes localized to a hyperplastic region: Implications for mechanisms of cyanophage evolution. Environ Microbiol. 2009;11:2370–87.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
26.Mann NH, Cook A, Millard A, Bailey S, Clokie M. Bacterial photosynthesis genes in a virus. Nature 2003;424:741–741.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
27.Lindell D, Sullivan MB, Johnson ZI, Tolonen AC, Rohwer F, Chisholm SW. Transfer of photosynthesis genes to and from Prochlorococcus viruses. Proc Natl Acad Sci USA. 2004;101:11013–8.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
28.Puxty RJ, Millard AD, Evans DJ, Scanlan DJ. Viruses inhibit CO2 fixation in the most abundant phototrophs on Earth. Curr Biol. 2016;26:1585–9.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
29.Kelly L, Ding H, Huang KH, Osburne MS, Chisholm SW. Genetic diversity in cultured and wild marine cyanomyoviruses reveals phosphorus stress as a strong selective agent. ISME J. 2013;7:1827–41.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
30.Yin Y, Fischer D. Identification and investigation of ORFans in the viral world. BMC Genomics. 2008;9:24.PubMed 
PubMed Central 
Article 
CAS 

Google Scholar 
31.Clokie MR, Millard AD, Mann NH. T4 genes in the marine ecosystem: studies of the T4-like cyanophages and their role in marine ecology. Virol J. 2010;7:291.PubMed 
PubMed Central 
Article 
CAS 

Google Scholar 
32.Rihtman B, Bowman‐Grahl S, Millard A, Corrigan RM, Clokie MRJ, Scanlan DJ. Cyanophage MazG is a pyrophosphohydrolase but unable to hydrolyse magic spot nucleotides. Environ Microbiol Rep. 2019;11:448–55.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
33.Dammeyer T, Bagby SC, Sullivan MB, Chisholm SW, Frankenberg-Dinkel N. Efficient phage-mediated pigment biosynthesis in oceanic cyanobacteria. Curr Biol. 2008;18:442–8.CAS 
PubMed 
Article 

Google Scholar 
34.Roitman S, Hornung E, Flores-Uribe J, Sharon I, Feussner I, Béjà O. Cyanophage-encoded lipid desaturases: Oceanic distribution, diversity and function. ISME J. 2018;12:343–55.CAS 
PubMed 
Article 

Google Scholar 
35.Thompson LR, Zeng Q, Kelly L, Huang KH, Singer AU, Stubbe J, et al. Phage auxiliary metabolic genes and the redirection of cyanobacterial host carbon metabolism. Proc Natl Acad Sci USA. 2011;108:E757–64.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
36.Puxty RJ, Perez-Sepulveda B, Rihtman B, Evans DJ, Millard AD, Scanlan DJ. Spontaneous deletion of an “ORFanage” region facilitates host adaptation in a “photosynthetic” cyanophage. PLoS One. 2015;10:e0132642.PubMed 
PubMed Central 
Article 
CAS 

Google Scholar 
37.Howard-Varona C, Hargreaves KR, Abedon ST, Sullivan MB. Lysogeny in nature: mechanisms, impact and ecology of temperate phages. ISME J. 2017;11:1511–20.PubMed 
PubMed Central 
Article 

Google Scholar 
38.Ranade K, Poteete AR. Superinfection exclusion (sieB) genes of bacteriophages P22 and λ. J Bacteriol. 1993;175:4712–8.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
39.Fogg PCM, Allison HE, Saunders JR, McCarthy AJ. Bacteriophage Lambda: a paradigm revisited. J Virol. 2010;84:6876–9.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
40.van Houte S, Buckling A, Westra ER. Evolutionary ecology of prokaryotic immune mechanisms. Microbiol Mol Biol Rev. 2016;80:745–63.PubMed 
PubMed Central 
Article 

Google Scholar 
41.Tuttle MJ, Buchan A. Lysogeny in the oceans: lessons from cultivated model systems and a reanalysis of its prevalence. Environ Microbiol. 2020;22:4919–33.PubMed 
Article 
PubMed Central 

Google Scholar 
42.Knowles B, Silveira CB, Bailey BA, Barott K, Cantu VA, Cobian-Guëmes AG, et al. Lytic to temperate switching of viral communities. Nature. 2016;531:466–70.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
43.Touchon M, Bernheim A, Rocha EPC. Genetic and life-history traits associated with the distribution of prophages in bacteria. ISME J. 2016;10:2744–54.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
44.Roux S, Hallam SJ, Woyke T, Sullivan MB. Viral dark matter and virus–host interactions resolved from publicly available microbial genomes. Elife. 2015;4:e08490.PubMed Central 
Article 
PubMed 

Google Scholar 
45.Luo E, Eppley JM, Romano AE, Mende DR, DeLong EF. Double-stranded DNA. virioplankton dynamics and reproductive strategies in the oligotrophic open ocean water column. ISME J. 2020;14:1304–15.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
46.Touchon M, Moura de Sousa JA, Rocha EP. Embracing the enemy: the diversification of microbial gene repertoires by phage-mediated horizontal gene transfer. Curr Opin Microbiol. 2017;38:66–73.CAS 
PubMed 
Article 

Google Scholar 
47.Coleman ML, Sullivan MB, Martiny AC, Steglich C, Barry K, Delong EF, et al. Genomic islands and the ecology and evolution of Prochlorococcus. Science. 2006;311:1768–70.CAS 
PubMed 
Article 

Google Scholar 
48.Zhaxybayeva O, Gogarten JP, Charlebois RL, Doolittle WF, Papke RT. Phylogenetic analyses of cyanobacterial genomes: quantification of horizontal gene transfer events. Genome Res. 2006;16:1099–108.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
49.Lindell D, Jaffe JD, Coleman ML, Futschik ME, Axmann IM, Rector T, et al. Genome-wide expression dynamics of a marine virus and host reveal features of co-evolution. Nature. 2007;449:83–6.CAS 
PubMed 
Article 

Google Scholar 
50.Raytcheva DA, Haase-Pettingell C, Piret JM, King JA. Intracellular assembly of cyanophage Syn5 proceeds through a scaffold-containing procapsid. J Virol. 2011;85:2406–15.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
51.Flores-Uribe J, Philosof A, Sharon I, Fridman S, Larom S, Béjà O. A novel uncultured marine cyanophage lineage with lysogenic potential linked to a putative marine Synechococcus ‘relic’ prophage. Environ Microbiol Rep. 2019;11:598–604.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
52.Malmstrom RR, Rodrigue S, Huang KH, Kelly L, Kern SE, Thompson A, et al. Ecology of uncultured Prochlorococcus clades revealed through single-cell genomics and biogeographic analysis. ISME J. 2013;7:184–98.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
53.Kiro R, Shitrit D, Qimron U. Efficient engineering of a bacteriophage genome using the type I-E CRISPR-Cas system. RNA Biol. 2014;11:42–4.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
54.Sarkis GJ, Jacobs WR, Hatfull GF. L5 luciferase reporter mycobacteriophages: a sensitive tool for the detection and assay of live mycobacteria. Mol Microbiol. 1995;15:1055–67.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
55.Tanji Y, Furukawa C, Na SH, Hijikata T, Miyanaga K, Unno H. Escherichia coli detection by GFP-labeled lysozyme-inactivated T4 bacteriophage. J Biotechnol. 2004;114:11–20.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
56.Brahamsha B. A genetic manipulation system for oceanic cyanobacteria of the genus Synechococcus. Appl Environ Microbiol. 1996;62:1747–51.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
57.Mahichi F, Synnott AJ, Yamamichi K, Osada T, Tanji Y. Site-specific recombination of T2 phage using IP008 long tail fiber genes provides a targeted method for expanding host range while retaining lytic activity. FEMS Microbiol Lett. 2009;295:211–7.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
58.Le S, He X, Tan Y, Huang G, Zhang L, Lux R, et al. Mapping the tail fiber as the receptor binding protein responsible for differential host specificity of Pseudomonas aeruginosa bacteriophages PaP1 and JG004. PLoS One. 2013;8:e68562.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
59.Murphy KC. Phage recombinases and their applications. Adv Virus Res. 2012;83:367–414.60.Bujarski JJ. Recombination of viruses. In: Encyclopedia of Virology. Elsevier; 1999. p. 1446–53.61.Pires DP, Cleto S, Sillankorva S, Azeredo J, Lu TK. Genetically engineered phages: a review of advances over the last decade. Microbiol Mol Biol Rev. 2016;80:523–43.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
62.Qimron U, Marintcheva B, Tabor S, Richardson CC. Genomewide screens for Escherichia coli genes affecting growth of T7 bacteriophage. Proc Natl Acad Sci USA. 2006;103:19039–44.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
63.Selick HE, Kreuzer KN, Alberts BM. The bacteriophage T4 insertion/substitution vector system. A method for introducing site-specific mutations into the virus chromosome. J Biol Chem. 1988;263:11336–47.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
64.Martel B, Moineau S. CRISPR-Cas: an efficient tool for genome engineering of virulent bacteriophages. Nucleic Acids Res. 2014;42:9504–13.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
65.Dale JW, Greenaway PJ. Identification of recombinant phages by plaque hybridization. In: Walker JM, editor. Nucleic Acids. Totowa, NJ: Humana Press; 1984. p. 285–8.Chapter 

Google Scholar 
66.Dekel-Bird NP, Avrani S, Sabehi G, Pekarsky I, Marston MF, Kirzner S, et al. Diversity and evolutionary relationships of T7-like podoviruses infecting marine cyanobacteria. Environ Microbiol. 2013;15:1476–91.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
67.Doron S, Fedida A, Hernndez-Prieto MA, Sabehi G, Karunker I, Stazic D, et al. Transcriptome dynamics of a broad host-range cyanophage and its hosts. ISME J. 2016;10:1437–55.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
68.Krupovic M, Forterre P. Single-stranded DNA viruses employ a variety of mechanisms for integration into host genomes. Ann NY Acad Sci. 2015;1341:41–53.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
69.Morris RM, Cain KR, Hvorecny KL, Kollman JM. Lysogenic host–virus interactions in SAR11 marine bacteria. Nat Microbiol. 2020;5:1011–5.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
70.Martínez-García E, Jatsenko T, Kivisaar M, de Lorenzo V. Freeing Pseudomonas putida KT2440 of its proviral load strengthens endurance to environmental stresses. Environ Microbiol. 2015;17:76–90.PubMed 
Article 
CAS 
PubMed Central 

Google Scholar 
71.Mruwat N, Carlson MCG, Goldin S, Ribalet F, Kirzner S, Hulata Y, et al. A single-cell polony method reveals low levels of infected Prochlorococcus in oligotrophic waters despite high cyanophage abundances. ISME J. 2021;15:41–54.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
72.Oppenheim AB, Kobiler O, Stavans J, Court DL, Adhya S. Switches in bacteriophage lambda development. Annu Rev Genet. 2005;39:409–29.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
73.Ray U, Sakalka A. Lysogenization of Escherichia coli by bacteriophage Lambda: complementary activity of the host’s DNA polymerase I and ligase and bacteriophage replication proteins Q and P. J Virol. 1976;18:511–7.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
74.Greengrass E. Resistance of marine Synechococcus to podovirus infection: genetic basis and phenotypic characterization. M.Sc. thesis. Technion – Israel Inst Technol. 2013.75.Fedida A, Lindell D. Two Synechococcus genes, two different effects on cyanophage infection. Viruses. 2017;9:136.PubMed Central 
Article 
CAS 

Google Scholar 
76.Shao Q, Trinh JT, McIntosh CS, Christenson B, Balázsi G, Zeng L. Lysis-lysogeny coexistence: prophage integration during lytic development. Microbiol Open. 2017;6:e00395.Article 
CAS 

Google Scholar 
77.Chen F, Lu J. Genomic sequence and evolution of marine cyanophage P60: a new insight on lytic and lysogenic phages. Appl Environ Microbiol. 2002;68:2589–94.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
78.Brum JR, Hurwitz BL, Schofield O, Ducklow HW, Sullivan MB. Seasonal time bombs: dominant temperate viruses affect Southern Ocean microbial dynamics. ISME J. 2016;10:437–49.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
79.Pratama AA, van Elsas JD. The ‘neglected’ soil virome—potential role and impact. Trends Microbiol. 2018;26:649–62.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
80.Srinivasiah S, Bhavsar J, Thapar K, Liles M, Schoenfeld T, Wommack KE. Phages across the biosphere: contrasts of viruses in soil and aquatic environments. Res Microbiol. 2008;159:349–57.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
81.Zhao Y, Temperton B, Thrash JC, Schwalbach MS, Vergin KL, Landry ZC, et al. Abundant SAR11 viruses in the ocean. Nature. 2013;494:357–60.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
82.Zhao Y, Qin F, Zhang R, Giovannoni SJ, Zhang Z, Sun J, et al. Pelagiphages in the Podoviridae family integrate into host genomes. Environ Microbiol. 2018;21:1989–2001.PubMed 
Article 
CAS 
PubMed Central 

Google Scholar 
83.Kashtan N, Roggensack SE, Berta-Thompson JW, Grinberg M, Stepanauskas R, Chisholm SW. Fundamental differences in diversity and genomic population structure between Atlantic and Pacific Prochlorococcus. ISME J. 2017;11:1997–2011.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
84.Berube PM, Biller SJ, Hackl T, Hogle SL, Satinsky BM, Becker JW, et al. Single cell genomes of Prochlorococcus, Synechococcus, and sympatric microbes from diverse marine environments. Sci Data. 2018;5:1–11.Article 
CAS 

Google Scholar 
85.Wyman M, Gregory RPF, Carr NG. Novel role for phycoerythrin in a marine cyanobacterium, Synechococcus strain DC2. Science. 1985;230:818–20.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
86.Lindell D, Padan E, Post AF. Regulation of ntcA expression and nitrite uptake in the marine Synechococcus sp. strain WH 7803. J Bacteriol. 1998;180:1878–86.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
87.Moore LR, Coe A, Zinser ER, Saito MA, Sullivan MB, Lindell D, et al. Culturing the marine cyanobacterium Prochlorococcus. Limnol Oceanogr Methods. 2007;5:353–62.CAS 
Article 

Google Scholar 
88.Lindell D. The genus Prochlorococcus, phylum cyanobacteria. In: The Prokaryotes: Other Major Lineages of Bacteria and The Archaea. Springer-Verlag Berlin Heidelberg; 2014. p. 829–45.89.Morris JJ, Kirkegaard R, Szul MJ, Johnson ZI, Zinser ER. Facilitation of robust growth of Prochlorococcus colonies and dilute liquid cultures by “helper” heterotrophic bacteria. Appl Environ Microbiol. 2008;74:4530–4.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
90.Wolk CP, Fan Q, Zhou R, Huang G, Lechno-Yossef S, Kuritz T, et al. Paired cloning vectors for complementation of mutations in the cyanobacterium Anabaena sp. strain PCC 7120. Arch Microbiol. 2007;188:551–63.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
91.Bryksin A, Matsumura I. Overlap extension PCR cloning: a simple and reliable way to create recombinant plasmids. Biotechniques. 2010;48:463–5.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
92.Simon R, Priefer U, Pühler A. A broad host range mobilization system for in vivo genetic engineering: Transposon mutagenesis in gram negative bacteria. Nat Biotechnol. 1983;1:784–91.CAS 
Article 

Google Scholar 
93.Henn MR, Sullivan MB, Stange-Thomann N, Osburne MS, Berlin AM, Kelly L, et al. Analysis of high-throughput sequencing and annotation strategies for phage genomes. PLoS One. 2010;5:e9083.PubMed 
PubMed Central 
Article 
CAS 

Google Scholar 
94.Zinser ER, Coe A, Johnson ZI, Martiny AC, Fuller NJ, Scanlan DJ, et al. Prochlorococcus ecotype abundances in the North Atlantic Ocean as revealed by an improved quantitative PCR method. Appl Environ Microbiol. 2006;72:723–32.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
95.Lindell D, Jaffe JD, Johnson ZI, Church GM, Chisholm SW. Photosynthesis genes in marine viruses yield proteins during host infection. Nature. 2005;438:86–9.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
96.Mitra RD, Church GM. In situ localized amplification and contact replication of many individual DNA molecules. Nucleic Acids Res. 1999;27:e34.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
97.Schwartz DA, Lindell D. Genetic hurdles limit the arms race between Prochlorococcus and the T7-like podoviruses infecting them. ISME J. 2017;11:1836–51.PubMed 
PubMed Central 
Article 

Google Scholar 
98.Bull JJ, Badgett MR, Wichman HA, Huelsenbeck JP, Hillis DM, Gulati A, et al. Exceptional convergent evolution in a virus. Genetics. 1997;147:1497–507.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
99.Quinlan AR. BEDTools: the Swiss-army Tool for genome feature analysis. Curr Protoc Bioinform. 2014;47:1–34. 11.12.Article 

Google Scholar 
100.Biller SJ, Berube PM, Dooley K, Williams M, Satinsky BM, Hackl T, et al. Marine microbial metagenomes sampled across space and time. Sci Data. 2018;5:180176.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
101.Haro-Moreno JM, López-Pérez M, de la Torre JR, Picazo A, Camacho A, Rodriguez-Valera F. Fine metagenomic profile of the Mediterranean stratified and mixed water columns revealed by assembly and recruitment. Microbiome. 2018;6:128.PubMed 
PubMed Central 
Article 

Google Scholar 
102.Philosof A, Yutin N, Flores-Uribe J, Sharon I, Koonin EV, Béjà O. Novel abundant oceanic viruses of uncultured marine group II Euryarchaeota. Curr Biol. 2017;27:1362–8.CAS 
PubMed 
PubMed Central 
Article 

Google Scholar 
103.Wilson ST, Aylward FO, Ribalet F, Barone B, Casey JR, Connell PE, et al. Coordinated regulation of growth, activity and transcription in natural populations of the unicellular nitrogen-fixing cyanobacterium Crocosphaera. Nat Microbiol. 2017;2:17118.CAS 
PubMed 
Article 
PubMed Central 

Google Scholar 
104.Sunagawa S, Coelho LP, Chaffron S, Kultima JR, Labadie K, Salazar G, et al. Structure and function of the global ocean microbiome. Science. 2015;348:1261359.PubMed 
Article 
CAS 
PubMed Central 

Google Scholar 
105.Bushnell B. BBMap: a fast, accurate, splice-aware aligner. 2014;URL: https://www.osti.gov/servlets/purl/1241166.106.Li H. Aligning sequence reads, clone sequences and assembly contigs with BWA-MEM. arXiv. 2013;1303:3997.
Google Scholar 
107.Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, et al. The sequence alignment/map format and SAMtools. Bioinformatics. 2009;25:2078–9.PubMed 
PubMed Central 
Article 
CAS 

Google Scholar  More