Bacteria acquire phosphate (Pi) by maintaining a periplasmic concentration below environmental levels. We recently described an extracellular Pi buffer which appears to counteract the gradient required for Pi diffusion. Here, we demonstrate that various treatments to outer membrane (OM) constituents do not affect the buffered Pi because bacteria accumulate Pi in the periplasm, from which it can be removed hypo-osmotically. The periplasmic Pi can be gradually imported into the cytoplasm by ATP-powered transport, however, the proton motive force (PMF) is not required to keep Pi in the periplasm. In contrast, the accumulation of Pi into the periplasm across the OM is PMF-dependent and can be enhanced by light energy. Because the conventional mechanism of Pi-specific transport cannot explain Pi accumulation in the periplasm we propose that periplasmic Pi anions pair with chemiosmotic cations of the PMF and millions of accumulated Pi pairs could influence the periplasmic osmolarity of marine bacteria.
Phosphorus (P) is a macronutrient universally required by all organisms including bacteria. Bacteria have a number of ways to acquire P. Scavenging P from organic molecules and reducing growth dependence on P by substituting P with sulfur in their membranes1,2,3,4,5,6 are both beneficial for bacteria living in phosphate (Pi)-depleted marine environments. However, there is compelling evidence that bacteria prefer Pi to organic P molecules7,8,9. Marine bacteria, which typically have two membranes (and therefore stain Gram negative), acquire Pi using a diffusion gradient between the environment and the periplasm10,11. In their cells, the periplasm is separated from the environment by the outer membrane (OM), whose permeability to small, hydrophilic solutes is controlled mainly by hydrated channels—porins (Fig. 1a†). Porins select solutes on the basis of their size, shape and charge12,13. When nutrients such as Pi become depleted, cells increase the number of porins to facilitate nutrient diffusion14.
There are the three known bacterial transport systems to import Pi from the periplasm into the cytoplasm: a low affinity-high velocity phosphate inorganic transport (Pit) system, a low affinity-high velocity Na-dependent phosphate transport (Npt) system and a high affinity-low velocity phosphate-specific transport (Pst) system15,16,17. Bacteria living in Pi-depleted waters use only the Pst system18,19. The PstCAB transporter is ATP-powered and receives Pi from a carrier protein, a PstS subunit, when the latter docks at its periplasmic side (Fig. 1a†). Although the Pi concentration of ~10−7 mol l−1 required for efficient import of Pi by PstSCAB20 is relatively high, it should not restrict Pi diffusion into the periplasm, because the periplasmic volume of even a relatively large bacterial cell, e.g., Synechococcus cyanobacteria (Supplementary Table 1) with an estimated periplasmic depth21 of 10−8 m, is only about 2 × 10−17 l. In such a tiny volume, the presence of only a few free Pi molecules would exceed the threshold 10−7 mol l−1 Pi concentration (Fig. 1a†).
Bacteria maximize the diffusive flux of nutrients through the OM by maintaining a steep nutrient concentration gradient between the environment and periplasm10,11. Consequently, to allow efficient diffusion of Pi into the periplasm in Pi-depleted (10−9–10−8 mol l−1) oceanic surface waters7,22, the periplasmic Pi concentration should hypothetically be <10−9 mol l−1. This means that there should be no free Pi molecules in the periplasm, i.e., every Pi molecule entering the periplasm should be instantly bound by a PstS subunit requiring an affinity >100 times above the known PstS affinity limit. However, the PstS affinity requirement does not seem to limit the growth of ubiquitous SAR11 alphaproteobacteria and Prochlorococcus cyanobacteria—the two bacterial populations comprising ≤¾ of oceanic surface bacterioplankton in the Pi-depleted North Atlantic subtropical gyre7. Furthermore, the ecological success of these bacteria is probably related to the high rates of Pi uptake measured in the gyre7,22. Surprisingly, measured rates of Pi acquisition by Prochlorococcus and SAR11 are lower in tropical surface waters where bacterial growth and Pi concentrations are higher22. The counter-intuitive reduction in the Pi acquisition rate by faster growing bacteria was attributed to the presence of an intermediate buffer, in which both SAR11 and Prochlorococcus cells store Pi: the fuller the Pi buffer is, e.g., in Pi-replete tropical surface waters, the fewer Pi molecules a cell acquires slower from seawater to top the buffer up22. As every Pi molecule acquired, or more precisely accumulated, by a cell first enters the buffer before being imported for assimilation, bacterial Pi uptake into the buffer and Pi import from the buffer can be decoupled. Although Prochlorococcus and SAR11 bacteria possess genes related to synthesis of polyphosphates23, their cells are perhaps too small to store Pi as polyphosphates intracellularly and they somehow accumulate Pi extracellularly.
Our aim here is to explain how marine bacteria accumulate Pi extracellularly. We demonstrate that Pi accumulation rates by oceanic SAR11, Prochlorococcus and Synechococcus cells are extremely variable in the Pi-depleted North Atlantic. Lower rates can be stimulated by light energy but their maximal rates are insensitive to light and approach the theoretical upper limit of diffusion. To relate the accumulated Pi with cellular Pi requirements, the P contents of flow-sorted cells of oceanic Prochlorococcus and SAR11 were determined by microbeam synchrotron X-ray fluorescence (μ-SXRF) using flow-sorted cultured Synechococcus cells for calibration. Using pulse-chase experiments with 33Pi and 32Pi (*Pi) radiotracers, we showed that cultured Synechococcus cells accumulate Pi in a similar way to SAR11 and Prochlorococcus cells22. This justified the use of Synechococcus isolates for more intrusive tests (e.g., of retention of accumulated Pi), to curtail artefacts resulting from the low tolerance of oceanic SAR11 and Prochlorococcus bacteria to harsh experimental manipulations. Moreover, by working with artificial seawater (ASW), we expanded the experimental range of Pi concentrations below those encountered in natural Pi-depleted seawater. To examine how the extracellular Pi is retained, we developed a living cellular model system with the extracellular *Pi-labelled buffer (Fig. 1a) and treated them with hydrolytic enzymes, surfactants, inhibitors, amended ASW and hypotonic solutions to locate whether the Pi buffer is on the cell surface (Fig. 1b), e.g., retained by a charged S-layer, as was recently proposed for archaea24, or in the periplasm (Fig. 1c). We used specific metabolic inhibitors25,26,27,28 to determine whether the proton motive force (PMF) is driving extracellular Pi accumulation and retention.
Based on our field and laboratory experiments, we conclude that the PMF is essential for Pi accumulation (but not retention) in the periplasm of marine bacteria. As the PstS-driven import cannot explain Pi accumulation in the periplasm, we propose a mechanism of periplasmic ionic pairing of inwardly diffusing Pi anions with chemiosmotic cations of the membrane potential (Fig. 1a‡), which could conserve cellular energy and control periplasmic turgor.
Bacterial clearance rates of Pi
Pi clearance rates (the volume of water cleared of Pi by a cell per unit of time, while the cell accumulates the cleared Pi) of oceanic SAR11, Prochlorococcus and Synechococcus cells vary by orders of magnitude in the patchily Pi-depleted North Atlantic (Supplementary Figs. 1 and 2a, and Supplementary Table 2). Living in such a patchy environment warrants the use of an extracellular buffer, where Pi is temporarily stored22. Mostly the measured rates are lower in waters adjacent to the subtropical gyre with higher in situ Pi concentrations, whereas the rates are higher in the central gyre waters with lower Pi concentrations (18–28°N, Supplementary Fig. 2a and Supplementary Table 2). In the central gyre waters, maximal Pi clearance rates of SAR11, Prochlorococcus and Synechococcus cells (Fig. 2a) are only 30×, 40× and 25×, respectively, below the theoretical maximal rate of nutrient acquisition by diffusion10,11. Apparently, a Synechococcus cell can accumulate Pi faster than either a Prochlorococcus or a SAR11 cell. To understand why the abundance of Synechococcus in the central gyre waters is three orders of magnitude lower than the abundance of Prochlorococcus and SAR11, we need to take into account cell sizes and cellular P requirements of these bacteria.
Once their maximal Pi clearance rates are normalized to cellular biovolumes (Supplementary Table 1 and Fig. 2a) then a Prochlorococcus, a Synechococcus or a SAR11 cell can theoretically clear all dissolved Pi from seawater ≤2.5 × 104, 2.6 × 104 or 1.2 × 105 times its cellular volume min−1, respectively. In agreement with the surface area-to-volume concept29, these biovolume-specific clearance rates show that a SAR11 cell is noticeably (4–5×) more effective in uptake of Pi than either a Prochlorococcus cell or a Synechococcus cell probably owing to the 2× higher surface area-to-volume ratio of a small, curved-rod shape SAR11 cell compared with the cocci cyanobacteria (Supplementary Fig. 3 and Supplementary Table 1).
To account for the cellular Ps requirements of oceanic bacterial cells, we determined their cellular P content. The cellular P content of a Synechococcus cell (1.93 × 107 P atoms cell−1) and hence their P demand is 5–7× higher than the P content of a Prochlorococcus and a SAR11 cell (4.14 and 2.68 × 106 P atoms cell−1, respectively, Fig. 2b and Supplementary Table 3). This implies that Synechococcus growth in Pi-depleted waters is constrained by its high P demand rather than by its ability to acquire Pi. It is also notable that a larger Synechococcus and a smaller Prochlorococcus have similar biovolume-specific Pi clearance rates, despite the 1.2× higher cell surface-to-volume ratio of the latter (Supplementary Table 1). As porin-restricted diffusion constrains the Pi clearance rate, the OM of Synechococcus ought to be more permeable to Pi, e.g., to have more Pi-specific porins per µm2 of the OM surface.
The above comparisons show that despite its high P demand, a Synechococcus cell is a good model system for studying the process of Pi acquisition by oceanic bacteria. In our laboratory studies, we focused on the oligotrophic Synechococcus sp. strain WH8102 but also made comparisons with the mesotrophic Synechococcus strains WH8109 and WH780330. The 40–90× lower Pi clearance rate of larger cells of cultured Synechococcus compared with oceanic bacterioplankton suggests that Synechococcus cells grown at a Pi concentration >10−5 mol l−1 have fewer porins12,13 than oceanic bacteria even when the Pi concentration is downshifted (Fig. 2a). Such porin-restricted diffusion explains why a cultured Synechococcus cell cleared Pi from about the same volume of water, while the Pi concentration ranged over more than seven orders of magnitude. The Pi clearance rate of cultured Synechococcus cells remained relatively constant between the three strains down to the low end of the tested Pi concentrations (Fig. 2a), such that within a few hours these cells removed virtually all 33Pi or 32Pi tracers from the medium (Supplementary Table 4). Using carrier-free 32Pi tracer, we determined that the final Pi concentration in the surrounding seawater was depleted by Synechococcus to 10−15 mol l−1 (the instrument detection limit), or six orders of magnitude below oceanic Pi levels (Supplementary Table 2).
Fast growth-decoupled accumulation of Pi
The rate of Pi accumulation (the amount of Pi molecules accumulated by a cell per unit of time) by cultured Synechococcus cells was proportional to the Pi concentration over a range of concentrations from 10−13 to 10−6 mol l−1 (Fig. 2b), much wider than the range of 10−9–10−8 mol l−1 offered by the Pi-depleted North Atlantic (Supplementary Table 2). At Pi concentrations similar to Pi-depleted oceanic waters, cultured cells accumulated Pi orders of magnitude slower than oceanic SAR11, Prochlorococcus and Synechococcus cells (Fig. 2b and Supplementary Fig. 2b). At 10−6 mol Pi l−1, a cultured Synechococcus cell can, however, double its cellular P content within merely 3 h (Fig. 2b and Supplementary Table 3), while they divide once every 1–2 days. Similarly, when related to their cellular P content, oceanic Prochlorococcus and SAR11 cells obtained sufficient Pi for cell division in merely 1–2 h, whereas in such oligotrophic waters Prochlorococcus divide only once every 3.5 days31 and SAR11 once every 2–3 days (calculated based on their cellular leucine uptake rates32). This decoupling of Pi accumulation and growth rates in Synechococcus, Prochlorococcus and SAR11 cells suggests that, apart from being a reserve for steady growth, the accumulated Pi could have an additional, important cellular function.
Synechococcus cells accumulate Pi extracellularly
To assess whether cultured Synechococcus cells accumulate Pi in the extracellular buffer, the accumulation of the 33Pi-pulse and 32Pi-chase was determined in live WH8102 cells (total Pi, Fig. 1a) and in their cellular macromolecules (biomass fixed with paraformaldehyde (PFA) that immobilizes DNA and RNA—the two principal Pi-containing macromolecules—as well as some periplasmic constituents, including PstS subunits) (Fig. 1d). After addition of the 10−6 mol l−1 32Pi-chase, accumulation of the 33Pi-pulse into cellular macromolecules should be halted, because any of the 4.5–7 × 10−9 mol l−1 33Pi-pulse that remains in the seawater would be diluted with the chase >100×. However, similar to that observed in oceanic bacteria22, the pulsed 33Pi continued to be assimilated into cell macromolecules at an unchanged rate, which started slowing down only after 5 h (Fig. 3a, c). Usually in pulse-chase experiments, incorporation of a pulsed molecule (e.g., amino acid or nucleoside) into bacterial macromolecules halts abruptly upon dilution with the chase, because the intermediate cellular labile pool of the pulsed molecule is small, e.g., 5% of the tracer incorporated into macromolecules33. In the case of Pi, that labile pool was 10×–20× larger than the amount assimilated into cellular macromolecules (Fig. 3b, d at the 3 h time point).
We then used the difference in *Pi label between the total Pi in live cells and macromolecule-bound Pi to estimate the amount of labile Pi in three Synechococcus strains. This showed the total *Pi content of tested live cells (Fig. 4a) to be consistently several times higher than the *Pi content of cellular macromolecules (fixed with either PFA or trichloroacetic acid (TCA), Fig. 1d,e and Supplementary Fig. 4). Furthermore, in the *Pi pre-labelled cells, which were washed and suspended in ASW with no *Pi (see Methods), the *Pi content in macromolecules increased with time and approached the total *Pi that remained stable (Fig. 4b). With no *Pi in the medium, the additional *Pi gained by the macromolecules can only be sourced from *Pi accumulated in the intermediate buffer. These data also show that the accumulated *Pi is semi-labile and can be imported into the cytoplasm to synthesize nucleic acids.
To confirm the extracellular (i.e., not in the cytoplasm) location of the labile Pi, we washed cells for 1–2 min with deionized water (DW). Washing live cells with DW effectively strips Pi and other ions adsorbed to the OM components. In addition, such brief hypo-osmotic shock also releases the content of the periplasm without disrupting the plasma membrane. Western blotting analysis confirmed the presence of the periplasmic Pi-binding subunits PstS in the DW-washed fraction but detected no Rubisco, arguably the most abundant soluble protein in the cyanobacterial cytoplasm (Fig. 5a, b). In fact, even a 30 min incubation of Synechococcus in DW did not lyse the cells (Fig. 5c). As a substantial fraction of the accumulated *Pi was removed with a brief rinse of live Synechococcus with DW (Fig. 3), we therefore conclude that their cells accumulate Pi outside the plasma membrane, i.e., on the surface of the OM or in the periplasm (Fig. 1a†).
Periplasmic location and retention of the Pi in Synechococcus
To identify the exact location of the accumulated Pi and to assess whether Pi was associated with specific cell surface ligands, we examined the response of live *Pi pre-labelled Synechococcus cells to a variety of treatments, which are inapplicable to delicate oceanic bacteria (Fig. 1b, c and Supplementary Tables 5 and 6). Treatments with surfactants released no *Pi (Fig. 6), suggesting that it is not associated directly with OM lipids. High concentrations of a broad-spectrum protease Proteinase K, which was shown to efficiently digest extracellular proteins in bacteria34,35, did not release the accumulated *Pi, implying no direct involvement of extracellular proteins (including the S-layer proteins of WH810236) in Pi accumulation (Fig. 6). Release of the accumulated Pi did not happen after destabilization of the spatial conformation of lipopolysaccharides (LPS) by depletion of Ca2+ and Mg2+ di-cations37 (ASW-Pi-Ca and NaCl + EDTA) or by lysozyme treatment38 (Fig. 6), suggesting no direct involvement of LPS in Pi accumulation. Retention of Pi by phosphorylation of extracellular macromolecules was ruled out, because alkaline phosphatase had no effect on the accumulated Pi (Fig. 6). Incubations with either acidic (pH 6) or alkaline (pH 10) ASW failed to remove the accumulated Pi (Fig. 6), demonstrating an insignificant effect of (de)protonation on the stability of the accumulated Pi.
To assess the role of PMF in retention of the accumulated Pi, we treated *Pi pre-labelled Synechococcus cells with the inhibitor carbonyl cyanide m-chloro-phenyl-hydrazone (CCCP) (Supplementary Table 7). Dissipation of the PMF had a minor effect on the already accumulated Pi: the amount of released Pi was comparable to the amount of Pi lost by washing live cells with ASW-Pi (Supplementary Fig. 4). In contrast to the above treatments, accumulated Pi was easily removed by short (1–2 min) rinses of live cells with a range of hypotonic solutions (Fig. 7), i.e., ASW diluted 1:10–1:100, phosphate-buffered saline or DW (Supplementary Table 8), which release periplasmic cell contents27,39 but do not cause osmotic cell lysis of marine Synechococcus (Fig. 5). Hypotonic washes reproducibly removed an amount of *Pi less than, or similar to, that removed from the PFA-fixed cellular macromolecules (Figs. 3 and 7). Thus, resistance to a variety of extracellular treatments (Fig. 6) in conjunction with the hypotonic removal of the accumulated Pi (Figs. 3, 5 and 7) strongly indicates the periplasmic location of the accumulated Pi.
The PMF is essential for Pi accumulation
The inhibitor CCCP has previously been used to investigate the role of the PMF in various aspects of cyanobacterial physiology26,27. Although we used this inhibitor to specifically elucidate the role of the PMF in Pi accumulation by cultured Synechococcus cells, we compared the CCCP effect with the effect of three other metabolic inhibitors (Supplementary Table 7) on Pi acquisition by oceanic bacterioplankton to safeguard from potential artefacts. A similar time course of Pi clearance by the control and 3-(3,4-dichloro-phenyl)-1,1-dimethylurea (DCMU)-treated bacterioplankton demonstrates that DCMU (which reduces photosynthetic electron flow and proton extrusion) slows down the microbial Pi clearance rate by ~30% (Supplementary Fig. 5). This reduction is consistent with the percentage (~30%) of cyanobacterial cells within the total bacterioplankton. The other three inhibitors completely terminated Pi clearance by bacterioplankton, as no time course was observed (Supplementary Fig. 5). The effect of N,N'-dicyclohexyl-carbodiimide (DCCD, an ATPase inhibitor) was delayed three times longer compared with 2,5-dibromo-3-methyl-6-isopropylbenzo-quinone (DBMIB) and CCCP, which halted Pi accumulation after 4 min (Fig. 8a, grey bars).
Likewise, CCCP was a highly effective inhibitor of Pi accumulation by cultured Synechococcus cells (Fig. 8a). The time delay of 3–5 min common to the three strains was similar to the 4 min delay of CCCP-treated bacterioplankton. This abrupt halt of Pi accumulation by CCCP-treated Synechococcus cells indicates that, as in experiments with bacterioplankton (Fig. 8a), membrane depolarization rather than ATP depletion is behind the inhibition, whereas the generic 4 min delay presumably reflects the time required for CCCP molecules to integrate into cellular membranes. As there was <1 min to incorporate *Pi into the PFA-fixed macromolecules and even less time to incorporate it into TCA-fixed nucleic acids (Fig. 1d, e), the bulk of the accumulated Pi in CCCP-treated cells (32% and 11% of the total Pi was in the PFA-fixed macromolecules of oceanic bacteria and isolates, respectively) remained labile (Fig. 8a). This comparison of live and fixed cells worked equally well for bacterioplankton and for cultures (Fig. 8a), confirming that in all bacteria tested Pi is first accumulated in the periplasm before being imported into the cytoplasm and assimilated.
The PMF consists of the transmembrane chemical H+ gradient (ΔpH) and the transmembrane electric gradient (ΔΨ) that can be sustained by other cations. The protonophore CCCP dissipates both the ΔpH and ΔΨ. To specifically link one of the components to Pi accumulation in the periplasm, we abolished ΔΨ in WH8102 using the three cation-specific ionophores: a K+-specific ionophore (Valinomycin), a monovalent cation-specific (e.g., Na+ and K+) ionophore (Monensin) and a divalent metal-specific (e.g., Mn2+, Ca2+ and Mg2+) ionophore (A23187, Supplementary Table 7), which facilitate the electroneutral transport of cations through a membrane down the chemical gradient. Compared with the control, the three ionophores reduced Pi accumulation into live cells by 0.4×, 1.5× and 2.6×, respectively (Fig. 8b). A significant fraction (20–37%) of the accumulated Pi remained in the periplasm. As relative to the protonophore CCCP, the three ionophores had a minor effect on Pi accumulation by Synechococcus cells (Fig. 8b), we conclude that ΔpH rather than ΔΨ is crucial for the periplasmic Pi accumulation. To validate conclusions about energy-dependent accumulation of Pi, drawn from the inhibitor-based experiments, we tested whether Pi accumulation by SAR11, Prochlorococcus and Synechococcus cells could be energy-stimulated. Conveniently, these bacteria use light energy in their metabolism.
Light stimulation of Pi accumulation
To test whether light can energize bacterial Pi accumulation, we compared Pi clearance by oceanic bacterioplankton cells incubated in the light vs. dark (Fig. 9). Indeed, light enhanced Pi accumulation by all cells studied (Supplementary Table 9). Compared with spectacular (≤10 times) but sporadic light stimulation of Pi clearance by Prochlorococcus cells (and to a lesser extent by Synechococcus cells), light stimulation of Pi clearance by SAR11 cells was more uniform, averaging 40% (Fig. 9). Counter-intuitively, light stimulation was distinctly more pronounced at lower rates of Pi clearance in moderately Pi-depleted waters bordering the North Atlantic subtropical gyre but absent at the maximal rates of Pi clearance in the Pi-depleted gyre (Fig. 9 and Supplementary Table 2).
Accumulation of Pi in the periplasm starts with Pi diffusion from seawater. The theoretical maximal rate of Pi diffusion into the periplasm of a Gram-negative cell depends on a Pi diffusion coefficient and the permeable surface area10,11. The Pi diffusion coefficeint is a seawater trait affecting all cells equally. Cells could, however, control their Pi-permeable surface area by varying the number of OM porins. Prochlorococcus and Synechococcus strains respond to Pi limitation by the increased expression of Pi-specific porin genes and the increased abundance of porin proteins14,40,41. However, the number of these porins in the OM cannot increase indefinitely. Indeed, the maximal Pi clearance rates of oceanic cyanobacterial and SAR11 cells (Fig. 2a) converge towards ~30× below the theoretical maximum. The apparent upper limit to the number of Pi-specific OM porins is probably a result of the competition for space between porins selective for different small solutes, acquisition of which is essential for a living cell.
In contrast to other solutes, Pi somehow accumulates in the periplasm. Synechococcus cells rapidly accumulate labile Pi, which is then steadily internalized and assimilated into macromolecules (Figs. 3 and 4). The semi-labile nature of the accumulated Pi (hypotonic removal, Figs. 1c, 3 and 5–7) in conjunction with its resistance to a variety of extracellular treatments (Fig. 6) points to its periplasmic location. Diffusion-driven Pi acquisition11 can deplete environmental Pi concentrations at a constant clearance rate down to 10−12 mol l−1 (Fig. 2a). This is feasible only if the periplasmic Pi concentration of Synechococcus cells is <10−12 mol l−1. Even if we assume that the affinity of the Synechococcus PstS subunit for Pi is sufficient to bind Pi at <10−12 mol l−1 concentrations (five orders of magnitude below the determined affinity20) then every Pi molecule that enters and stays in the Synechococcus periplasm needs to be instantly bound by a PstS subunit (Fig. 1a†). Sequestration of 6.7 × 106 molecules (Fig. 2b) of labile Pi in the periplasm would require an equal number of PstS subunits. The volume of a PstS subunit of 3.5 × 4 × 7 nm dimensions42 is ~5.1 × 10−23 l, whereas the periplasmic volume of Synechococcus is ~2 × 10−17 l. This is sufficient to accommodate merely 4 × 105 PstS subunits, or 17× less than the number required. However, even if 4 × 105 PstS subunits cram into the periplasm, there would be no room left for other known periplasmic constituents: substrate binding subunits of at least some of 41 other ABC-type transporters in WH810243, periplasmic enzymes, e.g., alkaline phosphatase6,44 and aminopeptidase45, and peptidoglycan gel matrix—the major structural component of periplasm46. As Prochlorococcus cells that have a smaller periplasm47 can have as many ABC-type transporters and periplasmic enzymes as WH810243 but accumulate more Pi in their buffer (1.2 × 107 molecules22), it makes it even less probable that all periplasmic Pi is bound by PstS subunits. Finally, up to 20× more Pi (e.g., at the 3 h time point in Fig. 3b, d, yellow) remains labile in living Synechococcus cells than is incorporated into bacterial macromolecules (which should include PFA-cross-linked PstS subunits, Fig. 1d). As the PstS-mediated Pi uptake model (Fig. 1a†) cannot explain the results of our experiments, an alternative explanation is required.
The evident capacity of Synechococcus to import the accumulated Pi into the cytoplasm (Figs. 3 and 4b) is inconsistent with passive sorption of Pi to the outer cell surface48,49, because passively adsorbed Pi cannot effectively diffuse into the periplasm (Fig. 1a†). Passive sorption of Pi by marine bacteria is furthermore questionable, because metabolically inactive but structurally intact bacterioplankton cells (e.g., CCCP-, DBMIB-, DCCD-incapacitated cells) should retain their capacity to adsorb Pi passively, but instead, they do not adsorb *Pi present in seawater for hours (Supplementary Fig. 5). In contrast, the rapid cessation or slowing down of Pi accumulation caused by the specific inhibitors (Fig. 8) and light stimulation of Pi accumulation (Fig. 9) indicates that Pi accumulation in the periplasm is a metabolically active process.
Light could enhance Pi accumulation by SAR11 cells through a proteorhodopsin-generated PMF50. Cyanobacteria could use photosynthetically generated PMF for the same purpose. The former energy generator consistently increased Pi accumulation by 40%, whereas the latter energy generator showed a higher (≤10×) potential to boost Pi accumulation by Prochlorococcus cells (Fig. 9). Although such considerable light stimulation is notable, the most unexpected result of the light vs. dark experiments was the lack of light stimulation at the higher rates of Pi accumulation (Fig. 9) in the Pi-depleted waters of the North Atlantic subtropical gyre22. Apparently, under severe Pi depletion, bacteria do not rely on light as the primary energy source for Pi acquisition but use their internal energy sources. Pi accumulation seems so important for bacteria that their maximal Pi clearance rates (~30× below the theoretical maximal rate of nutrient acquisition by diffusion) are probably constrained by the limited OM surface for incorporating Pi-specific porins rather than by the availability of cellular energy.
It seems inexplicable that bacteria can accumulate millions of Pi molecules (Figs. 2b and 3b, d, and Supplementary Fig. 2b) in their periplasm, while maintaining diffusion even at exceedingly low environmental concentrations. The high percentage of *Pi present in the periplasm (Figs. 4–7) and the gradual assimilation of accumulated Pi (Figs. 3–4) indicates that the rate of periplasmic Pi accumulation by far exceeds the rate of Pi import. The parallel import of 33Pi pulse and 32Pi chase (Fig. 3) means that the spatial distribution of Pi in the periplasm is not ordered. Bacteria do not synthesize polyphosphates in the periplasm but somehow make Pi molecules semi-labile: accessible for PstS subunits but prevented from diffusing back into seawater. On the one hand, periplasmic Pi needs to remain in ionic form rather than phosphorylate organic molecules, because PstS subunits specifically bind HPO42− and H2PO4− ions42. On the other hand, Pi cannot be maintained as free ions because a few free Pi ions would make the periplasmic concentration of Pi above the environmental concentration and hence reverse diffusion. The nearly instantaneous termination of Pi accumulation (Fig. 8a) by the inhibitors CCCP and DBMIB (which dissipate the PMF and thereby halt ATP generation), compared with the delayed reduction of Pi accumulation by DCCD (which primarily blocks ATP generation and thus destroys ATP-sustained membrane potential), strongly indicates that Pi accumulation is linked to cellular energetics through the PFM rather than being directly driven by ATP through PstSCAB-type transporters (Fig. 1a†). The mutually exclusive conditions (listed above) of PMF-dependent Pi accumulation in the periplasm cannot be explained by PstS-mediated Pi import (Fig. 1a†). Therefore, an alternative mechanism is needed.
An alternative mechanism of PMF-dependent Pi accumulation in the periplasm is a conjecture, because we know little about the organization and functioning of the periplasm46 in a living bacterial cell. A cell maintains the PMF by extruding protons (H+ ions) across the plasma membrane against the electrochemical gradient using the energy of respiration and photosynthesis50,51,52. Accumulation of H+ in the periplasm would make it acidic relative to both the cytoplasm (pH ~ 7.253) and seawater (pH 8.0–8.354) (Fig. 1a‡). However, it is unlikely that many free H+ ions would accumulate in the periplasm of marine bacteria, because their OM is permeable to the smallest H+ ions. Hence, H+ ions should diffuse out into the environment to be neutralized by >100× excess of OH− ions in alkaline (pH 8.0–8.3) seawater54, thereby dissipating the membrane potential. To prevent the dissipation of the H+-based gradient, H+/Na+ antiporters may exchange at least some of the periplasmic H+ ions for Na+ ions55. This substitution could preserve the electrical gradient and support ATP production through H+-ATP synthases, which can transport Na+56. Genes for both the H+/Na+ antiporter and the Na+-driven ATP synthase are present in genomes of Prochlorococcus56 and Synechococcus isolates (e.g., NP_87599956, WP_010315318, WP_025922676, WP_002805629 and WP_062436653).
The electric potential between the acidic positively charged periplasm and the alkaline seawater would facilitate diffusion, or more precisely mass transfer, of anions with higher negative charge more strongly than anions with lower negative charge. Thereby, mass transfer through anion-selective porins13 of HPO42− and PO43− anions (which comprise 29% and 0.01% of the total Pi in seawater at pH 8.0, respectively57,58) would be particularly favourable. Stronger cationic association of the main seawater anions (e.g., Cl−, HSO4−, HCO3− and NO3−) would also favour mass transfer of free Pi anions. Once in the H+-enriched periplasm, the HPO42− and PO43− anions would associate with one or two H+ and metal cations to reach pH-dependent speciation equilibrium57,58. Kinetic stability of neutral Pi molecules would explain why disruption of the membrane potential only has a minor immediate effect on the Pi already accumulated in the periplasm (Supplementary Fig. 4).
Neutralization of cations with Pi anions would reduce the PMF, but that could be restored via continuous extrusion of H+ and cations across the plasma membrane to maintain mass transfer of Pi anions through porins as long as free Pi anions can mass transfer from the environment (Fig. 1a‡). Mass transfer would stop when equilibrium is reached, i.e., all Pi anions remaining in seawater are too strongly associated with environmental cations and molecules of water. The strength of their association depends on the total ionic composition of environmental solutions58; in seawater, the association of Pi anions is apparently weak, because we observed bacterial accumulation of Pi anions to continue down to environmental concentrations <10−12 mol l−1 (Fig. 2) and even <10−15 mol l−1 (Supplementary Table 4).
Although the periplasm becomes saturated with Pi in 1 h (e.g., 4 × 106 Pi molecules in 2.1 × 10−17 l periplasm equals 0.3 mol l−1, Fig. 3b, d), a cell somehow averts precipitation of the Pi salts in the periplasm. To prevent formation of insoluble Pi salts, a cell would need to minimize concentrations of divalent cations (e.g., Ca2+ and Mg2+, which associate with Pi to form salts with low solubility) in the periplasm, while balancing the concentrations of monovalent cations (e.g., Na+ and K+) and Pi anions to form neutral soluble ion pairs in a way analogous to the common phosphate buffer. Although cation-associated, the H2PO4− and HPO42− ions remain accessible to PstS-mediated (the PstS subunits have specific affinity to these two anion forms42) import, because they remain soluble (Fig. 1a‡). Formation of neutral ion pairs would maintain mass transfer of HPO42− and PO43− into the periplasm, and prevent H+ from escaping into seawater. To simplify the explanations we give below, we suggest to term this periplasmic H+-driven phosphate-cation association—phosphatation.
Maintaining maximal rates of Pi acquisition irrespective of the availability of light energy (Fig. 9) suggests that periplasmic phosphatation is functionally important for marine bacteria as different as cyanobacteria and SAR11 alphaproteobacteria. We propose the following three physiological functions of periplasmic phosphatation: (i) The periplasmic association of Pi with chemiosmotic cations accumulates not only Pi, but also cations. Accumulation of the latter could be viewed as energy conservation, because H+/Na+ import through the plasma membrane can generate ATP59,60. (ii) The high concentration of Pi accumulated in the periplasm could also ensure that the PstS transport system is always saturated with Pi and operates near its maximal rate. (iii) Periplasmic phosphatation could have an osmotic function. The concentration of Pi in the periplasm can increase by 0.3 mol l−1 in just 1 h (e.g., 4 × 106 Pi molecules in 2.1 × 10−17 l periplasm, Fig. 3b, d) and reach >0.5 mol l−1 within 3 h. Accumulation of an additional 0.5 mol l−1 of Pi salt would increase periplasmic osmolarity by ~0.5 osmol l−1. Considering that there are other osmotically active molecules in the periplasm, and that the Pi salt concentration could raise further the periplasmic osmolarity, this would easily exceed the osmolarity of seawater, i.e., ~1 osmol l−1 57. To equilibrate, the resultant osmotic differential water molecules would enter the periplasm increasing its volume and the OM turgor (Fig. 1a‡), explaining how the OM could work as a load-bearing element61. Therefore, we predict that periplasmic phosphatation is essential for bacterial osmotic regulation—a hypothesis worth testing experimentally.
There is little doubt that the cationic attraction of Pi anions predates life. It was proposed that back in the earliest Archean, positively charged clay particles in oceanic cold alkaline seeps could be viewed as primordial membranes of protocells62. Their positive charge would similarly attract Pi anions from seawater and this initial enrichment of clay particles with Pi might have been inherited by the first living forms, who gradually engaged Pi in cellular energetics and later genetics. Thus, periplasmic phosphatation is inherently essential for bacterial cells, because it could conserve energy as well as store and attract the vital Pi.
Data were collected on five oceanographic cruises (Supplementary Table 2) on board the Royal Research Ships James Clark Ross (JCR), James Cook (JC), Discovery III (D) and Discovery IV (DY), and the Research Vessel Maria S. Merian (MSM) in the North Atlantic during Atlantic Meridional Transect cruises AMT17-D299, AMT20-JC053, AMT22-JC079 and AMT27-DY084 in September–October 2005, 2010, 2012 and 2017, respectively, and during the MSM03 cruise in September–October 2006. At predawn, midday and occasional evening stations seawater samples were collected from 20 m (a representative depth from the surface mixed layer unaffected by the ship’s movement and contamination) using a 20 × 20 l Niskin bottle rosette (Miami, FL, USA) mounted on the stainless steel frame of a conductivity-temperature-depth profiler. Samples were decanted into 10 l polyethylene carboys. Experiments commenced within 1–2 h after sample collection. The concentrations of bioavailable Pi were determined using isotope dilution, concentration series bioassays7,22 with carrier-free 32P-labelled or 33P-labelled orthophosphoric acid (Hartmann Analytic GmbH, Braunschweig, Germany). Bioassay experiments were performed in polypropylene, screw cap, 2 ml micro-centrifuge tubes (Starlab, Milton Keynes, UK). Flow-sorting experiments and experiments with inhibitors were performed in Pyrex glass bottles (Fisher Scientific, Loughborough, UK). To minimize artificial contamination (e.g., Pi, other nutrients, heavy metals), all plastic-ware, glassware and silicone tubing were soaked in 10% HCl and repeatedly rinsed with DW and sampled seawater.
Synechococcus strains and growth conditions
We chose Synechococcus sp. WH8102, because its Pi acquisition and utilization strategies are known1,41. This strain belongs to clade III that is generally considered to be adapted to low-nutrient conditions63,64 and abundant in Pi-depleted Atlantic waters65. Strain Synechococcus sp. WH8109 is a member of clade II, a clade that dominates tropical and subtropical regions of the world’s ocean66,67, tolerates intermediate nutrient-depleted conditions64,67 and lives in nutrient-richer upwelling waters65. It was reasonable to expect that these strains accumulate Pi extracellularly, because they grow better under Pi-depleted rather than under Pi-replete conditions30. To elucidate whether the Pi accumulation was specific to cyanobacterial ecotypes that had evolved to cope with Pi deprivation, we also included an opportunistic strain Synechococcus sp. WH7803 (clade V) that reaches a maximal growth rate when supplemented with ≥10−6 mol l−1 Pi49.
Cultures of Synechococcus sp. WH8102 and Synechococcus sp. WH7803 were axenic. An axenic culture of Synechococcus sp. WH8109 strain was established by flow sorting. Live bacterial cells in a mixed culture were stained with 0.1 μg ml−1 Hoescht 33342 (final concentration) and Synechococcus sp. WH8109 cells were flow cytometrically differentiated from other bacteria by their specific autofluorescence. Using a custom-built MoFlo XDP instrument68, WH8109 cells were flow-sorted in batches of 10–200 cells directly into sterile tubes with ASW medium. Tubes were placed into an illuminated growth chamber and incubated until growth became evident by colour.
Axenic cultures of Synechococcus sp. strains WH8102, WH8109 and WH7803 were cultivated in an ASW medium70 prepared using American Chemical Society grade reagents and supplemented with 8 × 10−5 mol l−1 K2HPO4. Semi-continuous cultures were maintained at 23 °C with gentle agitation at 100 r.p.m. on an orbital shaker at a light intensity of ~20 μmol photons m−2 s−1. Axenic cultures were monitored regularly for contamination by plating ~106 cells on ASW agar plates containing 0.5 g l−1 yeast extract and were contaminant free. To maintain trace Pi conditions, glassware used for Pi-free cultivation and experiments was soaked in 10% HCl and thoroughly rinsed with DW.
Cell enumeration and flow sorting
Cultured Synechococcus cells were counted live, unstained, to assess their cell integrity during laboratory experiments (e.g., see Fig. 5c). Oceanic samples were fixed with PFA (1% w/v final concentration), stained with SYBR Green I DNA dye69 at 20 °C in the dark for 1 h and flow cytometrically counted and sorted22,70 (Supplementary Fig. 1). Bacterioplankton cells flow-sorted during the D299 and JC053 cruises were taxonomically identified using fluorescence in situ hybridization7,9. High nucleic acid-containing bacteria with virtually undetectable chlorophyll autofluorescence71 were identified as Prochlorococcus. Low nucleic acid-containing bacteria were identified as SAR11 alphaproteobacteria. Oceanic Synechococcus were identified flow cytometrically based on their specific orange autofluorescence7.
Western blot analyses
Cells from an exponentially growing Synechococcus WH8102 culture (OD750 = 0.25) were starved for Pi to induce PstS synthesis. After 60 h incubation, 800 ml cells were pelleted by centrifugation, washed twice with 20 ml ASW and subjected to 1 min hypotonic shock in 6 ml DW. Following centrifugation, supernatant (6 ml) was collected. The remaining cell pellet was suspended in 6 ml cold DW, supplemented with Protease Inhibitor Cocktail II (Abcam) and the cells were disrupted by two passes through a high-pressure Cell Disruptor (Constant Systems Ltd, Northants, UK). Intact cells were removed from hypotonic DW supernatant and disrupted cells by short centrifugation (5000 × g, 10 min, 4 °C). Large membrane debris of disrupted cells were removed using high-speed centrifugation (20,000 × g, 60 min, 4 °C) and the remaining cell lysate (6 ml) was collected. Volumes (5, 10 or 20 μl) of DW supernatant and cell lysate were denatured with 1 : 3 (v/v) sample buffer at 90 °C for 5 min and separated by SDS-polyacrylamide gel electrophoresis using 14% acrylamide. A reference gel was stained with Coomassie blue. Western blot immunolabelling used antibodies against the periplasmic phosphate-binding subunit PstS72 (1 : 10,000 dilution) and the large Rubisco subunit RbcL (1 : 10,000; AS03 037, Agrisera). For western blotting, proteins were transferred to the polvinylidene difluoride (PVDF) membrane and analysed using standard protocols, following the use of a horseradish peroxidase (HRP)-conjugated goat anti-rabbit IgG HRP-linked secondary antibody (1 : 5000; #7074, Cell Signaling Technology®).
Exponentially growing Synechococcus sp. WH8102 cells (0.3–3 × 107 cells ml−1) were collected on a 0.2 μm pore size Nuclepore track-etched filter membrane (Whatman International, Ltd, Maidstone, UK) mounted in a filter holder (Swinnex, Millipore, Ireland) and washed with 40 ml Pi-free ASW (ASW-Pi) at a flow rate of 2 ml min−1 using a syringe pump (KD Scientific, Holliston, MA, USA). The washed cells were pulsed with 10−8 mol l−1 Pi traced with 33Pi (H333PO4, specific activity ~111 TBq mmol−1; Hartmann Analytic GmbH, Braunschweig, Germany) for 2 h. The pulse was chased with 100× Pi concentration (10−6 mol l−1) traced with carrier-free 32Pi (H332PO4, specific activity ~222 TBq mmol−1; Hartmann Analytic GmbH, Braunschweig, Germany) and incubated for up to 23 h. Retention of 33Pi and 32Pi was determined for live and PFA-fixed cells (Fig. 3). Dissolved tracer was washed from live cells with ASW-Pi using a 0.2 μm pore size PVDF membrane spin column (Proteus Mini, Generon, Maidenhead, UK): 0.5 ml cells were placed in a column, spun down at 5000 × g for 1 min and washed thrice with 0.1 ml of ASW-Pi. The radioactivity was determined separately for the cells retained in the spin column insert and for the effluent in the collection tube. To fix cells, 0.5–1.5 ml cells were fixed with 1% PFA, incubated at ambient temperature for 1 h and filtered on 0.2 μm pore size polycarbonate filters. Filters were rinsed twice with 4 ml DW, placed in 20 ml scintillation vials (Meridian, Epsom, UK), mixed with 10 ml scintillation cocktail (Goldstar, Meridian) and radioassayed using a low-activity liquid scintillation analyser (Tri-Carb 3180 TR/SL, PerkinElmer, Beaconsfield, UK) with QuantaSmart™ software. To assess their reproducibility, the pulse-chase experiments were repeated thrice. The results of two experiments are presented in Fig. 3.
Extracellular Pi labelling of Synechococcus cells
Cells washed of free Pi were re-suspended in 20 ml ASW-Pi, supplemented with 80–160 Bq ml−1 of H333PO4 or H332PO4 and incubated in a glass bottle under standard cultivation conditions for 1–3 h. The labelled cells were collected on 0.2 μm pore size Nuclepore filters and effluent was sampled to determine the efficiency of *Pi clearance (Supplementary Table 4). Extracellularly *Pi pre-labelled Synechococcus cells were washed free of residual *Pi with 20 ml ASW-Pi and re-suspended in solutions appropriate for further experiments.
Retention of the accumulated Pi
*Pi pre-labelled Synechococcus cells were treated with enzymes (Supplementary Table 6) or re-suspended in a range of solutions (Supplementary Tables 5 and 8) using either a 0.2 μm pore size PVDF membrane spin column or a 0.2 μm pore size Nuclepore membrane filter fitted in a 13 mm diameter Swinnex filter holder. For treatments in spin columns, 0.3–0.5 ml treated cells were incubated in a column, spun down at 5000 × g for 1 min and washed thrice with 0.1 ml of an appropriate washing solution. The radioactivity was determined separately for the cells retained in the spin column insert and for the effluent in the collection tube. Alternatively, cells were collected on a Nuclepore membrane, washed with 10 ml of a treatment solution at a rate of 0.5–1 ml min−1 and the radioactivity determined for the filter membrane only. The activity of the protease Proteinase K in ASW was confirmed using the PDQ protease assay (Athena Environmental Sciences, Inc., Baltimore, MD, USA). To assess their reproducibility, all experiments were repeated independently at least thrice.
Inhibitors were dissolved in dimethyl sulfoxide (DMSO) and added to samples to a final concentration of 0.05–1 × 10−4 mol l−1 (Supplementary Table 7) simultaneously with the 33Pi or 32Pi tracer. To provide a control, samples were supplemented with the appropriate amount of DMSO. Accumulation of *Pi in DMSO-amended samples and control samples without DMSO addition was similar (Shapiro–Wilk normality test passes P = 0.932, paired t-test t = −0.155 with 8 degrees of freedom, two-tailed P-value 0.881). Samples were incubated in the light under ambient conditions and sub-samples were withdrawn periodically to determine the total microbial Pi accumulation. To assess their reproducibility, all experiments were repeated independently at least thrice.
Microbeam synchrotron X-ray fluorescence analysis
These measurements were performed at the Diamond Light Source, the UK National Synchrotron Facility. Beamline I18 (Microfocus Spectroscopy) was used to measure the amount of P in cells of Prochlorococcus cyanobacteria (range 1–5 × 105 cells) and Synechococcus sp. strain WH8102 (0.1–1 × 106 cells) (Supplementary Table 3). Cells were flow-sorted onto 0.2 μm pore size polycarbonate filters (P free) using the custom-built MoFlo XDP instrument68. The filters with dried sorted cells were mounted on a custom-made sample holder and the cell-containing filter areas (~1000 × 2000 µm) were probed with a beam spot of 30 × 30 µm2 in a helium atmosphere (Supplementary Fig. 3a-c). An approximation of the number of photons on the sample was derived by measuring a thin-film reference material with known mass deposition of a number of metals. This calculated photon flux applied to the cells and accounting for differences in sample matrix, density and volume, leads to an estimated P mass per pixel for the cells. From these values, the total number of P atoms was derived for each cell preparation by integrating signal from all P containing pixels using PyMCA X-ray Fluorescence Toolkit (http://pymca.sourceforge.net) and ImageJ 1.50i (https://imagej.nih.gov/ij/). Using the cultured Synechococcus cells as calibrants, sensitivity (≥2 × 104 cells) and linear range (≤7.5 × 105) of the method were determined.
The determined P content was 4.14 ± 1.06 × 106 P atoms per cell for oceanic Prochlorococcus equivalent to 1.3–3.1 genomes (accessions: ASM1148v1, ASM1246v1, ASM1264v1, ASM1564v1, ASM1566v1, ASM1858v1, ASM75985v1, ASM792v1 and ASM1146v1). The extracellularly accumulated Pi of cells was removed during sorting by the sheath fluid of low osmotic concentration (~0.03 osmol l−1), which strips the extracellularly bound Pi (Figs. 1, 3, and 5–7). Therefore, only intracellular phosphorus was measured for the sorted cells. In comparison, the P content calculated for an exponentially growing extracellular Pi-free Synechococcus sp. WH8102 cell was 1.9 ± 0.16 × 107 P atoms, which is ~4 times the genomic P content (ASM19597v1).
Bacterioplankton cells were imaged and their dimensions measured68 (Supplementary Fig. 3d-f and Supplementary Table 1). Cells of Synechococcus sp. WH8102 were imaged using a LSM780 Confocal Laser Scanning Microscope (Carl Zeiss, Jena, Germany), using 488 nm (Ar) and 594 nm (HeNe) laser lines.
Statistics and reproducibility
Each experiment was repeated independently at least three times and often more than ten times.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
The auxiliary data collected on research cruises are archived indefinitely at the British Oceanographic Data Centre (BODC) and are available at www.bodc.ac.uk. The μ-SXRF data are available from Open Science Framework at http://osf.io/6RH2V. All other data supporting the findings of this study are available within the paper and its Supplementary Information files. The source data underlying Figs. 2–4 and 6–9, and Supplementary Figs. 2, 4 and 5 are provided as a Source Data file.
Moore, L. R., Ostrowski, M., Scanlan, D. J., Feren, K. & Sweetsir, T. Ecotypic variation in phosphorus acquisition mechanisms within marine picocyanobacteria. Aquat. Microb. Ecol. 39, 257–269 (2005).
Sohm, J. A. & Capone, D. G. Phosphorus dynamics of the tropical and subtropical north Atlantic: Trichodesmium spp. versus bulk plankton. Mar. Ecol. Prog. Ser. 317, 21–28 (2006).
Dyhrman, S. T. & Haley, S. T. Phosphorus scavenging in the unicellular marine diazotroph Crocosphaera watsonii. Appl. Environ. Microbiol. 72, 1452–1458 (2006).
Van Mooy, B. A., Rocap, G., Fredricks, H. F., Evans, C. T. & Devol, A. H. Sulfolipids dramatically decrease phosphorus demand by picocyanobacteria in oligotrophic marine environments. Proc. Natl Acad. Sci. USA 103, 8607–8612 (2006).
Mather, R. L. et al. Phosphorus cycling in the North and South Atlantic Ocean subtropical gyres. Nat. Geosci. 1, 439 (2008).
Sebastian, M. & Ammerman, J. W. The alkaline phosphatase PhoX is more widely distributed in marine bacteria than the classical PhoA. ISME J. 3, 563 (2009).
Zubkov, M. V. et al. Microbial control of phosphate in the nutrient-depleted North Atlantic subtropical gyre. Environ. Microbiol. 9, 2079–2089 (2007).
Michelou, V. K., Lomas, M. W. & Kirchman, D. L. Phosphate and adenosine-5′-triphosphate uptake by cyanobacteria and heterotrophic bacteria in the Sargasso Sea. Limnol. Oceanogr. 56, 323–332 (2011).
Gomez-Pereira, P. R. et al. Comparable light stimulation of organic nutrient uptake by SAR11 and Prochlorococcus in the North Atlantic subtropical gyre. ISME J. 7, 603–614 (2013).
Koch, A. L. in Advances in Microbial Ecology (ed. Marshall, K.C.) 37–70 (Springer, USA, 1990).
Jumars, P. A. et al. Physical constraints on marine osmotrophy in an optimal foraging context. Aquat. Microb. Ecol. 7, 121–159 (1993).
Nikaido, H. & Vaara, M. Molecular basis of bacterial outer membrane permeability. Microbiol. Rev. 49, 1–32 (1985).
Pongprayoon, P., Beckstein, O., Wee, C. L. & Sansom, M. S. P. Simulations of anion transport through OprP reveal the molecular basis for high affinity and selectivity for phosphate. Proc. Natl Acad. Sci. USA 106, 21614–21618 (2009).
Fuszard, M. A., Wright, P. C. & Biggs, C. A. Cellular acclimation strategies of a minimal picocyanobacterium to phosphate stress. FEMS Microbiol. Lett. 306, 127–134 (2010).
Rosenberg, H., Gerdes, R. & Chegwidden, K. Two systems for the uptake of phosphate in Escherichia coli. J. Bacteriol. 131, 505–511 (1977).
Willsky, G. R. & Malamy, M. H. Characterization of two genetically separable inorganic phosphate transport systems in Escherichia coli. J. Bacteriol. 144, 356–365 (1980).
Lebens, M., Lundquist, P., Söderlund, L., Todorovic, M. & Carlin, N. I. The nptA gene of Vibrio cholerae encodes a functional sodium-dependent phosphate cotransporter homologous to the type II cotransporters of eukaryotes. J. Bacteriol. 184, 4466–4474 (2002).
Sowell, S. M. et al. Transport functions dominate the SAR11 metaproteome at low-nutrient extremes in the Sargasso Sea. ISME J. 3, 93 (2008).
Scanlan, D. J. et al. Ecological genomics of marine picocyanobacteria. Microbiol. Mol. Biol. Rev. 73, 249–299 (2009).
Neznansky, A., Blus-Kadosh, I., Yerushalmi, G., Banin, E. & Opatowsky, Y. The Pseudomonas aeruginosa phosphate transport protein PstS plays a phosphate-independent role in biofilm formation. FASEB J. 28, 5223–5233 (2014).
McCarren, J. et al. Inactivation of swmA results in the loss of an outer cell layer in a swimming Synechococcus strain. J. Bacteriol. 187, 224–230 (2005).
Zubkov, M. V., Martin, A. P., Hartmann, M., Grob, C. & Scanlan, D. J. Dominant oceanic bacteria secure phosphate using a large extracellular buffer. Nat. Commun. 6, 7878 (2015).
Temperton, B., Gilbert, J. A., Quinn, J. P. & McGrath, J. W. Novel analysis of oceanic surface water metagenomes suggests importance of polyphosphate metabolism in oligotrophic environments. PLoS ONE 6, e16499 (2011).
Li, P. -N. et al. Nutrient transport suggests an evolutionary basis for charged archaeal surface layer proteins. ISME J. 12, 2389–2402 (2018).
Li, W. K. W. & Dickie, P. M. Metabolic inhibition of size-fractionated marine plankton radiolabeled with amino acids, glucose, bicarbonate, and phosphate in the light and dark. Microb. Ecol. 11, 11–24 (1985).
Spiller, H., Stallings, W. Jr, Tu, C. & Gunasekaran, M. Dependence of H+ exchange and oxygen evolution on K+ in the marine cyanobacterium Synechococcus sp. strain UTEX 2380. Can. J. Microbiol. 40, 257–265 (1994).
Ikeya, T., Ohki, K., Takahashi, M. & Fujita, Y. Study on phosphate uptake of the marine cyanophyte Synechococcus sp. NIBB 1071 in relation to oligotrophic environments in the open ocean. Mar. Biol. 129, 195–202 (1997).
Strahl, H. & Hamoen, L. W. Membrane potential is important for bacterial cell division. Proc. Natl Acad. Sci. USA 107, 12281–12286 (2010).
Kendall, A. I. Bacterial metabolism. Physiol. Rev. 3, 438–455 (1923).
Mazard, S., Wilson, W. H. & Scanlan, D. J. Dissecting the physiological response to phosphorus stress in marine Synechococcus isolates (Cyanophyceae). J. Phycol. 48, 94–105 (2012).
Zubkov, M. V. Faster growth of the major prokaryotic versus eukaryotic CO2 fixers in the oligotrophic ocean. Nat. Commun. 5, 3776 (2014).
Mary, I. et al. Light enhanced amino acid uptake by dominant bacterioplankton groups in surface waters of the Atlantic Ocean. FEMS Microbiol. Ecol. 63, 36–45 (2008).
Zubkov, M. V. & Sleigh, M. A. Ingestion and assimilation by marine protists fed on bacteria labeled with radioactive thymidine and leucine estimated without separating predator and prey. Microb. Ecol. 30, 157–170 (1995).
Frece, J. et al. Importance of S‐layer proteins in probiotic activity of Lactobacillus acidophilus M92. J. Appl. Microbiol. 98, 285–292 (2004).
Parsot, C., Ménard, R., Gounon, P. & Sansonetti, P. J. Enhanced secretion through the Shigella flexneri Mxi‐Spa translocon leads to assembly of extracellular proteins into macromolecular structures. Mol. Microbiol. 16, 291–300 (2006).
Brahamsha, B. An abundant cell-surface polypeptide is required for swimming by the nonflagellated marine cyanobacterium Synechococcus. Proc. Natl Acad. Sci. USA 93, 6504–6509 (1996).
Martin, N. L. & Beveridge, T. J. Gentamicin interaction with Pseudomonas aeruginosa cell envelope. Antimicrob. Agents Chemother. 29, 1079–1087 (1986).
Masschalck, B. & Michiels, C. W. Antimicrobial properties of lysozyme in relation to foodborne vegetative bacteria. Crit. Rev. Microbiol. 29, 191–214 (2003).
Heppel, L. A. Selective release of enzymes from bacteria. Science 156, 1451–1455 (1967).
Martiny, A. C., Coleman, M. L. & Chisholm, S. W. Phosphate acquisition genes in Prochlorococcus ecotypes: evidence for genome-wide adaptation. Proc. Natl Acad. Sci. USA 103, 12552–12557 (2006).
Tetu, S. G. et al. Microarray analysis of phosphate regulation in the marine cyanobacterium Synechococcus sp. WH8102. ISME J. 3, 835–849 (2009).
Luecke, H. & Quiocho, F. A. High specificity of a phosphate transport protein determined by hydrogen bonds. Nature 347, 402–406 (1990).
Scanlan, D. J. & West, N. J. Molecular ecology of the marine cyanobacterial genera Prochlorococcus and Synechococcus. FEMS Microbiol. Ecol. 40, 1–12 (2002).
Palenik, B. et al. The genome of a motile marine Synechococcus. Nature 424, 1037–1042 (2003).
Martinez, J. & Azam, F. Aminopeptidase activity in marine Chroococcoid cyanobacteria. Appl. Environ. Microbiol. 59, 3701–3707 (1993).
Hobot, J. A., Carlemalm, E., Villiger, W. & Kellenberger, E. Periplasmic gel: new concept resulting from the reinvestigation of bacterial cell envelope ultrastructure by new methods. J. Bacteriol. 160, 143–152 (1984).
Ting, C. S., Hsieh, C., Sundararaman, S., Mannella, C. & Marko, M. Cryo-electron tomography reveals the comparative three-dimensional architecture of Prochlorococcus, a globally important marine cyanobacterium. J. Bacteriol. 189, 4485–4493 (2007).
Sanudo-Wilhelmy, S. A. et al. The impact of surface-adsorbed phosphorus on phytoplankton Redfield stoichiometry. Nature 432, 897–901 (2004).
Fu, F.-X., Zhang, Y., Feng, Y. & Hutchins, D. A. Phosphate and ATP uptake and growth kinetics in axenic cultures of the cyanobacterium Synechococcus CCMP 1334. Eur. J. Phycol. 41, 15–28 (2006).
Walter, J. M., Greenfield, D., Bustamante, C. & Liphardt, J. Light-powering Escherichia coli with proteorhodopsin. Proc. Natl Acad. Sci. USA 104, 2408–2412 (2007).
Scholes, P., Mitchell, P. & Moyle, J. The polarity of proton translocation in some photosynthetic microorganisms. Eur. J. Biochem. 8, 450–454 (1969).
Nitschmann, W. H. & Peschek, G. A. Oxidative phosphorylation and energy buffering in cyanobacteria. J. Bacteriol. 168, 1205–1211 (1986).
Belkin, S. & Packer, L. Determination of pH gradients in intact cyanobacteria by electron spin resonance spectroscopy. Methods Enzymol. 167, 670–677 (1988).
Marion, G. M. et al. pH of seawater. Mar. Chem. 126, 89–96 (2011).
Padan, E. & Schuldiner, S. Molecular physiology of Na+/H+ antiporters, key transporters in circulation of Na+ and H+ in cells. Biochim. Biophys. Acta Bioenerg. 1185, 129–151 (1994).
Dufresne, A. et al. Genome sequence of the cyanobacterium Prochlorococcus marinus SS120, a nearly minimal oxyphototrophic genome. Proc. Natl Acad. Sci. USA 100, 10020–10025 (2003).
Atlas, E., Culberson, C. & Pytkowicz, R. M. Phosphate association with Na+, Ca2+ and Mg2+ in seawater. Mar. Chem. 4, 243–254 (1976).
Dickson, A. G. & Riley, J. P. The estimation of acid dissociation constants in sea-water media from potentiometric titrations with strong base. II. The dissociation of phosphoric acid. Mar. Chem. 7, 101–109 (1979).
Skulachev, V. P. The sodium cycle: a novel type of bacterial energetics. J. Bioenerg. Biomembr. 21, 635–647 (1989).
Müller, V. & Grüber, G. ATP synthases: structure, function and evolution of unique energy converters. Cell Mol. Life Sci. 60, 474–494 (2003).
Rojas, E. R. et al. The outer membrane is an essential load-bearing element in Gram-negative bacteria. Nature 559, 617 (2018).
Martin, W. & Russell, M. J. On the origins of cells: a hypothesis for the evolutionary transitions from abiotic geochemistry to chemoautotrophic prokaryotes, and from prokaryotes to nucleated cells. Philos. Trans. R. Soc. B 358, 59–85 (2003).
Zwirglmaier, K. et al. Basin-scale distribution patterns of picocyanobacterial lineages in the Atlantic Ocean. Environ. Microbiol. 9, 1278–1290 (2007).
Post A. F. in Harmful Cyanobacteria (eds Huisman, J., Matthijs, H. C. P. & Visser, P. M.) Ch. 5 (Springer, 2005).
Mazard, S., Ostrowski, M., Partensky, F. & Scanlan, D. J. Multi-locus sequence analysis, taxonomic resolution and biogeography of marine Synechococcus. Environ. Microbiol. 14, 372–386 (2012).
Farrant, G. K. et al. Delineating ecologically significant taxonomic units from global patterns of marine picocyanobacteria. Proc. Natl Acad. Sci. USA 113, E3365–E3374 (2016).
Sohm, J. A. et al. Co-occurring Synechococcus ecotypes occupy four major oceanic regimes defined by temperature, macronutrients and iron. ISME J. 10, 333–345 (2016).
Kamennaya, N. A., Kennaway, G., Fuchs, B. M. & Zubkov, M. V. “Pomacytosis”—Semi-extracellular phagocytosis of cyanobacteria by the smallest marine algae. PLoS Biol. 16, e2003502 (2018).
Zubkov, M. V., Burkill, P. H. & Topping, J. N. Flow cytometric enumeration of DNA-stained oceanic planktonic protists. J. Plankton Res. 29, 79–86 (2007).
Marie, D., Partensky, F., Jacquet, S. & Vaulot, D. Enumeration and cell cycle analysis of natural populations of marine picoplankton by flow cytometry using the nucleic acid stain SYBR Green I. Appl. Environ. Microbiol. 63, 186–193 (1997).
Hartmann, M. et al. Efficient CO2 fixation by surface Prochlorococcus in the Atlantic Ocean. ISME J. 8, 2280 (2014).
Scanlan, D. J., Mann, N. H. & Carr, N. G. The response of the picoplanktonic marine cyanobacterium Synechococcus species WH7803 to phosphate starvation involves a protein homologous to the periplasmic phosphate‐binding protein of Escherichia coli. Mol. Microbiol. 10, 181–191 (1993).
We thank the captains, officers, crew and chief scientists of the research cruises JCR091, MSM03, JC053, JC079 and DY084. We are grateful to M. A. Sleigh and D. Green for their supportive criticism of earlier versions of the manuscript, to J. Wulf, I. Mary, C. Grob and M. Hartmann for help with experiments at sea, and to H. Zer for help with the biochemical analysis. This study was supported by the UK Natural Environment Research Council through Research grants NE/M014363/1 and NE/J02273X/1. We acknowledge the Diamond Light Source for beam time under proposal NT15802-1.
The authors declare no competing interests.
Peer review information Nature Communications thanks Adam Martiny, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Kamennaya, N.A., Geraki, K., Scanlan, D.J. et al. Accumulation of ambient phosphate into the periplasm of marine bacteria is proton motive force dependent. Nat Commun 11, 2642 (2020). https://doi.org/10.1038/s41467-020-16428-w
Nature Geoscience (2021)
Nanomolar phosphate supply and its recycling drive net community production in the subtropical North Pacific
Nature Communications (2021)