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Applied and Environmental Microbiology, January 2008, p. 23-31, Vol. 74, No. 1
0099-2240/08/$08.00+0     doi:10.1128/AEM.01007-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Ni Uptake and Limitation in Marine Synechococcus Strains{triangledown}

Christopher L. Dupont,1 Katherine Barbeau,2 and Brian Palenik3*

Scripps Institution of Oceanography, University of California, San Diego, La Jolla, California 92093,1 Geosciences Research Division, Scripps Institution of Oceanography, University of California, San Diego, La Jolla, California 92093,2 Marine Biology Research Division, Scripps Institution of Oceanography, University of California, San Diego, La Jolla, California 920933

Received 4 May 2007/ Accepted 11 October 2007


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ABSTRACT
 
Ni accumulation and utilization were studied in two strains of marine Synechococcus, isolated from both coastal (CC9311; clade I) and open-ocean (WH8102; clade III) environments, for which complete genome sequences are available. Both strains have genes encoding an Ni-containing urease and when grown on urea without Ni become Ni-N colimited. The Ni requirements of these strains also depend upon the genomic complement of genes encoding superoxide dismutase (SOD). WH8102, with a gene encoding only an Ni-SOD, has a novel obligate requirement for Ni, regardless of the N source. Reduced SOD activity in Ni-depleted cultures of WH8102 supports the link of this strain's Ni requirement to Ni-SOD. The genome of CC9311 contains a gene for a Cu/Zn-SOD in addition to a predicted pair of Ni-SODs, yet this strain cannot grow without Ni on NO3– and can grow only slowly on NH4+ without Ni, implying that the Cu/Zn-SOD cannot completely replace Ni-SOD in marine cyanobacteria. CC9311 does have a greater tolerance for Ni starvation. Both strains increase their Ni uptake capabilities and actively bioconcentrate Ni in response to decreasing extracellular and intracellular Ni. The changes in Ni uptake rates were more pronounced in WH8102 than in CC9311 and for growth on urea or nitrate than for growth on ammonia. These results, combined with an analysis of fully sequenced marine cyanobacterial genomes, suggest that the growth of many marine Synechococcus and all Prochlorococcus strains is dependent upon Ni.


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INTRODUCTION
 
In surface seawater, nickel is generally present in low nanomolar amounts (3 to 4 nM) (6) with elevated concentrations in coastal waters due to natural terrestrial and anthropogenic inputs (7, 12). Ni concentrations in the ocean exhibit a "nutrient-like" depth profile, being depleted in the euphotic zone and increasing with depth (7), indicating that biological uptake and remineralization processes control Ni geochemistry. While no marine phytoplankton have been shown to have an obligate requirement for Ni, the role of Ni in urea assimilation is well established and historically has been used to explain the "nutrient-like" depth profile of Ni in seawater (25). Urease, an amidohydrolase with Ni in the active site (18), catalyzes the dissociation of urea into ammonia and carbon dioxide. For numerous algal taxa, it has been shown that growth with urea as the sole nitrogen source is nickel dependent (11, 29, 31), with nickel-deprived cultures becoming Ni-N colimited (37). These Ni-N-colimited phytoplankton can subsequently grow with the addition of Ni, which presumably results in a functional urease, or with the addition of ammonia, which does not require Ni for assimilation (37). The presence and regulation of urease in Synechococcus has been characterized (8), though the Ni dependency of growth on urea has not been shown. Most marine cyanobacterial genomes sequenced to date contain an operon encoding the multidomain Ni-containing urease (Fig. 1). The sole other previously characterized use for Ni among marine phytoplankton is in Ni/Fe hydrogenases found in some nitrogen-fixing cyanobacteria, though the Ni starvation-induced deficiency of this enzyme may not affect growth or nitrogen fixation (9).


Figure 1
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  		FIG. 1. Phylogenomic mapping of genes coding for SODs and urease within the fully sequenced genomes of marine cyanobacteria. The phylogenetic tree was drawn using the full-length amino acid sequences of rpoC2. The numbers at branch points denote the bootstrap support for the topology (100 replicates), with numbers on the left and right corresponding to protein parsimony and protein distance analyses, respectively.

It has long been recognized that phytoplankton metal physiology varies between closely related species or strains from different oceanic regimes. For example, centric diatoms from open-ocean environments tend to have lower Fe requirements than coastal species (5, 42), though increased copper requirements are a consequence (34, 35). The discovery of plastocyanin in Thalassiosira oceanica provided a mechanistic explanation for the increased Cu requirements in oceanic diatoms (34). By providing a complementary approach to physiological experiments, genomics may facilitate the elucidation of the molecular and genetic reasons for ecotypic differences in metal physiology. Of particular relevance, a comparison of the genomes of coastal (CC9311) and oceanic (WH8102) strains of marine Synechococcus from phylogenetically distinct clades recently revealed a number of proteomic differences in the number and type of metalloenzymes, implying differences in metal physiology (32).

Available genome sequences of marine cyanobacteria suggest another physiological use of Ni in addition to urease. Many marine cyanobacterial genomes sequenced to date contain a gene (sodN) potentially encoding an Ni-containing superoxide dismutase (SOD) (Fig. 1). SODs, of which there are Fe-, Mn-, Ni-, and Cu/Zn-containing isoforms, catalyze the breakdown of superoxide into hydrogen peroxide and molecular oxygen (16). As both photosynthesis and respiration generate toxic superoxide radicals, SODs play a critical role in protecting photosynthetic organisms from self- and environmentally induced oxidative stress (49). Previously only observed in the soil actinomycete Streptomyces (20), Ni-SOD is the only predicted isoform of SOD in Synechococcus sp. strain WH8102 (30) and in all strains of Procholorococcus sequenced to date (Fig. 1). One strain of Synechococcus, CC9311, also has a second gene encoding a protein with low sequence similarity (35%) to the biochemically characterized Streptomyces Ni-SOD (Fig. 1), with conservation of the Ni-binding residues (metal-binding residues [3]). The genomes of several strains of Synechococcus, including CC9311, also contain a gene (sodC) encoding a Cu/Zn SOD (Fig. 1), an isoform of SOD previously unobserved in cyanobacteria (32). A few other marine Synechococcus genomes contain a gene (sodB) encoding an Fe-SOD in lieu of sodN (Fig. 1), implying an evolutionary exchange of these two isoforms. Euryhaline cyanobacteria contain either an Fe-SOD (RCC307) (Fig. 1) or both an Fe and an Mn-SOD (PCC7002) (44). The genomes of the nitrogen-fixing cyanobacteria Trichodesmium and Crocosphaera contain genes encoding Mn (sodA) and Ni isoforms of SOD (Fig. 1), as well as an Ni/Fe-hydrogenase (not shown).

Here, we hypothesize that some marine cyanobacteria (e.g., WH8102) have an obligate growth requirement for Ni due to the obligate use of Ni-SOD, one independent of the nitrogen source used for growth. However, other strains (e.g., CC9311) should not have this obligate requirement due to the ability to utilize a different isoform of SOD. The requirements of these organisms also should depend upon the nitrogen source provided; growth on urea should impose a greater need for nickel than growth on NH4+. Finally, the increased, and in the case of WH8102 likely constitutive, need for Ni necessitates regulated Ni uptake and homeostasis. To test these hypotheses, we assayed the growth rates and cellular Ni concentrations of cultures of CC9311 and WH8102 over a range of free Ni2+ concentrations. SOD enzyme activities were also determined for both Ni-replete and -depleted cultures. Finally, the feedback regulation of Ni transport capabilities by extracellular and intracellular Ni was determined using radiotracer-based uptake experiments.


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MATERIALS AND METHODS
 
Bioinformatic analyses.
Full genome sequences were searched using Blastp (1) and the sodN, ureABCDE, and rpoC2 amino acid sequences of WH8102 (30). The resulting amino acid sequences were aligned using ClustalX (45) with manual intervention. The alignment of the sodN sequences was used to screen the putative sodN genes for the "Ni hook" motif, which is proposed to be characteristic of Ni-SOD (3). The aligned full-length rpoC2 sequences were bootstrapped and analyzed with Phylip using both maximum parsimony and protein distance algorithms (13). To generate the maximum parsimony trees, SEQBOOT, PROTPARS, and CONSENSE were used sequentially, while SEQBOOT, PROTDIST, NEIGHBOR, and CONSENSE were used for protein distance analysis.

Media and culture manipulations.
Synechococcus sp. strains WH8102 and CC9311 were grown in synthetic ocean water (SOW) minimally modified from the original recipe of Waterbury and Willey (46) (Table 1). Several steps were taken in the medium preparation and culturing process to prevent metal contamination. The media and added macronutrients were made trace metal free using column-based solid-phase extraction with precleaned Chelex-100 resin (pretreated according to the method in reference 36) and microwave sterilized prior to the addition of filter-sterilized trace metals, vitamins, and EDTA (Sigma ultra grade). All growth containers were rigorously cleaned as follows: (i) soaking in 1% citronox soap for at least 1 day, followed by multiple rinses with 18.2-m{Omega} water (milli-Q; Millipore); (ii) soaking in 10% trace metal grade HCl (Fisher) for 2 days; and (iii) multiple rinses with milli-Q water. Containers were sterilized by microwaving them while they were filled halfway with milli-Q water. Microwaving times varied according to volume, but a water temperature of 95°C was attained, as determined by an infrared thermometer (Fisher). All additions were made with sterile pipette tips rinsed twice with 0.2-µm-filtered HCl (10%) and milli-Q water, and all manipulations were conducted in HEPA-filtered laminar-flow benches.


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  		TABLE 1. Composition of SOW

MINEQL (48) was used to determine the chemical speciation of trace metals, and specifically Ni, in SOW (Fig. 2). Due to the constant presence of excess (13.45 µM) EDTA, the [Ni2+] is over 1,000 times lower than the total Ni concentration (Fig. 2) and remains stable despite uptake by phytoplankton. For media containing 13.45 µM EDTA and 50 nM total Ni, the [Ni2+] will only change from pNi (pNI = –log10[Ni2+]) 10.3 to pNi 10.6, with a pH change from 8.0 to 8.6. In order to vary the [Ni2+], only the total Ni added was varied; this kept all other micronutrient/EDTA ratios constant and also avoided potentially confounding effects that different concentrations of EDTA might have on cyanobacteria.


Figure 2
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  		FIG. 2. Ni speciation in SOW. The line shows the relationship between total and free nickel in SOW (the recipe is described in Table 1). Most of the Ni is bound by EDTA, with a small percentage bound by citrate and only minute amounts being in the bioavailable free ion (Ni2+ is shown on the y axis).

Cultures were grown at 19°C (CC9311) or 24°C (WH8102) under continuous light (unless otherwise noted, 80 µM photons m–2 s–1). For the growth assays and Ni quota experiments, 25-ml cultures were grown in capped 60-ml glass tubes. Cultures for protein assays and short-term uptake experiments were grown in 1-liter or 2.7-liter polycarbonate bottles with internal stir bars (100 rpm). The axenicities of the cultures were verified using agar plates supplemented with bactone and tryptone.

Determination of the specific growth rates and cellular Ni over a range of [Ni2+].
Cultures (25 ml) were grown in semicontinuous fashion at fixed free-ion concentrations of Ni. If transferred during mid-log phase of exponential growth, exponential growth is perpetuated, and due to the buffering by excess EDTA, [Ni2+] remain relatively constant and the cultures mimic trace metal "chemostats" (43). To further reduce batch effects, macronutrients were added in excess (Table 1). After growth equilibration (four or five transfers of 1:200 dilutions during mid-log-phase growth; ~4 weeks), the cells were transferred (1:200 dilution) to media containing 63Ni instead of "cold" Ni (63Ni from Perkin-Elmer diluted in Optima grade HCl, 0.1% in milli-Q). Following 10 to 12 doublings in the radioactive media, 10 ml of culture was gently filtered (Supor; 0.2 µm; 25 mm), and the filter was rinsed sequentially with 5 ml of 8-hydroxyquinoline-5-sulfonic acid (sulfoxime, an Ni chelator; 1 mM in SOW, pH 8.0; Avocado Biochemicals) and SOW to remove surface-bound 63Ni (37) and placed in 15 ml of scintillation fluid (Ecolyte). Aliquots (150 µl) of the culture were also spiked into separate scintillation vials to determine the total 63Ni (and therefore the total Ni and [Ni2+]) in each culture. The radioactivities of these samples were determined using standard scintillation counting with quench correction. Counts per minute were converted to dpm and compared to a dilution series (e.g., a standard curve) of the 63Ni stock solution to convert them to molar concentrations. Finally, glutaraldehyde (25%; Sigma) was added to 1-ml aliquots of the cultures to a final concentration of 0.25%. These samples were allowed to fix for 10 min and were frozen at –70°C until they were used for cell counts.

Culture phycoerythrin fluorescence was monitored as a proxy for culture biomass (Turner AU-10; excitation = 544 nM; emission = 577 nM). The specific growth rates of these cultures were determined using geometric mean linear regression of a plot of ln (culture fluorescence) versus time during mid-exponential phase. While the amount of phycoerythrin fluorescence per cell varied according to the Ni concentration in the media (not shown), the use of equilibrated semicontinuous cultures allowed reliable growth rate measurements once the fluorescence-per-cell relationship stabilized (23).

To determine the cell densities of the cultures, the glutaraldehyde-fixed (0.25%; stored at –70°C) samples were thawed at room temperature. Following 20-fold dilution with SOW, samples were filtered onto 0.2-µm black polycarbonate filters (Millipore) using 11-µm Nitex mesh filters (Millipore) as a support in glass chimney filter funnels (Fisher). The filters were mounted with immersion oil on glass slides, and cell counts were conducted on an epifluorescence microscope. Fields were counted in triplicate for each slide, and if the counts for a given slide deviated by more than 10%, a new filter and slide were made.

Ni uptake kinetics.
Growth-equilibrated semicontinuous cultures of WH8102 were harvested during mid-log phase via centrifugation (9,000 x g; 10 min) in acid-washed centrifuge tubes. The cells were washed twice by resuspension in Chelex-100 resin-purified (Chelexed) and microwave-sterilized SOW with no nutrients, EDTA, or trace metals (SOW–) and centrifugation. The resulting cell pellet was resuspended with Chelexed SOW– and subdivided into acid-washed polycarbonate bottles (60 ml). Half of these bottles were treated with 0.25% glutaraldehyde for 30 min (kill controls), at which point the live-kill pairs received staggered additions of 63Ni. For the time course study, 10 nM 63Ni was added, and for the kinetics study, additions ranged from 100 pM to 100 nM. For the kinetics study, cell suspensions were filtered after 30 min. Cells were collected on 0.2-µm filters (25 mm; polyethersulfone; Supor; Pall Corporation, East Hills, NY) by gentle vacuuming and rinsed sequentially with 5 ml of sulfoxime (1 mM in SOW–) and Chelexed SOW. Samples were also taken to determine the total 63Ni and cell densities in the suspensions; these were handled as described above. Uptake is presented as live-kill (kills are typically 20 to 30% of the live numbers).

Measurements of SOD activity.
Cultures (1 to 2 liters) of Synechococcus were grown under Ni-replete (50 nM Ni total) and Ni-depleted (no added nickel) conditions. Ni-replete cultures were harvested during mid-log phase, while Ni-depleted cultures were harvested in late-log phase prior to the Ni-dependent cessation of exponential growth (initial decline in exponential growth). Cells were harvested by centrifugation (9,000 rpm for 10 min in sterile acid-washed high-density polyethylene bottles), immediately frozen at –70°C, and stored until extraction. For protein extraction, the cell pellet was resuspended in 2 ml of sterile, cold buffer (0.1 M potassium phosphate, 0.1 mM EDTA, 0.1% Triton 100, 1 mM phenylmethylsulfonyl fluoride, pH 7.8) and passed twice through a French pressure cell (20,000 lb/in2). Unbroken cells were removed from the resulting suspension by centrifugation (5,000 x g for 5 min, twice). SOD activity was calculated using the ferricytochrome c reduction assay as described by Flohe and Otting (15). SOD activity was normalized to total protein concentrations determined using the bicinchoninic acid method (MicroBCA; Pierce).


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RESULTS
 
Essentiality of Ni to marine Synechococcus.
To test if Synechococcus CC9311 and WH8102 require Ni, cultures were grown in SOW with and without Ni and with either ammonium, urea, or nitrate as the added nitrogen source. As predicted, WH8102 could not grow without Ni, though the nature of the resulting limitation was dependent upon the nitrogen source in the media. When grown on urea, WH8102 became Ni-N colimited after one transfer to Ni-free medium, with either Ni or NH4+ additions restoring normal growth (Fig. 3A). After two or three transfers in Ni-free media, WH8102 was also unable to grow on NH4+ (Fig. 3B) or NO3– (Fig. 3B) without added Ni, and Ni additions restored growth (Fig. 3B and C). CC9311 also became Ni-N colimited when grown on urea without Ni (Fig. 3D), yet contrary to the original hypothesis, CC9311 was also not able to grow on NO3– without Ni (Fig. 3F) and could grow on NH4+ only at reduced rates compared to cultures provided with Ni (Fig. 3E).


Figure 3
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  		FIG. 3. Ni limitation of marine Synechococcus. Representative growth curves are shown for WH8102 grown on urea without Ni and subsequently amended with Ni or NH4+ (the triangles indicate addition to a limited culture) (A), WH8102 grown on NH4+ without Ni and amended with Ni (B), WH8102 grown on NO3– without Ni and amended with Ni (C), CC9311 grown as in panel A (D), CC9311 grown on NH4+ with and without Ni (E), and CC9311 grown as in panel C (F).

Ni-limited cultures of WH8102 consistently exhibited greatly reduced SOD activity compared to Ni-replete cultures (Table 2), an intuitive consequence of Ni starvation for WH8102, which contains a gene only for the Ni-containing isoform of SOD. In contrast, Ni-limited cultures of CC9311 contained SOD activity equal to or greater than that of Ni-replete cultures, except for growth on urea (Table 2). Ni additions restored maximal growth to Ni-limited CC9311 grown on NO3– or NH4+; therefore, either the Ni limitation observed in CC9311 is attributable to an unknown Ni metalloenzyme or the SOD activities observed in Ni-replete and Ni-limited cultures of CC9311 are not functionally equivalent.


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  		TABLE 2. SOD activities in Synechococcus

To address the hypothesis that, when Ni starved, CC9311 is more susceptible to oxidative stress despite measurable SOD activity, cultures were grown with and without Ni and with NH4+ as a nitrogen source at several light intensities from 0.5 to 150 microeinsteins m–2 s–1. At 0.5 and 10 microeinsteins m–2 s–1, the Ni-depleted cultures grew only marginally more slowly, or even faster, than the Ni-replete cultures (Fig. 4), but at 50 and 80 microeinsteins m–2 s–1, the Ni-depleted cultures grew significantly more slowly (Fig. 4). Even higher light levels (150 microeinsteins m–2 s–1) inhibited growth in Ni-replete cultures but were lethal to Ni-depleted cultures (Fig. 4). The decreased ability of CC9311 to tolerate high light when Ni starved leads us to believe that the SOD activity observed in Ni-depleted CC9311 extracts is not functionally equivalent to that observed in Ni-replete CC9311.


Figure 4
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  		FIG. 4. Growth rates of Synechococcus sp. strain CC9311 over a range of irradiance intensities when supplied with NH4+ as a nitrogen source and either no Ni or 50 nM total Ni. The error bars are for duplicate cultures.

Growth rates of WH8102 and CC9311 over a range of [Ni2+].
While the above-mentioned studies are diagnostic for the absolute Ni requirements of Synechococcus, their environmental relevance is unclear given the observed nanomolar concentrations of Ni in seawater. However, most trace metals in seawater are bound by organic ligands that reduce the concentrations of the bioavailable free and inorganic species of a metal (26). With this in mind, the specific growth rates (µ) were determined for Synechococcus supplied with a range of [Ni2+] and either urea, nitrate, or ammonia as a nitrogen source. The total Ni concentrations for each experiment are summarized in Table 3, while the relationship between total Ni and free Ni concentrations in the SOW media is shown in Fig. 2. The experiments tested a range of Ni concentrations that roughly reflected the potential environmental range of [Ni2+].


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  		TABLE 3. Maximum µ for each strain and nitrogen source and total concentration of Ni added for the experiments presenteda

Normalizing the specific growth rates using the highest observed growth rates for each strain and nitrogen source (Table 3), the growth efficiency (percent of maximal µ) over a range of [Ni2+] was calculated (Fig. 5). CC9311 was less affected by low [Ni2+] than WH8102 for growth on urea and NO3– (Fig. 5A and C). Nitrogen source-specific differences in Ni requirements for growth were also observed. For both strains, growth efficiency declined most severely with decreasing [Ni2+] when urea was supplied as the nitrogen source, as was originally hypothesized (Fig. 5A). The growth efficiencies of both strains were less affected by low [Ni2+] when they were grown on NH4+ than when they were grown on urea (Fig. 5A and B). When growing on nitrate, the growth efficiencies of the strains were affected by decreasing [Ni2+] in different fashions (Fig. 5C). WH8102 growth rates declined with decreasing Ni to a lesser extent than urea-grown cells (Fig. 5A and C). In contrast, CC9311 supplied with NO3– grew at near-maximal rates over a broad range of [Ni2+] (Fig. 5C).


Figure 5
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  		FIG. 5. Growth efficiencies over a range of [Ni2+] for Synechococcus sp. strains WH8102 and CC9311 grown on urea (A), NH4+ (B), and NO3– (C). Growth efficiency was calculated using the measured growth rates at a given [Ni2+] and the maximum growth rate observed for that strain and nitrogen source (Table 3). The error bars are the ranges for duplicate cultures.

While not a focus of this study, reduced growth efficiency at higher Ni concentrations, presumably due to Ni toxicity, was also observed (Fig. 5). The decline in growth rates was most severe for WH8102 growing on urea and nitrate, while CC9311 exhibited reduced growth rates at high [Ni2+] when grown on NH4+. This suggests that both strains have narrow ranges of [Ni2+] in which optimal growth efficiency can be achieved.

Cellular Ni concentrations and Ni accumulation in WH8102 and CC9311.
In order to determine directly if Synechococcus adjusts its uptake capabilities for Ni in response to Ni starvation, short-term Ni uptake rates were determined for cultures grown at different [Ni2+]. When supplied with a saturating pulse of Ni (10 nM), Synechococcus rapidly takes up Ni for 30 to 60 min, with declining rates over time (Fig. 6A). Cultures grown at lower Ni concentrations took up Ni more rapidly than those grown at higher Ni (Fig. 6A). Ni uptake by WH8102 can be modeled well by traditional Michaelis-Menten kinetics {{rho} = {rho}max [Ni]/(K{rho} + [Ni])}, where {rho}, {rho}max, and K{rho} are the uptake rate, the saturated uptake rate, and the half saturation constant, respectively). The uptake capacity ({rho}max) appears to be regulated to a greater extent than affinity (K{rho}) by Ni; cultures grown on NH4+ at 100-fold-lower Ni concentrations exhibited a fivefold increase in {rho}max with a less significant change in affinity (2.5 versus 3.5 nM) (Fig. 6B). CC9311 also had a high affinity for Ni (~3 nM), but the changes in {rho}max for similar conditions were less than for WH8102 (2.5-fold versus 5-fold) (data not shown).


Figure 6
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  		FIG. 6. Regulated Ni uptake in WH8102. (A) Time course of Ni uptake by cells grown at pNi 10.3 or pNi 11.3. The error bars are the ranges for duplicate kill-control-corrected cultures. (B) Kinetics of Ni uptake. The symbols represent the measured uptake rates for cells grown at either 0.5 nM (Ni depleted) or 50 nM (Ni replete) total Ni (pNi 12.3 and 10.3, respectively). The curves are the least squares regression fit of the hyperbolic Michaelis-Menten relationship. The fitted {rho}max was 0.35 and 0.07 zetamoles (10–21) Ni cell–1 day–1 for Ni-starved and Ni-replete cells, respectively. K{rho} was 2.5 and 3.5 nM for Ni-starved and Ni-replete cells, respectively.

The cellular concentrations of Ni were determined for both strains during the growth experiments shown in Fig. 4 and were used to provide insight into the regulation of Ni uptake rates and accumulation. For interpretation, the amount of Ni per cell was converted to intracellular molar concentrations (Ni quota [QNi]) by assuming a spherical cell with a radius of 1 µm (Fig. 7). While cellular size or carbon might actually change with decreasing Ni, this was not determined here. The Ni per cell concentrations were also converted into steady-state uptake rates ({rho}ss = µQ, where µ is the specific growth rate) (24) and bioconcentration factors (BCF) (BCF = QNi/NiT, where NiT is the total medium Ni concentration). Finally, {rho}max was calculated using {rho}ss, [Ni2+], a K{rho} value of 3 nM (from Fig. 6), and the Michaelis-Menten equation.


Figure 7
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  		FIG. 7. Cellular QNi, {rho}ss, BCF, and {rho}max for Synechococcus sp. strain WH8102 and CC9311 grown on urea (A), NH4+ (B), and NO3– (C). The error bars are the range of duplicate cultures.

For growth on urea, both strains reduced their cellular Ni quotas with the reduction of [Ni2+] from ~pNi 10.7 to pNi 12 (Fig. 7A). Below pNi 12, the urea-grown cells showed increased Ni uptake capacity (Fig. 7A), maintaining their QNi above 300 nM through heightened uptake rates and BCF (Fig. 7A). A similar trend was observed for the growth of WH8102 on nitrate (Fig. 7C). Elegantly, the observation of heightened Ni uptake rates at low [Ni2+] provides a mechanistic explanation for the sustained growth observed in Fig. 5A and C and rules out Ni contamination as a potential explanation (all Ni added was 63Ni). When grown on NH4+, QNi and {rho}ss in CC9311 varied in a nearly linear fashion dependent upon the extracellular [Ni2+], with little change in {rho}max or BCF until [Ni2+] fell below pNi 12, when a slight increase in the uptake rate was observed (Fig. 7B). WH8102 cells grown on NH4+ increased their BCF and {rho}max in a nearly linear fashion with decreasing [Ni2+] (Fig. 7B), thereby allowing QNi to decrease only 2.2 orders of magnitude over a 3.5-order-of-magnitude change in extracellular [Ni2+]. NO3–-grown CC9311 exhibited only minor changes in {rho}max or BCF. Comparing the strains, CC9311 contained upwards of twofold more cellular Ni than WH8102 and exhibited higher uptake rates at comparable [Ni2+], except at the lower [Ni2+], where the trend was reversed.


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DISCUSSION
 
Ni requirements of marine cyanobacteria.
For growth on urea, our genome-based predictions of the Ni requirements for CC9311 and WH8102 were correct, as both strains became Ni-N colimited when grown without Ni. Biochemically, the Ni-dependent deficiency of urease prevents the cell from acquiring nitrogen, thereby halting growth. While reduced SOD activity was also observed for urea-fed Ni-limited cultures (Table 2), the ability to at least initially recover growth with NH4+ additions (Fig. 3A) suggests that the cell may be able to reallocate intracellular Ni from urease to Ni-SOD with the change in nitrogen source. Note that the recovery of growth with NH4+ additions persists for only a single transfer to NH4+ medium with no added Ni, after which Ni becomes limiting. WH8102 cannot grow on NO3– or NH4– without Ni (Fig. 3B and C), a phenotype attributable to the deficiency in Ni-SOD activity (Table 2). While other marine phytoplankton have been shown to require Ni for growth on urea (28), this is the first marine phytoplankton shown to have an obligate growth requirement for Ni.

After equilibration in media without any added Ni, CC9311 cannot grow well on NH4+ or at all on nitrate, a phenotype contrary to our original hypothesis. While SOD activity was observed in protein extracts from Ni-depleted CC9311 (Table 2), it does not appear to be a completely functional replacement for Ni-SOD, as implied by the Ni-dependent cessation of growth. The light dependency of the Ni limitation observed for NH4+-grown CC9311 (Fig. 4) supports this hypothesis, as superoxide production would be predicted to increase with increasing light. The non-Ni-dependent SOD activity did not allow growth on NO3– without Ni.

In Synechococcus sp. strain PCC7942, the Fe and Mn SODs are localized to the cytoplasm and thylakoid membranes, respectively, with mutants with an inactivated Fe-SOD being more susceptible to methyl viologen, which generates superoxide radicals in the cytoplasm (44). If Ni-SOD in marine cyanobacteria is localized to the cytoplasm, with a separate targeting for Cu/Zn-SOD, cells growing on NO3– without Ni would be particularly susceptible to superoxide radicals created by the reduction of oxygen by nitrate reductase (2).

The competition for intracellular Ni between urease and Ni-SOD may be an important determinant of Ni requirements and the resulting phenotype. The urease operon is upregulated for growth on NO3– relative to NH4+ (41), and NO3–- or urea-grown Synechococcus strains have higher urease activities than NH4+-grown cultures (8). High levels of urease, as well as the attendant Ni-binding metallochaperone encoded by ureE (27), may scavenge Ni away from Ni-SOD. Consistent with this, NO3–-grown cultures are more susceptible to low Ni than NH4+-grown cells (Fig. 7).

The growth experiments conducted over a broad range of [Ni2+] show that WH8102 is affected by decreasing extracellular and intracellular Ni to a greater extent than CC9311. This is surprising, given that WH8102 was isolated from an oligotrophic environment with lower Ni concentrations, but this result is also consistent with CC9311 being able to at least partially employ the non-Ni-dependent SOD activity as a replacement for Ni-SOD. For example, when grown on urea, if CC9311 can partially scavenge superoxide radicals using a Cu/Zn-SOD, then more intracellular Ni can be allocated to urease, whereas WH8102 must devote more of the QNi to Ni-SOD. The greater reduction in the growth rates with decreasing [Ni2+] observed for growth on urea compared to ammonia for both strains is also consistent with the dependence upon a pair of Ni metalloenzymes. The phylogenomic mapping in Fig. 1 shows that the sodN gene (Ni-SOD) does not cooccur within a genome containing sodB (Fe-SOD). The replacement of an Fe-SOD with a Ni-SOD would be a logical evolutionary adaptation allowing the reduction of Fe requirements in an environment where Fe concentrations are low and can be limiting to cyanobacterial growth (22). Indeed, a strain utilizing the Fe-SOD (WH7803) is more susceptible to Fe limitation than Synechococcus sp. strain A2169 (17), and the use of an Ni-SOD by A2169 could be a ready explanation, though the phylogeny of this strain is not known.

We have shown that at least one strain of marine Synechococcus has an obligate growth requirement for Ni, something not previously observed in phytoplankton. Given the requirements of our two model marine cyanobacteria and the complements of Ni metalloenzymes found in cyanobacterial genomes (Fig. 1), we suspect that all marine Prochlorococcus strains also have obligate Ni requirements, while many strains of Synechococcus, Trichodesmium, and Crocosphaera are partially dependent upon Ni for growth. Fe-SOD-containing strains of Synechococcus (WH5701, RCC107, and WH7805) likely do not have a requirement for Ni, except for growth on urea, and WH7803 might have no Ni requirement, as it lacks both Ni-SOD and urease (Fig. 1). The strains containing Fe-SOD instead of Ni-SOD probably have increased Fe requirements as a consequence.

Regulated Ni accumulation by Synechococcus.
To date, Ni-specific transporters or regulatory proteins have not been molecularly characterized in marine phytoplankton. The physiological studies presented here attest to the presence of a regulated Ni uptake system in marine Synechococcus. Both strains actively accumulate Ni in a regulated fashion: both strains concentrated Ni out of the media at 100 to 10,000 times (Fig. 7C), and Synechococcus strains grown at lower [Ni2+] have increased maximal uptake rates (Fig. 6). The increase in Ni uptake rates restored, or at least maintained, growth rates in both strains at low [Ni2+] by maintaining QNi (Fig. 5 and 7). The regulation of the Ni uptake capacity was different for each strain and nitrogen source. In general, much greater changes in {rho}max were observed in WH8102 than in CC9311 (Fig. 7), indicating more stringent regulation or possibly a greater transport potential. For both strains, the greatest changes in {rho}max were observed for growth on urea compared to growth on NO3– or NH4+ (Fig. 7B and D). The dependence of Ni uptake rates upon nitrogen sources has been observed in the marine diatom Thalassiosira weissflogii (37). The nitrogen source-specific differences observed here suggest that the global nitrogen-sensing regulator encoded by ntcA (41) may play a role in modulating Ni uptake.

Using the published cellular carbon and phosphorus quotas for Synechococcus sp. strain WH8103 grown under nutrient-replete conditions (4), we calculated ranges of Ni/C and Ni/P ratios of 5.6 x 10–7 to 5.6 x 10–4 and 1.5 x 10–4 to 1.5 x 10–1 moles mole–1, respectively. The Ni/C ratios observed here bracket the highest Ni concentrations observed for diatoms growing on urea (Ni/C = 1.7 x 10–6 moles mole–1) (37), and Ni/P ratios of 5 x 10–5 to 2 x 10–1 have been observed in Cyanothece sp., a marine nitrogen-fixing cyanobacterium (14). Due to the sheer abundance of cyanobacteria in both coastal and open-ocean ecosystems (33, 40), the Ni/C and Ni/P quotas for Synechococcus indicate that these organisms must contribute to the observed surface depletion in oceanic Ni concentrations. For example, a bloom of cyanobacteria assimilating 0.1 µM P (organic or inorganic) will also take up to 15 nM Ni, well in excess of the typical surface ocean concentration of 3 nM Ni. This would suggest that Ni is recycled within the euphotic zone prior to export to deep waters, or else surface seawater would be entirely depleted of Ni given the calculated nutrient drawdown.

Ni limitation in marine ecosystems?
Environmental relevance is a pertinent question for any culture-based study; to what extent are the results laboratory oddities or ecological realities? The strains used in this study have genomes that are well represented in metagenomic libraries of marine environments (38); therefore, the question lies in the geochemistry of Ni. The few published measurements of total and chemically labile Ni in natural marine systems using competitive ligand exchange adsorptive cathodic-stripping voltametry have suggested that 10 to 60% of the Ni in coastal and open-ocean environments is bound by organic ligands (10, 39). Given these numbers and our measurements of growth rates at fixed [Ni2+] (Fig. 5), the environmental [Ni2+] seems unlikely to limit cyanobacterial growth, except on urea. However, the 2 to 3 nM nickel in surface seawater has been suggested not to be bioavailable based on both ecological stoichiometry (21) and physicochemical speciation measurements (47). Ni also possesses extremely slow coordination kinetics, theoretically retarding the maximum possible biological uptake rates relative to other metals (19). Given the lack of a consensus on the bioavailability of Ni from chemical measurements, it is difficult to use our uptake and growth rate data to confidently argue for or against the possibility of Ni limitation in natural systems at this time.


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ACKNOWLEDGMENTS
 
C.L.D. was supported by a National Defense Science and Engineering Graduate Research Fellowship, and this work was supported by a grant from the Princeton Center for Environmental Bioinorganic Chemistry to C.L.D. and National Science Foundation grants EF0333162 and DBI0542119 to B.P.


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FOOTNOTES
 
* Corresponding author. Mailing address: Mail Code 0202, University of California, San Diego, La Jolla, CA 92093. Phone: (858) 534-7505. Fax: (858) 534-7313. E-mail: bpalenik{at}ucsd.edu Back

{triangledown} Published ahead of print on 19 October 2007. Back


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Applied and Environmental Microbiology, January 2008, p. 23-31, Vol. 74, No. 1
0099-2240/08/$08.00+0     doi:10.1128/AEM.01007-07
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