Introduction

Understanding the global N cycle is critical to ocean biogeochemical models, as nitrogen is arguably the single most limiting nutrient for oceanic primary production. A major current challenge is to determine how N biogeochemistry will change as we transition from the Holocene to the Anthropocene [1]. Nitrogen fixation is one of the key pathways predicted to change as the surface ocean becomes warmer and more acidified [2], [3], [4], [5], [6], [7] and as progressive anthropogenic eutrophication increases fixed N loading in many marine ecosystems [8], [9].

Modeled estimates of N input from marine biological N₂ fixation are dependent on concentrations of other chemical species of fixed N such as nitrate (NO₃⁻) [10], [11]. This is largely because fixed N has been shown in past studies to have relatively strong “inhibitory” effects on N₂-fixation by the ubiquitous oceanic diazotroph Trichodesmium [12], [13], [14], [15], most likely due to differences in the energetic costs involved in assimilating different N species such as NO₃⁻ and N₂ [16]. Several recent laboratory studies, however, have suggested that N₂ fixation by unicellular diazotrophs such as Crocosphaera watsonii may not be as strongly inhibited by NO₃⁻ as has been previously suggested for Trichodesmium [14], [15], [17].

While this major physiological difference may relate to differences in N₂-fixation strategies (Trichodesmium fixes N₂ during the day; Crocosphaera fixes N₂ during the night, similar to unicellular organismal physiology described by Berman-Frank et al. [18]), these recent findings imply that the ratios of N-assimilation kinetic parameters for different N sources (e.g. V_max,N2:V_max,NO3−) may be very different between Trichodesmium and Crocosphaera. In addition to these laboratory-based results, field studies indicate that N₂-fixation rates by unicellular diazotrophs increase with decreasing depth and increasing light in upwelling water where NO₃⁻ concentrations are high [19], [20]. Trichodesmium blooms are also frequently observed in upwelling regions that are known to have high NO₃⁻ concentrations [21]. Lastly, Deutsch et al. [22] presented a model proposing that N₂-fixation rates might be very high in the Peru upwelling system, based on the distribution of phosphorus, despite high concentrations of NO₃⁻ in this region. The general picture of how fixed N sources such as NO₃⁻ control N₂ fixation is still unclear.

In the context of these recent laboratory, field and modeling studies, we asked how the growth rate, as controlled by light, influences preferences for nitrogen substrates (e.g. NH₄⁺, NO₃⁻ and N₂) to support growth of the unicellular N₂ fixer Crocosphaera watsonii. Our data indicate that the N-source utilization ratio (NO₃⁻:N₂) changes in a predictable manner as a function of cell growth. We present experiments suggesting that three key parameters are necessary to determine how fixed N controls N₂-fixation rates by Crocosphaera watsonii: 1) the cellular demand for N, which is largely controlled by the growth rate, 2) the light-specific cellular-assimilation kinetics of the various forms of N (e.g. V_max) and 3) the relative concentrations of the various forms of N. Our basic model relies on the tenet that light energy is the driver of photoautotrophic growth rates while substrates such as NO₃⁻, N₂, PO₄³⁻ etc. do not drive growth but serve as nutrient supports. Thus, a gradient in the light-energy supply rate creates a gradient in the demand for nitrogen to support growth and a gradient in the ratio of nutrient assimilation rates of various nutrient substrates. Our conceptual model may serve as a framework to understand how fixed N availability controls N₂ fixation by oceanic diazotrophs. In light of expected future increases in anthropogenic fixed N inputs to both the coastal and open ocean [23], [24], these studies are needed to improve both physiological models and biogeochemical estimates of global biological N₂ fixation and overall predictions of primary production trends over the next century [10], [25], [26].

Materials and Methods

We investigated short-term and long-term effects of fixed N on N₂-fixation rates by C. watsonii cultures (strain WH0003) in which growth rates were controlled by different light levels. In preparation for both short- and long-term experiments, C. watsonii was pre-acclimated to light environments by growing cultures in triplicate 1-L polycarbonate bottles at 25 and 175 µmol quanta m⁻² s⁻¹ and 28°C, on a 12∶12 hour light:dark cycle for 5 or more generations (as in other laboratory culture experiments; Berman-Frank et al. [18]) with an artificial seawater medium prepared according to the YBCII recipe of Chen et al. [27]. Trace metals (FeCl₃·6H₂O 4.50×10⁻⁷ M, MnCl₂·4H₂O 1.21×10⁻⁷ M, NaMoO₄·2H₂O 1.00×10⁻⁷ M, ZnSO4·7H₂O 7.97×10⁻⁸ M, CoCl₂·6H₂O 5.03×10⁻⁸ M) and vitamins (Thiamine 2.96×10⁻⁷ M, B₁₂ 3.96×10⁻¹⁰ M, Biotin 2.50×10⁻⁹ M) were added with the dilution medium [28] with 4 µM phosphate added as HNa₂PO₄. Cultures were grown with a semi-continuous culturing method as in other studies [14], [18], [29], [30], [31], [32] by diluting cultures every 3 days. Cultures were diluted by enumerating cells and calculating a dilution factor to achieve a target culture cell density of 20×10³ cells mL⁻¹. We determined culture cell densities by agitating cultures just prior to collecting 5 ml of culture and enumerating live cells from subsamples microscopically. Although we did not continuously stir cultures, we did not observe cells or biomass sticking to the sides of the bottles. We calculated growth rates (µ) in between 3-day dilution periods with N_T = N₀e^µT, where N₀ is the cell density at the beginning of a 3-day period (T) and N_T is the cell density at the end of the period.

Long-term exposures

Based on results from our initial short-term experiment with NO₃⁻, we decided to expose Crocosphaera to NO₃⁻ for a longer time period to determine if long-term exposures elicited a different response relative to that in the short-term exposure. In long-term exposures to NO₃⁻, C. watsonii was pre-acclimated to experimental conditions in semi-continuous cultures using NO₃⁻ as a fixed N source (added as 30 µM NaNO₃), in parallel with control cultures growing without an added fixed N source. Particulate organic N of cultures was maintained at similar concentrations by semi-continuous dilution between the control (PN_lowlight = 6.6±3.3 µmoles N L⁻¹; PN_highlight = 7.0±0.8 µmoles N L⁻¹) and added NO₃⁻ treatments (PN_lowlight = 6.7±0.9 µmoles N L⁻¹; PN_highlight = 7.9±0.5 µmoles N L⁻¹). We measured N₂-fixation rates in 50 mL samples from each culture replicate with the acetylene reduction assay as described above at three experimental time points (Table 1). For estimates of NO₃⁻ concentrations, we passed 20 mL of culture through a 0.45 µm syringe filter and NO₃⁻ was measured by the analytical laboratory at the Marine Science Institute, University of California, Santa Barbara, CA, USA. We collected samples to measure the concentration of NO₃⁻ from culture replicates 18 h after the last dilution of cultures (initial measurement) and either 48 h (high-light treatment) or 96 h (low-light treatment) after the initial measurement. To estimate cellular NO₃⁻-assimilation rates, we normalized diminishing NO₃⁻ concentrations during this time to culture cell concentrations that were calculated at the mid-point between these two time points using the growth rate. We did not examine a long-term response to NH₄⁺ exposure primarily because it generally represents a small portion of fixed N relative to concentrations of NO₃⁻ in many natural oceanic waters.

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Table 1. Measurements of culture cell density (cells L⁻¹×10⁶), dissolved nitrate+nitrite concentrations (NO₃⁻ + NO₂⁻, µmol L⁻¹) and N₂-fixation rates (fmol cell⁻¹ hr⁻¹) at different time points (hours since culture dilution) in cultures used in the short- and long-term exposure experiments.

https://doi.org/10.1371/journal.pone.0114465.t001

Results

We observed large differences in growth rates of C. watsonii between light treatments. In control cultures growing on N₂ only, growth was significantly lower in low-light acclimated cultures (25 µmol quanta m⁻² s⁻¹; 0.23±0.02 d⁻¹) relative to cultures growing under higher light (175 µmol quanta m⁻² s⁻¹, 0.68±0.03 d⁻¹; t-test, p<0.05). The controlling effects of NH₄⁺ and NO₃⁻ on N₂ fixation were different in short-term exposures, but varied as a function of growth rate. In addition, the effect of NO₃⁻ on N₂ fixation was similar between short and long-term exposures.

Short-term exposures

In slow-growing cultures acclimated to low light, short-term additions of 0.4 µM NH₄⁺ inhibited N₂-fixation rates to <10% of rates in control treatments without added NH₄⁺ (Fig. 1a). In faster-growing cultures acclimated to 175 µmol quanta m⁻² s⁻¹, with biomass concentrations equivalent to those in low-light cultures (Table 1), short-term exposure to five times as much NH₄⁺ (2.0 µM) was needed to achieve the same inhibitory effect on N₂ fixation (Fig. 1a). The short-term inhibitory effects of NO₃⁻ on N₂ fixation also varied as a function of growth rate. In slow-growing, low-light acclimated cultures, short-term exposure to NO₃⁻ reduced mean N₂-fixation rates by ∼47–62% relative to rates in control treatments without added NO₃⁻ (Fig. 1b). In fast-growing cultures acclimated to high light, however, short-term additions of NO₃⁻ at any concentration up to 40 µM did not inhibit mean N₂-fixation rates by more than 9%, relative to N₂-fixation rates in control cultures without added NO₃⁻ (Fig. 1b).

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Figure 1. Short-term inhibitory effects of ammonium (NH₄⁺, 0–1.5 µmol L⁻¹) and nitrate (NO₃⁻, 0–40 µmol L⁻¹) on N₂ fixation by Crocosphaera watsonii (WH0003) (percent of control with no added nitrogen).

Cultures were grown in steady state under high light (175 µmol quanta m⁻² s⁻¹, growth rate (µ) = 0.68 d⁻¹, open symbols) and low light (25 µmol quanta m⁻² s⁻¹, µ = 0.23 d⁻¹, closed symbols) before adding nitrogen. Error bars represent standard deviations on means from 3 culture replicates.

https://doi.org/10.1371/journal.pone.0114465.g001

Long-term exposures

In high-light-acclimated cultures, long-term exposure to 30 µM NO₃⁻ yielded significantly higher growth rates (µ = 0.87 d⁻¹) than those in control cultures without added NO₃⁻ (µ = 0.68 d⁻¹; p<0.05), indicating that growth was limited by the N₂-assimilation rate (Fig. 2a). Diminishing NO₃⁻ concentrations over time suggested that NO₃⁻-assimilation rates in fast-growing cultures (µ = 0.87 d⁻¹) were 2.8 times higher than those in slow-growing cultures (µ = 0.23 d⁻¹; Fig. 3a; p<0.05), but the contribution of NO₃⁻ to the total daily N assimilation still varied as a function of growth rate. In high-light-acclimated cultures exposed to NO₃⁻ (µ = 0.87 d⁻¹), NO₃⁻ assimilation represented 40% of the total daily N assimilation while N₂ assimilation represented 60% (Fig. 2b). When combined, NO₃⁻ and N₂ assimilation yielded a higher total daily N-assimilation rate (187 fmol N cell⁻¹ d⁻¹) than that in the control treatment growing on N₂ only (122 fmol N cell⁻¹ d⁻¹; p<0.05; Fig. 2b). Furthermore, N₂-fixation rates in cultures with added NO₃⁻ were not significantly different than those in control cultures without NO₃⁻ (p<0.05; Fig. 2b).

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Figure 2. Effects of long-term exposure to 30 µmol L⁻¹ nitrate (NO₃⁻).

(a) cellular growth rates and (b) nitrogen-assimilation rates of Crocosphaera watsonii (WH0003) acclimated to high light (175 µmol quanta m⁻² s⁻¹) and low light (25 µmol quanta m⁻² s⁻¹). (b) N₂-fixation rates (solid bars) are overlain on total N assimilation (N₂ + NO₃⁻ assimilation, hashed bars). Control cultures did not receive added NO₃⁻. Error bars represent standard deviations on means from 3 culture replicates.

https://doi.org/10.1371/journal.pone.0114465.g002

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Figure 3. Changes in nitrate (triangles) and cell concentrations (circles) in cultures of Crocosphaera watsonii (WH0003) in long-term exposure experiments.

Cultures were grown in steady state under (a) high light (175 µmol quanta m⁻² s⁻¹) and (b) low light (25 µmol quanta m⁻² s⁻¹) with added nitrate (30 µmol L⁻¹; open symbols) or with N₂ only (closed symbols). *Calculated NO₃⁻ concentrations (see Methods section for details). Error bars represent standard deviations on means from 3 culture replicates.

https://doi.org/10.1371/journal.pone.0114465.g003

Under low light, long-term exposure to 30 µM NO₃⁻ did not support faster growth rates (Fig. 2a, 3b) even though NO₃⁻-uptake supported 61% of the total daily N assimilation. Instead, N₂-fixation rates were reduced by 55% relative to those in cultures without added NO₃⁻ (p<0.05; Fig. 2a). Thus, in cultures that were grown with NO₃⁻, there was a clear shift in the ratio of N source utilization where growth-specific NO₃⁻-assimilation rates increased by 55% with decreasing light,while growth-specific N₂-assimilation rates increased by 46% with increasing light (Fig. 4). In both the high- and low-light treatments with 30 µM NO₃⁻ added, the concentration of NO₃⁻ was high (>16 µmol NO₃⁻ L⁻¹) throughout the entire 66 h or 114 h sampling period (Fig. 3).

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Figure 4. Growth-specific assimilation rates of nitrate (NO₃⁻; open bars) and dinitrogen (N₂; closed bars) in cultures of C. watsonii (WH0003) with added NO₃⁻ (30 µmol L⁻¹).

Growth-specific NO₃⁻ and N₂-assimilation rates change inversely relative to each other as a function of light-limited growth. Error bars represent standard deviations on means from 3 culture replicates.

https://doi.org/10.1371/journal.pone.0114465.g004

Discussion

Our main finding is that N-source utilization by C. watsonii varied as a function of the growth rate, which we controlled in our experiments with the supply of light energy. Thus, we interpret the variation in N-source utilization (e.g. NO₃⁻:N₂ or NH₄⁺:N₂) to be caused by a gradient in the demand for nitrogen as a substrate to support cell division. This N-source utilization ratio seems to change as a function of energy supply and growth rate because of differences in uptake kinetics between N sources (e.g. V_maxNO3−: V_maxN2) and energy requirements for the reduction and assimilation of each N source.

In our short-term exposure experiment with NH₄⁺, fast-growing cultures of C. watsonii (µ = 0.68 d⁻¹) needed a much higher concentration of NH₄⁺ (5x) to satisfy the nitrogen substrate demand relative to slow-growing cultures (µ = 0.23 d⁻¹; Fig. 1a). An alternate way to view this relationship is that as the amount of the added NH₄⁺ decreased, increasing amounts of N₂ were fixed to satisfy the remaining nitrogen demand to support cell growth (Fig. 1). These results suggest that the magnitude of assimilation of various N substrates depends on the cellular N demand that is needed to support the light-controlled growth rate relative to the light-specific cellular assimilation rate kinetics of each N source. Thus, we propose that when the light-controlled, growth-modulated demand for N exceeds the cellular-assimilation rate of NH₄⁺ or NO₃⁻, N₂ fixation provides fixed N to fill the resulting N deficit.

The variable controlling effects of NO₃⁻ and NH₄⁺ on N₂ fixation suggest that there are large differences in the assimilation kinetics of these different N species (Fig. 1). Under low light, low concentrations of NO₃⁻ and NH₄⁺ (0.5 µM) had maximum inhibitory effects on N₂ fixation, suggesting that the half-saturation constants (K_s) with respect to NO₃⁻ and NH₄⁺ are similar for C. watsonii. The incomplete inhibitory effect of NO₃⁻ on N₂-fixation rates even at high concentrations of NO₃⁻ (Fig. 1b, 3b), however, suggests that the maximum NO₃⁻-assimilation rate (V_max) by C. watsonii (WH0003) is low relative to that of NH₄⁺.

In our long-term experiment, we pre-acclimated Crocosphaera with high NO₃⁻ concentrations (∼15–30 µM, Fig. 3) for 5 or more generations before sampling cultures over a 48–96 h period. In these long-term exposures to NO₃⁻, we measured residual NO₃⁻-concentrations in the culture medium to estimate the cellular NO₃⁻-assimilation rate. The ratio of NO₃⁻ -assimilation:N₂ fixation varied as a function of energy supply and growth (Fig. 2), further supporting these variables as controls of fixed N inhibition of N₂ fixation. Exposure to NO₃⁻ did not affect N₂ fixation by fast-growing cultures of C. watsonii, yet NO₃⁻ comprised 40% of the total daily N, thereby supporting growth rates that were 27% higher than those in control cultures without added NO₃⁻ (Fig. 2b). Thus, the growth of high-light cultures of C. watsonii, similar to Cyanothece, another marine unicellular N₂ fixer [34], was clearly limited by the N₂-assimilation rate, as the addition of 30 µM NO₃⁻ supported higher growth rates (Fig. 2a).

These results indicate that growth rates of C. watsonii benefits from assimilating multiple N sources simultaneously, as individual assimilation rates of N₂ or NO₃⁻ alone cannot support maximum growth rates in high-light environments. Under low light, NO₃⁻-assimilation did not support faster growth as it did under high light, but instead comprised 61% of the total daily assimilated N (Fig. 2). This higher contribution of NO₃⁻ to the total N demand inhibited N₂ fixation by 55% relative to rates in control cultures without added NO₃⁻. Thus, we conclude that the inhibitory effect of NO₃⁻ on N₂ fixation by C. watsonii varies as a function of energy supply and growth rate.

Although we did not separate the direct effect of light-energy supply and growth rate in our long-term experiment, our analyses of the short-term effects of NH₄⁺ and NO₃⁻ exposure on N₂ fixation were done only during dark hours when Crocosphaera fixes N₂. Thus, Crocosphaera offers a unique advantage in comparison with Trichodesmium (which fixes CO₂ and N₂ simultaneously in the light) because it is possible to separate direct effects of light-energy supply from the effects of the light-limited growth rate on N-source utilization preferences. Future experiments might consider experiments that separate these effects by modulating growth rates in other ways.

The assimilation rates of the various chemical forms of N (e.g. NH₄⁺, NO₃⁻, N₂) seem to be dictated in part by the energetic cost of reduction [16]. Many phytoplankton species are known to assimilate NH₄⁺ more easily than NO₃⁻ because of the lower energetic investment associated with assimilating NH₄⁺ [35]. Although N-uptake kinetics have not been described for C. watsonii, Mulholland et al. [29] documented a maximum uptake rate for NH₄⁺ by Trichodesmium that was presumably more than an order of magnitude higher than that for NO₃⁻. Based on the relatively weak inhibitory effect of NO₃⁻ on N₂ fixation by C. watsonii relative to that observed for NH₄⁺ (Fig. 1, 3), we infer that the maximum assimilation rate of NO₃⁻ by C. watsonii (V_max) must be considerably lower than that of NH₄⁺.

Although NH₄⁺ assimilation carries a cost associated with transport across the cell membrane, it is generally thought to be less expensive to assimilate than NO₃⁻ and N₂ [36], [37] because of the high costs associated NO₃⁻ and N₂ assimilation, which must first be reduced to NH₄⁺ before being assimilated onto glutamic acid (ΔG  = +69 Kcal mol N⁻¹ for NO₃⁻ and +87 Kcal mol N⁻¹ for N₂) [16]. A lower assimilation cost for NH₄⁺ might afford a high V_max relative to that for more energetically expensive forms of nitrogen. Thus, the lower cost associated with NO₃⁻ reduction to NH₄⁺ relative to N₂ reduction to NH₄⁺ appears to benefit C. watsonii in a light-limited environment where growth is slow relative to a maximum NO₃⁻-assimilation rate (Fig. 4). In a high-light environment, the maximum assimilation rate of NO₃⁻ relative to the growth rate is reduced in comparison with that in low-light cultures (Fig. 4), where N₂ supports a higher portion of the daily N demand for growth. Future studies should quantify NO₃⁻-assimilation kinetics for N₂ fixers and identify how they might change as a function of other environmental conditions.

In addition to the energetic costs for reducing NO₃⁻ and N₂, the difference between energetic and material investments associated with the production of assimilatory proteins such as nitrogenase and nitrate reductase may be at least partially responsible for the differential ratios of NO₃⁻:N₂ reduction as function of growth. Tradeoffs in energetic investments for NO₃⁻ and N₂ reduction may come from balancing differential cellular nitrogen demands that are associated with variable growth rates [38] or from the supply of light. Further separating the effect of light-energy supply from the effect of growth on the ratio of fixed N:N₂ utilization may lead to a better understanding of the release of fixed N by diazotrophs [5], [33], [39], [40], [41].

Contrary to findings by Ohki et al. [12] that suggest a strong time dependence of exposure to NO₃⁻, NH₄⁺ and urea in controlling inhibitory effects on N₂ fixation in Trichodesmium, we documented consistent inhibitory effects of NO₃⁻ on N₂ fixation of Crocosphaera regardless of the duration of exposure. The results presented by Ohki et al. [12] are difficult to interpret in a context of supply and demand for N, however, because growth rates between treatments were not defined.

Although previous studies have not discussed inhibitory effects of fixed N on N₂ fixation in a context of the supply rate of fixed N relative to the growth-modulated demand for N, four relatively recent studies have collectively examined inhibitory effects of fixed N on N₂ fixation in batch cultures of Crocosphaera and/or Trichodesmium growing under 30–40, 80, 128 and 180 µmol quanta m⁻² s⁻¹, all at 26 or 27°C [14], [15], [17], [42]. In batch cultures, the biomass concentration of the culture is important to consider because of the accelerating effect of increasing biomass on the rate of disappearance of NO₃⁻ or NH₄⁺. Interpretation of these studies in a context of the supply rate of fixed N relative to the growth-modulated demand for N is also difficult, mainly because biomass and/or growth rates between treatments were not defined during batch-mode growth.

In our experiments, we maintained constant exponential growth rates with a semi-continuous culturing method and we maintained equivalent biomass concentrations between treatments so that differences in NH₄⁺ and NO₃⁻ drawdown due to biomass differences would not affect cellular N₂-fixation rates between treatments and between time points (Fig. 3; Table 1). In addition to our experiments with Crocosphaera, all of these previous studies indicate that NO₃⁻ and/or NH₄⁺ have controlling effects on N₂ fixation by oceanic N₂ fixers. Future studies that examine N-source preferences should focus on growth-modulated controls of fixed N on N₂ fixation in both Trichodesmium and Crocosphaera. Although we presume that this model would be similar for Trichodesmium, there may be unforeseeable differences due to the major differences between the physiological mechanisms that these species use to separate oxygen generated by photosynthesis from the nitrogenase enzyme; Trichodesmium seems to use a spatial separation mechanism, as it fixes both inorganic carbon and N₂ during the light period. In contrast, Crocosphaera uses a temporal separation mechanism, as it stores fixed carbon during the light period and respires it for energy during the night to fuel N₂ fixation in the dark, similar to the unicellular strategy described by Berman-Frank et al. [18].

In the open ocean, the primary limiting nutrients for growth of N₂-fixing cyanobacteria are iron (Fe) and phosphorus (P) [43], [44]. In combination with light, Fe and P have an indirect effect on N demand through their support of cellular growth. Capone and Knapp [8] originally proposed that the N:P ratio is important in controlling N₂-fixation rates, and recently Ward et al. [11] suggested that the N:Fe ratio is a dominant controlling factor of marine N₂ fixation. Our basic model suggests that the ratio of N:X is important in controlling N₂-fixation rates where “X” is a resource that influences growth rates (such as light, P and Fe), and thereby, the demand for N. Laboratory data support this, where high concentrations of P supported high N₂-fixation rates relative to cultures with lower P concentrations, despite equivalent N:P supply ratios [15]. In a modeling study, Ward et al. [11] demonstrated that the N:P supply ratio is a secondary factor in defining boundaries of N₂ fixation, while the N:Fe supply ratio is more important in an ecological context through competitive interactions with non-N₂-fixing phytoplankton. Further, Garcia et al. [32] suggest that the Fe:P supply ratio may be more important in controlling N₂ fixation than the absolute concentration of either of these limiting nutrients. Collectively, these studies suggest that links between C, N, P and Fe biogeochemical cycles depend on the relative supply of each of these nutrients and our study further suggests that the energy-supply rate or the growth rate modulates interactions between these nutrients.

Our study indicates that global models of marine biological N₂ fixation should consider an interaction between assimilation kinetics of fixed N and a growth-modulated demand for N. Although our study did not focus on how Crocosphaera might respond in the natural environment, our data provide a framework around which future studies might structure investigations of N-source preferences by natural communities of N₂ fixers. Reactive nitrogen from atmospheric sources and agricultural runoff are expected to increase in the future and the effects of increased N input to the oceans on phytoplankton communities is uncertain [23], [24], [45]. Thus, a clear understanding of how reactive nitrogen affects N₂ fixation is needed to support predictions of how phytoplankton communities will change.

Two other relevant environmental factors that will certainly influence growth of N₂ fixers in the future are CO₂ and temperature [4], [5], [6], [7], [30], [34]. Both of these factors are predicted to increase, and will likely influence the controlling effects of fixed N on N₂ fixation through their effects on growth rates. Thus, our basic framework potentially has far-reaching implications for both current estimates of oceanic N₂ fixation, and for estimates of N₂-fixation rates that are likely to exist in the future surface oceans [3].

Acknowledgments

We thank Eric Webb for providing the isolate of WH0003 that we used in this study.