Study on the Characteristics and Mechanism of Inorganic Nitrogen Nutrients Preferential Assimilation by Phaeocystis globosa

Abstract

Phaeocystis globosa is an important harmful algal species that is globally distributed. Previous studies have indicated that P. globosa preferentially uptake nitrate, but the underlying mechanism is still unclear. To further verify this preference and reveal the underlying mechanism, the assimilation rates of nitrate and ammonium by P. globosa at different concentrations was quantitatively studied by using a nitrogen stable isotope (¹⁵N) tracer technique, and the regulatory mechanism was determined from the physiological and biochemical responses. The findings revealed that the preferential assimilation of nitrate by P. globosa was influenced by the ambient ammonium concentration. When the ambient concentration of ammonium was less than approximately 3.5 μmol·L⁻¹, the assimilation rates of nitrate form P. globosa were as high as 1.05 × 10⁻⁵ μmol·L⁻¹·h⁻¹·cell⁻¹. Even though the nitrate assimilation in P. globosa was inhibited at ammonium concentrations greater than this threshold, nitrate assimilation could not be completely suppressed. The activity of NR and the expression of related genes in P. globosa were also affected by ammonium. In addition, ¹⁵N signals originally labeled nitrate accumulated in ammonium. This indicated that P. globosa was likely to reduce excess nitrate to ammonium and subsequently release it into the substrate, which might be an important energy dissipation mechanism for P. globosa. These results expand the classical understanding of the utilization of nitrogen nutrients by marine phytoplankton, and offer valuable resources for comprehending the mechanism of harmful algal blooms.

1. Introduction

There are two forms of Phaeocystis globosa: solitary free-living cells and colonies [1]. Colonies consist of numerous flagellate-free cells that are embedded within an elastic membrane [2]. The colonies serve as the primary forms of algal blooms. The membrane of these colonies is composed of polysaccharides, which serve to protect against viral infections and consumption by zooplankton, as well as to store nutrients [3,4]. The presence of colonies is one of the reasons why this algal species competes well with other algae. Extensive blooms of P. globosa have been observed in various locations across Europe, the southeastern shores of Vietnam, and the southern coastline of China [2,5,6]. The blooms of P. globosa have been shown to produce hemolytic toxins, clog fish gills, and cause the death of planktonic organisms [2]. A large amount of white foam is produced due to P. globosa blooms, which leads to the deterioration of the aquatic environment [7,8]. Reports suggest that colonies pose a threat to the safety of nuclear power plants by blocking cooling systems [9]. Therefore, P. globosa blooms exert a profound influence on the structure and function of coastal ecosystems, and greatly hinder social and economic development.

Nitrogen (N) serves as the foundation of marine life. Ammonium (NH₄⁺) and nitrate (NO₃⁻) play crucial roles as nutrients for marine phytoplankton. According to previous studies, NH₄⁺ is considered the preferential N form for phytoplankton [10,11], which is determined by the physiological characteristics of microalgae. In terms of energy, algae consume less energy by assimilating NH₄⁺ than NO₃⁻. Therefore, NH₄⁺ is generally taken up by algae first, and only after its near depletion is NO₃⁻ taken up [12,13]. The NH₄⁺ preference for phytoplankton has been one of the classical physiological theories, and has been grounded in many studies. For instance, when the concentration of NH₄⁺ exceeds 1~2 μmol·L⁻¹, the uptake of NO₃⁻ by microalgae in Chesapeake Bay never exceeds 1% of the total N [14]. A previous study revealed that algae assimilated NO₃⁻ only when the concentration of NH₄⁺ decreased below 0.5~1 μmol·L⁻¹ [15]. Wheeler (1990) reported that NH₄⁺ completely inhibited the uptake of NO₃⁻ within the range of 0.1~0.3 μmol·L⁻¹ [16].

However, the NH₄⁺ preference may also depend on the algal species. Previous research on P. globosa revealed that NO₃⁻ significantly contributed to the growth and formation of colonies. In contrast to NH₄⁺ and urea, P. globosa displayed a greater propensity for colony formation when utilizing NO₃⁻ as the substrate [17], and the effect of NO₃⁻ on P. globosa has been shown to be significant (p < 0.05) [18]. Many field experiments have also shown that P. globosa blooms in most marine areas, such as the Belgian coastal zone and Ross Sea, are accompanied by high NO₃⁻ consumption [19,20,21,22]. Notably, Lv (2019) established the NO₃^—enriched group, the NH₄⁺-enriched group, and the NO₃⁻- and NH₄⁺-enriched group (both-enriched group), and it was reported that the uptake rate of NO₃⁻ was greater than that of NH₄⁺. NH₄⁺ accumulates in the fluid inside colonies of P. globosa [23]. These relevant results contradict the classical physiological theory of NH₄⁺ preference, challenging the in-depth understanding of the mechanism of P. globosa blooms.

Here, we speculate that the preference for NO₃⁻ for P. globosa may present as a high threshold of NH₄⁺ for inhibiting the uptake of NO₃⁻, while previous research failed to reach this threshold. Additionally, the accumulation of P. globosa intracolonial NH₄⁺ may originate from the excessive reduction of NO₃⁻ to NH₄⁺, but there is no direct evidence for this phenomenon. Therefore, this study expanded the substrate NH₄⁺ concentrations to study the threshold of NH₄⁺ for inhibiting the assimilation of NO₃⁻. The assimilation rates of NH₄⁺ and NO₃⁻ by P. globosa were accurately calculated synchronously by using ¹⁵N labeling for NO₃⁻ alone. Moreover, the accumulation of ¹⁵NH₄⁺ was monitored to verify the hypothesis that NH₄⁺ was released from excessive NO₃⁻ reduction. Enzyme activity and related gene expression were used to elucidate the relevant mechanism of action. These results clarify the ecological adaptation mechanism of P. globosa in terms of N utilization, providing a theoretical basis for understanding the formation and prevention of P. globosa blooms.

2. Materials and Methods

2.1. Algae Culture

P. globosa was procured from the northern region of the Beibu Gulf. All culture experiments used a modified formula of artificial seawater [24], which was sterilized at 121 °C for 30 min. P. globosa was cultured under conditions of 20 ± 1 °C and 50 μmol·m⁻²·s⁻¹ illuminance with a 12 h light: 12 h dark cycle.

2.2. Experimental Setup

The P. globosa culture was at the exponential growth stage. Then, the culture was diluted with artificial seawater sterilized at 121 °C at an algal culture/seawater ratio of 2:7 and acclimated through indoor culture for 3 to 4 days, after which the concentration of NO₃⁻ was monitored every day. After the concentration of NO₃⁻ was completely consumed, 12 μmol·L⁻¹ NO₃⁻ was added, and P. globosa were cultured for 2 days until the concentration of NO₃⁻ was lower than the detection limit and the algal density was 7.5 × 10⁵ cells·mL⁻¹ (the concentration of PON was 43~78 μmol·L⁻¹). The cultures were divided into three groups: (a) 20 μmol·L⁻¹ NO₃⁻ and 5 μmol·L⁻¹ NH₄⁺ (58% biomass N); (b) 20 μmol·L⁻¹ NO₃⁻ and 10 μmol·L⁻¹ NH₄⁺ (43% biomass N); and (c) 20 μmol·L⁻¹ NO₃⁻ and 20 μmol·L⁻¹ NH₄⁺ (51% biomass N). The above three groups of experiments were named group 1, group 2, and group 3, respectively. NO₃⁻ in the three mixed N media all contained 5% NaNO₃ (≥98 atom%, Sigma-Aldrich, St. Louis, MO, USA), with a high abundance of ¹⁵N as a tracer of the N cycle. The added algal culture mixture was shaken well and divided into 75 square breathable cell culture bottles (75 cm², Nest) for the experiment. The experimental environment was the same as that of the algal culture. The nutrient solution was added at the start of the experiment, seven time points were sampled at 0, 2, 4, 6, 8, 10, and 12 h, and three parallel points were designed for each time point to determine the parameters. The sampling points of each experimental group involved three parallel sets. Destructive sampling was used in all experiments.

2.3. Sample Collection

The N mainly included NO₃⁻, NH₄⁺, and NO₂⁻. The algal culture was filtered through membranes (Whatman GF/F). The filtrate was frozen in a −20 °C freezer until analysis by an automatic nutrient analyzer.

The stable N isotope standards included dissolved δ¹⁵N-NH₄⁺ and granular δ¹⁵N-PON. The algal culture was filtered through membranes heated at 450 °C for 4 h. The filtrate was frozen at −20 °C for freezing preservation for the detection of δ¹⁵N-NH₄⁺. The filter membrane was securely encased in clean aluminum foil and kept at a temperature of −20 °C for storage until the determination of the δ¹⁵N-PON value and concentration of the PON.

For enzyme activity and gene analysis, 150 mL of algal culture was collected at 0 h, 4 h, and 10 h in groups 1 and 3, and 150 mL of algal culture was collected at 0 h, 2 h, and 10 h in group 2. The algal culture was centrifuged at 4 °C. Then, the culture was centrifuged at 8000× g for 10 min to collect the algal cells. Samples of algal cells were immediately transferred to liquid nitrogen for flash freezing, and transferred to a freezer at −80 °C for storage until analysis.

2.4. Sample Analysis

The concentrations of NO₃⁻, NH₄⁺, and NO₂⁻ in the samples were determined by means of a continuous-flow analyzer (Skalar, Breda, The Netherlands). The detection ranges of NO₃⁻, NH₄⁺, and NO₂⁻ were 0.07~35.7 μmol·L⁻¹, 0.07~35.7 μmol·L⁻¹, and 0.07~14.3 μmol·L⁻¹, respectively. The data were evaluated using commercial nutrient reference material to control the data quality.

To determine the concentration of PON and its stable N isotope, the filter film sample was first wrapped in aluminum foil and loaded into a freeze-dryer for more than 48 h. Subsequently, the dried sample was wrapped in microspheres with 35 × 35 mm² aluminum foil (Elementar, Hanau, Germany), and the sample was compacted by a tablet press, which removed any air.

The δ¹⁵N-PON value and PON value were determined by a Thermo Fisher EA Flash element analyzer and a Delta Q isotope ratio mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). Each lot was calibrated using a single point of the USGS-41 reference material. The δ¹⁵N-NH₄⁺ value was ascertained using a Gas Bench MAT253 stable isotope mass spectrometer. The δ¹⁵N values in PON and NH₄⁺ are expressed relative to the content of atmospheric N according to the following equation: δ¹⁵N (‰) = (R_sample − R_standard)/R_standard × 1000, where R = ¹⁵N/¹⁴N and atom% = ¹⁵N/(¹⁵N + ¹⁴N) × 100.

Enzyme activity was determined by evaluating the activities of nitrate reductase (NR) and glutamine synthetase (GS). First, the cells were crushed using an ultrasonic cell crusher (Scientz, Ningbo, China). Subsequently, the activity of the NR detection kit (Solarbio, Beijing, China) was used to determine the activity of NR, and a GS kit (Solarbio, Beijing, China) was used to determine the activity of GS, both of which were utilized according to the kit instructions. The samples for qPCR were sent to Shengong Bioengineering (Shanghai, China) Co., Ltd., for RNA extraction and qPCR analysis.

2.5. Data Processing

The formula for calculating the growth rates is as follows:

$$\mathsf{\mu }=\frac{\mathrm{L}\mathrm{n}{\mathrm{N}}_2-\mathrm{L}\mathrm{n}{\mathrm{N}}_1}{{\mathrm{t}}_2-{\mathrm{t}}_1}$$

(1)

where N₁ and N₂ represent the algal density and t₁ and t₂ represent the time. In this study, the time interval was 0.5 days.

The uptake rate of N in this study was determined based on the modified uptake rate formula of Mulholland et al. (2006) [25]:

$${\mathrm{V}}_{{{\mathrm{N}\mathrm{O}}_3}^{-}}=\frac{{\left(\mathrm{a}\mathrm{t}\mathrm{o}\mathrm{m}\mathrm{\%}\mathrm{P}\mathrm{N}\right)}_{\mathrm{f}\mathrm{i}\mathrm{n}\mathrm{a}\mathrm{l}}-{\left(\mathrm{a}\mathrm{t}\mathrm{o}\mathrm{m}\mathrm{\%}\mathrm{P}\mathrm{N}\right)}_{\mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l}}}{(\mathrm{a}\mathrm{t}\mathrm{o}\mathrm{m}\mathrm{\%} \mathrm{N} \mathrm{s}\mathrm{o}\mathrm{u}\mathrm{r}\mathrm{c}\mathrm{e} \mathrm{p}\mathrm{o}\mathrm{o}\mathrm{l}-{\mathrm{a}\mathrm{t}\mathrm{o}\mathrm{m}\mathrm{\%}\mathrm{P}\mathrm{N})}_{\mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l}}\times \mathrm{t}\mathrm{i}\mathrm{m}\mathrm{e}}\times \mathrm{P}\mathrm{N}$$

(2)

$${\mathrm{V}}_{{{\mathrm{N}\mathrm{H}}_4}^{+}}=\frac{\mathrm{P}\mathrm{N}}{\mathrm{t}\mathrm{i}\mathrm{m}\mathrm{e}}-{\mathrm{V}}_{{{\mathrm{N}\mathrm{O}}_3}^{-}}$$

(3)

where (atom%PN)_final is the final atomic particle N value, (atom%PN)_initial is the initial atomic particle N value, the source pool is the enrichment dissolved N pool, PN is the particle N on the GF/F membrane (μmol·L⁻¹) (all the above data can be obtained from the isotope mass spectral data), and time is the culture time (h).

This formula is used to determine whether the increase in ammonium isotopes is due to the influence of processes in assimilation (Section 4.3). The isotopic fractionation (ε) of the NH₄⁺ assimilation and utilization processes was calculated by the Rayleigh model. The formula is as follows:

$${\mathsf{\delta }}_{\mathrm{s}\mathrm{u}\mathrm{b}\mathrm{s}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{e}}\left(f\right)={\mathsf{\delta }}_{\mathrm{s}\mathrm{u}\mathrm{b}\mathrm{s}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{e}}(f=1)-\epsilon \times lnf$$

(4)

where f represents the residual ratio of the reactant, and δ_substrate(f) and δ_substrate(f = 1) represent the stable isotope ratios of the reactant at that moment and at the initial stage, respectively.

Furthermore, the 64-bit version of the Origin 2018 software was used for graph generation.

3. Results

3.1. Growth of P. globosa

According to the growth curves (Figure 1b), the algal density of the three groups increased with time. Specifically, before 6 h, the algal density increased slowly, and after 6 h, the rapid growth in density was observed. According to the growth rate chart (Figure 1b), the growth rates of group 3, group 2, and group 1 increase successively, and the lowest growth rate of group 1 is 0.88 d⁻¹.

3.2. Variations in Inorganic N Nutrient Concentrations

In this study, the concentrations of NO₃⁻ and NH₄⁺ exhibited varying degrees of reduction over time. Specifically, in group 1, the concentration of NO₃⁻ decreased from an initial value of 19.99 μmol·L⁻¹ at 0 h to 0.62 μmol·L⁻¹ at 8 h, subsequently stabilizing at approximately 0.6 μmol·L⁻¹. The concentration of NH₄⁺ decreased slightly from 5.17 μmol·L⁻¹ at 0 h to 2.06 μmol·L⁻¹ at 4 h, followed by fluctuations within the range of 1.1~1.5 μmol·L⁻¹. The concentration of NO₂⁻ increased slightly initially, followed by a gradual decrease, maintaining a concentration between 0.05~0.6 μmol·L⁻¹ (Figure 2a). In group 2, the concentration of NO₃⁻ decreased slightly from 21.42 μmol·L⁻¹ at the initial level to 20.11 μmol·L⁻¹ at 2 h, then rapidly decreased to 1.68 μmol·L⁻¹ at 8 h, and further decreased to below 0.85 μmol·L⁻¹. The concentration of NH₄⁺ decreased from 9.22 μmol·L⁻¹ at 0 h to 2.09 μmol·L⁻¹ at 2 h and then remained stable. The concentration of NO₂⁻ increased slightly and then decreased, and the overall concentration was lower than 1 μmol·L⁻¹ (Figure 2b). In group 3, the concentration of NO₃⁻ changed little in the first 4 h, from 21.33 μmol·L⁻¹ to 20.02 μmol·L⁻¹, and then rapidly decreased to 3.98 μmol·L⁻¹ at 10 h. The concentration of NH₄⁺ decreased significantly from 22.18 μmol·L⁻¹ at the beginning to 3.51 μmol·L⁻¹ at 4 h and continued to decrease from 6~12 h, with the lowest concentration decreasing to 1.21 μmol·L⁻¹. The concentration of NO₂⁻ remained below 0.1 μmol·L⁻¹ throughout the experiment (Figure 2c).

3.3. Variation in Uptake Rates

The maximum uptake rate of NO₃⁻ was 5.28 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹ at 4~6 h (Figure 3a). In group 2, the uptake rate of NH₄⁺ was greater than that of NO₃⁻ for the initial 6 h. The largest and smallest uptake rates of NH₄⁺ in group 2 were 4.41 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹ and 3.96 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹, respectively (Figure 3b). In group 3, the uptake rate of NO₃⁻ was higher than that of NH₄⁺ during the 6~10 h (Figure 3c). Notably, the maximum uptake rate of NO₃⁻ in group 3 reached 1.05 × 10⁻⁵ μmol·L⁻¹·h⁻¹·cell⁻¹, while the maximum uptake rate of NH₄⁺ was 5.94 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹.

3.4. Variations in N Isotopes and Isotopic Fractionation (ε)

In this study, the δ¹⁵ N-PON values in all three groups significantly and continuously increased over time. In group 1, the δ¹⁵N-PON value increased rapidly from 4.4‰ at 0 h to 340.3‰ at 8 h, after which the increase slowed and reached its highest value (373.1‰) at 12 h (Figure 4a). In group 2, the overall change characteristics of the δ¹⁵ N-PON values were similar to those in group 1, showing a significant increase in the first 8 h, after which the increasing trend slowed. However, it is worth noting that the δ¹⁵ N-PON value in group 2 in the first 4 h was lower than that in group 1 at the same time. The δ¹⁵N-PON value in group 2 was greater than that in group 1 at 6 h until the results were similar at 8~12 h, with a maximum value of 368.5‰ (Figure 4b). In group 3, the increasing trend of δ¹⁵N-PON in the first 4 h was slower than that in the other two groups, increasing to only 122.3‰. Subsequently, the δ¹⁵N-PON value increased significantly until it reached the highest value of 283.1‰ at 12 h (Figure 4c).

The δ¹⁵N-NH₄⁺ values of the three groups of experiments first increased and then decreased with time. In group 1, the δ¹⁵N-NH₄⁺ value increased to 436.1‰ at 2 h and then fluctuated in the range of 60 to 420‰. After 6 h, the δ¹⁵N-NH₄⁺ value began to decline rapidly, reaching 61.32‰ at 10 h and 76.9‰ at 12 h (Figure 5a). In group 2, the δ¹⁵N-NH₄⁺ value showed an obvious upward trend in the first 4 h, reaching a maximum of 268.6‰, and then continued to decrease to 73.4‰ at 12 h (Figure 5b). In group 3, the δ¹⁵N-NH₄⁺ value continuously increased in the first 6 h, peaking at 271.3‰ and then continuing to decrease, and the δ¹⁵N-NH₄⁺ value decreased to 112.8‰ at 12 h (Figure 5c).

In the variation of isotopic fractionation (ε) of δ¹⁵N-NH₄⁺ (Figure 6), it was found that the isotopic fractionation (ε) of group 1, group 2, and group 3 was 2094.3‰,364.0‰, and 163.4‰, respectively, with the largest ε in group 1.

3.5. The Activities of NR and GS and the Expression of NR and GS Genes

In group 1, the activity of NR was 0.43 U/mg prot at 0 h, after which it increased first. The activity of GS at 0 h was 2.22 U/mg prot, which decreased significantly and remained at approximately 1.3 U/mg prot from 4~10 h (Figure 7a). This was related to nutrient consumption, and the activities of the two enzymes decreased when nutrients were almost exhausted at the late stage of the experiment. Their expression levels were downregulated at 4 h, and both the NR and GS gene expression levels were upregulated at 10 h (Figure 8a). In group 2, the activity of NR tended to increase. After 2 h, the change in activity was small; it remained at approximately 0.31 U/mg prot and subsequently showed a downward trend (Figure 7b). In group 2, the NR and GS gene expression levels were downregulated (Figure 8b). In group 3, the activity of NR increased significantly to 1.72 U/mg prot at 10 h. The activity of GS at 0 h was 2.04 U/mg prot after which it decreased first (Figure 7c). The gene expression level of GS and NR also tended to first decrease and then increase, reaching the highest relative expression level at 10 h (Figure 8c).

4. Discussion

4.1. The Characteristics of NO₃⁻ and NH₄⁺ Assimilation by P. globosa

4.1.1. Comparison of the Assimilation Rates of NO₃⁻ and NH₄⁺

This study revealed that the preferential assimilation of NO₃⁻ by P. globosa is influenced by the ambient concentration of NH₄⁺. In groups 1 and 2, as the concentration of NH₄⁺ decreased, the assimilation rate of NO₃⁻ gradually increased (Figure 3a,b). However, the assimilation of NO₃⁻ at 0~4 h was influenced by the ambient concentration of NH₄⁺ in group 3. After 6 h, the maximum assimilation rate of NO₃⁻ increased to 1.05 × 10⁻⁵ μmol·L⁻¹·h⁻¹·cell⁻¹, and the concentration of NO₃⁻ increased to 6.22 μmol·L⁻¹ (Figure 2c and Figure 3c). These findings indicated that when both NO₃⁻ and NH₄⁺ were present in the environment and the concentration of NH₄⁺ exceeded a certain threshold, the preferential assimilation of NO₃⁻ by P. globosa was modulated by the concentration of NH₄⁺. When the threshold was reached, the maximum assimilation rate of NO₃⁻ was greater than that of NH₄⁺, and the assimilation of NO₃⁻ was preferred at the later stage of the experiment.

When the concentration of NH₄⁺ in the environment was reduced to a certain extent, P. globosa notably exhibited NO₃⁻ uptake. For example, in group 3, when the concentration of NH₄⁺ was reduced to 3.51 μmol·L⁻¹, the concentration of NO₃⁻ decreased rapidly from 14.10 to 1.17 μmol·L⁻¹ at 10 h, while the concentration of NH₄⁺ decreased by only 1.61 μmol·L⁻¹ during the same period (Figure 2c). The uptake rate of NO₃⁻ began to increase at 4~6 h and rapidly increased to 1.05 × 10⁻⁵ μmol·L⁻¹·h⁻¹·cell⁻¹ at 6~8 h (Figure 3c). Compared with the previous experimental data of the other two groups, the uptake rate of NO₃⁻ in group 3 was lower. In group 2, when the concentration of NH₄⁺ was reduced to 2.09 μmol·L⁻¹ within 2 h, it remained in the range below 1.60 μmol·L⁻¹, while the concentration of NO₃⁻ decreased from 20.11 μmol·L⁻¹ to 0.67 μmol·L⁻¹ during this period (Figure 2b). The uptake rate of NO₃⁻ increased to 1.70 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹ at 2~4 h, which was consistent with the change in nutrients (Figure 3b). Through further verification by design group 1, it was found that the concentration of NH₄⁺ decreased by only 4.05 μmol·L⁻¹ during the entire experiment, while the concentration of NO₃⁻ continued to decrease rapidly and was mostly consumed at 8 h (Figure 2a). The study showed that the uptake rate of NO₃⁻ reached a maximum of 5.28 × 10⁻⁶ μmol·L⁻¹·h⁻¹·cell⁻¹ at 4~6 h (Figure 3a). This result is related to the preferential uptake of NO₃⁻ by P. globosa reported in many previous studies [20,23]. Malone’s research revealed that the uptake rate of NO₃⁻ by Chaetoceros sp. was 0.20~4.00 × 10⁻⁹ μmol·L⁻¹·h⁻¹·cell⁻¹ [26]. The rate of NO₃⁻ uptake by Pseudo-Nitzschia delicatissima cultured in artificial seawater is 1.48~1.77 × 10⁻⁹ μmol·L⁻¹·h⁻¹·cell⁻¹ [27]. The uptake rate of NO₃⁻ by Trichodesmium sp. is less than 1.75 × 10⁻⁹ μmol·L⁻¹·h⁻¹·cell⁻¹ [28]. Additionally, a study revealed higher uptake rates of NO₃⁻ in Nitzschia sp., and the uptake rate ranged from 2.64~5.71 × 10⁻⁹ μmol·L⁻¹·h⁻¹·cell⁻¹ [29]. The assimilation rate of nitrate in P. globosa was greater than that in other algae.

4.1.2. The Inhibitory Effect of Ambient Concentrations of NH₄⁺ on the Assimilation of NO₃⁻

According to the above results, it is speculated that when the concentration of NH₄⁺ in the environment is lower than approximately 3.5 μmol·L⁻¹, P. globosa will begin to assimilate and utilize NO₃⁻, which is considered delayed uptake. In this study, the inhibitory effect of NH₄⁺ on the uptake or assimilation of NO₃⁻ ranged from 3.51~22.18 μmol·L⁻¹. Increased concentrations of NH₄⁺ have been found in experiments in Central Bay to lead to delayed uptake of NO₃⁻ [30]. Some scholars believe that NH₄⁺ may hinder the utilization of NO₃⁻ by decreasing the availability of ATP, which is crucial for the active transport of NO₃⁻ [31]. The process by which NH₄⁺ inhibits the utilization of NO₃⁻ has been shown to be highly unpredictable [32]. Indeed, the extent to which NH₄⁺ inhibits NO₃⁻ metabolism, along with its threshold concentration, varies significantly based on species of algae, physiological state, and specific environmental conditions [20]. In Eppley’s study, N was assimilated by Phaeocystis sp. only when the concentration of NH₄⁺ was lower than 1 μmol·L⁻¹ [15]. When the concentration of NH₄⁺ exceeds 1~2 μmol·L⁻¹, the uptake of NO₃⁻ by microalgae in Chesapeake Bay never exceeds 1% of the total N [14]. The concentration of NH₄⁺ in San Francisco Bay must be reduced to 1~4 μmol·L⁻¹ for NO₃⁻ to be rapidly assimilated by phytoplankton [30]. In this study, P. globosa changed from NH₄⁺ to NO₃⁻ when the concentration of NH₄⁺ was 1~3.5 μmol·L⁻¹, which may be conducive to the production of colonies and may aid in the survival and competition of P. globosa.

Notably, although it is inferred above that the prophase P. globosa in group 2 and group 3 preferentially assimilated and used NH₄⁺, the relatively high concentration of NH₄⁺ in the environment did not completely repress the assimilation of NO₃⁻. On the one hand, the concentration of NO₃⁻ still slightly decreased in these groups in the early stage of these experiments; on the other hand, the ¹⁵N-PON value increased significantly. According to the fractionation effect of stable isotopes, when there is a natural abundance of N without the addition of a tracer, lighter ¹⁴N will have priority in the PON pool in the process of microalgal assimilation [33], resulting in a decrease in the δ¹⁵N-PON value. However, when a certain N in the environment is added with a high abundance of ¹⁵N, the assimilation of N by algae will cause highly labeled ¹⁵N signals to enter the PON pool, resulting in an increase in the δ¹⁵N-PON value [34]. There are two possible reasons for this persistent assimilation of NO₃⁻. On the one hand, Glibert reported that the assimilation of N and C is connected through a variety of biochemical pathways; there is an interaction between their assimilations, and the cellular energy balance acts as the coordinator of the two [12]. When NO₃⁻ and NH₄⁺ act as substrates for N growth, NO₃⁻ has a stronger affinity for carbon dioxide, and the addition of NO₃⁻ can change the electron distribution between different chloroplasts and mitochondrial pathways. The process of balancing the generation and utilization of energy in the survival mechanism of algae may be better coordinated by changing the N to achieve optimal selection [12]. On the other hand, this phenomenon may be determined by the evolutionary history of P. globosa. When NO₃⁻ is present in the medium, P. globosa promotes the formation of colonies, and few colonies are formed when NH₄⁺ is enriched alone [17,19,35,36]. Liang et al. (2021) reported that colonies have strong photosynthetic activity, which indicates that they have stronger carbon fixation functions and can produce more energy and organic matter [37]. Moreover, colonies also utilize N to secrete exopolysaccharide (EPS).

4.2. Regulation of Nitrogen-Metabolizing Enzymes

Nitrogen (N) metabolism is one of the basic physiological metabolic processes in plants and is completed by the catalysis of enzymes. Phytoplankton can reduce NO₃⁻ and NH₄⁺ to amino acids through the action of nitrogen-metabolizing enzymes and utilize these amino acids (Figure 9) [38]. Since the uptake and assimilation of NO₃⁻ require a two-step reductase process, NO₃⁻ requires much more energy than NH₄⁺, so most algae may preferentially use NH₄⁺ as N to conserve energy [17]. Algal cells can utilize different forms of nutrients by regulating the activities of GS, NR, and other enzymes. The presence of NH₄⁺ stimulates the GS gene and the activities of glutamine synthetase so that algal cells can assimilate and utilize NH₄⁺. NO₃⁻ is also an environmental signal that regulates many processes, such as gene expression [39]. Cells can sense and respond to environmental concentrations of NO₃⁻ through nitrate (NO₃⁻) transporters. For example, Arabidopsis thaliana acts as an ion sensor through the parental and receptor capacity of the nitrate (NO₃⁻) transporter AtNRT1.1 and a phosphorylation switch at H101, thereby mediating downstream gene expression through downstream kinase phosphorylation [40].

In fact, the transport and assimilation of NO₃⁻ are usually inhibited by NH₄⁺ [12], and the provision of NH₄⁺ can inhibit the uptake of NO₃⁻ by altering the activity of NR or preventing NR synthesis [41]. Moreover, NH₄⁺ can also affect NO₃⁻ metabolism by downregulating the transmembrane transport of NO₃⁻ [12]. Studies have shown that the presence of NH₄⁺ in media exerts an inhibitory effect on the expression of the NR gene and the activity of NR [42]. A similar phenomenon was observed in the present study. First, in group 2, both the GS and NR genes were expressed at 0 h, and the GS gene levels were downregulated at 2 h. In group 3, both genes were expressed at 4 h, and the NR gene expression was upregulated at 10 h (Figure 8). Furthermore, the presence of NH₄⁺ affects the activity of other enzymes, such as NR. In the three groups of experiments, NR was inhibited by NH₄⁺, and the activity of NR did not increase. In group 2, the concentration of NH₄⁺ decreased to 2.09 μmol·L⁻¹ after 2 h, and the activity of GS also showed a downward trend. At this time, NR was weakened by NH₄⁺ inhibition, and the related activity of NR increased. At 4 h in group 3, NO₃⁻ began to be assimilated and utilized. The activity of GS decreased at 4 h, and the activity of NR continued to increase after 4 h (Figure 7). When the glutamine level is low, the activity of NR is upregulated when NO₃⁻ is available. In addition, when the glutamine level is high, the activity of NR is inhibited [12]. The nutrient preference of phytoplankton for NO₃⁻or NH₄⁺ has been extensively studied in various environments, such as cultures or oceans, and the mechanism of preferential uptake is complex and regulated at multiple levels, including the uptake rate, gene expression control, and enzyme activity [42].

4.3. Mechanism of NH₄⁺ Production in P. globosa

The concentration of NH₄⁺ in fluid inside colonies in previous studies was significantly greater than that in the external environment. However, the underlying cause for this disparity remains uncertain [23]. We speculate that the difference in concentration may be related to the excess reduction of NO₃⁻ by algae and the release of excess NH₄⁺. To confirm this conclusion, NO₃⁻ with a high abundance of ¹⁵N was added to monitor the change in δ¹⁵N abundance in the NH₄⁺ pool to determine the transformation of ¹⁵N atoms from the NO₃⁻ pool to the NH₄⁺ pool. The results showed that the δ¹⁵N-NH₄⁺ value in all the experimental groups increased with time to varying degrees. For example, in group 3, group 2, and group 1, the δ¹⁵N-NH₄⁺ value increased to the highest value at 2 h, 4 h, and 6 h, respectively, with a correlation greater than 260‰, indicating a rapidly increasing ratio of ¹⁵N vs. ¹⁴N atoms (Figure 5). This variation may be attributed to the rapid removal of ¹⁴N and/or the rapid introduction of ¹⁵N. The former mainly refers to the assimilation and utilization of NH₄⁺ by P. globosa, which causes the ¹⁴N atom of the substrate to be utilized by P. globosa more quickly than the ¹⁵N atom, resulting in an increase in the δ¹⁵N-NH₄⁺ abundance of the substrate. Therefore, it is necessary to discuss whether the influence of the assimilation and utilization process is obvious. Taking the highest δ¹⁵N-NH₄⁺ value in each group as the reaction cutoff, the isotopic fractionation of assimilation and utilization was approximately 163~2094‰ (Figure 6), which was much greater than the normal level. It can be concluded that the increase in the abundance of the substrate δ¹⁵N-NH₄⁺ was not mainly caused by the assimilation of NH₄⁺ by P. globosa.

Notably, the increase in δ¹⁵N-NH₄⁺ abundance in each experimental group was consistent with the moment when the concentration of NO₃⁻ decreased, and the δ¹⁵N-NH₄⁺ value was positively correlated with the assimilation rate of nitrate. The isotopic fractionation of NH₄⁺ calculated for the high uptake rate of NO₃⁻ in group 1 reached 2094‰, and the isotopic fractionation of NH₄⁺ obtained for the relatively low uptake rate of NO₃⁻ in group 3 reached 163‰ (Figure 6). Therefore, it is concluded that the increase in δ¹⁵N-NH₄⁺ may be mainly due to the reduction of ¹⁵NO₃⁻, which introduces ¹⁵N atoms into the NH₄⁺ pool. The above transformation is thought to be driven by the dissipation/output of excess reducing agent energy under the action of cellular energy imbalance. The excess presence of reducing agents is usually caused by an imbalance between photochemistry and the assimilation of C and N, involving the lutein cycle, the Calvin cycle, etc. When photochemistry exceeds assimilation capacity, excess reducing agents may be produced [12]. Pressure causes the uptake rate of C and N to be slower than that of light, resulting in an energy imbalance. As a means of energy dissipation, algae are capable of taking up NO₃⁻ and subsequently releasing NH₄⁺ as a product [43]. When NO₃⁻ is reduced to NH₄⁺ by phytoplankton, substantial energy expenditure occurs. This reduction can be seen as a strategy to address excessive photosynthetic energy. Therefore, the combined action of reduction and release is an optimal method for dissipating that energy [44]. Kamp et al. (2011) used stable isotope labeling experiments to confirm that under anoxic or dark conditions, the utilization of ¹⁵NO₃⁻ was coupled with the generation and liberation of ¹⁵NH₄⁺ [45]. This study confirmed that the NH₄⁺ leakage process is synchronized with the uptake and assimilation of NO₃⁻ by P. globosa under light conditions.

Furthermore, there was a rapid decrease in δ¹⁵N-NH₄⁺ during the later stages of the experiment, raising speculation about the potential existence of dynamic equilibrium within the culture. Initially, it was observed in three experiments that the δ¹⁵N-NH₄⁺ value began to decrease after 6 h. Therefore, it was inferred that the decrease in δ¹⁵N-NH₄⁺ might be attributed to the weakened production of ¹⁵N or/and the introduction of ¹⁴N (Figure 10). Based on the observed changes in concentration, NO₃⁻ underwent consumption during this period, leading to a diminished generation of ¹⁵N. However, NO₃⁻ was not completely consumed after 6 h, especially in group 3. This result indicated that there might be an increase in mineralization in group 3 during this period, leading to an increase in ¹⁴N. Some evidence has suggested that the internal fluid of colonies is enriched in DON, which is vulnerable to degradation by heterotrophic bacteria [46]. Heterotrophic bacteria (such as Candidatus actinomarina minuta) have been observed to exist in the outer fluid and internal fluid of colonies and can act on organic nitrogen within algal cells, promoting mineralization and thereby generating new ¹⁴N [23]. Consequently, the weakened introduction of ¹⁵N and the production of new ¹⁴N may lead to a decrease in the δ¹⁵N-NH₄⁺ value.

5. Conclusions

Compared with other algae, P. globosa exhibited superior assimilation rates of NO₃⁻, reaching a maximum of 1.05 × 10⁻⁵ μmol·L⁻¹·h⁻¹·cell⁻¹. The inhibitory effect of ambient concentrations of NH₄⁺ on the assimilation of NO₃⁻ has also been observed in other algae. Similarly, this study revealed that the assimilation of nitrate is suppressed by NH₄⁺. However, when the ambient concentration of NH₄⁺ was below a certain threshold, the inhibitory effect diminished. Additionally, the presence of NH₄⁺ in the culture suppressed nitrate reductase activity and the expression of the NR gene. However, based on the δ¹⁵N-PON results, NO₃⁻ was still assimilated during the early stages of the experiment. Therefore, the assimilation of nitrate was incompletely suppressed by NH₄⁺. P. globosa releases NH₄⁺ into the environment during its growth phase, potentially utilizing this mechanism to release excess energy and address intracellular energy imbalances. The results accurately captured the process of NH₄⁺ release by P. globosa, contributing significantly to the understanding of the assimilation of inorganic nitrogen. Based on this study, further research can be conducted to investigate the nitrogen cycling processes related to NH₄⁺ released from P. globosa. It is hoped that this study can provide scientific data on the mechanism of algal blooms.