Light regulates inorganic nitrogen uptake and storage, but not nitrate assimilation, by the red macroalga Hemineura frondosa (Rhodophyta)

ABSTRACT

Macroalgal growth in temperate coastal ecosystems is primarily regulated by light and inorganic nitrogen availability. The effect of light (photon irradiance) on NO₃⁻ and NH₄⁺ uptake, and NO₃⁻ assimilation, were studied in the red macroalga, Hemineura frondosa, which does not operate a carbon concentrating mechanism (non-CCM). Non-CCM macroalgae grow in low and high light environments but become increasingly dominant with depth, suggesting a mechanism for ‘preserving energy’ under low light levels. H. frondosa was acclimated to limiting (30 µmol photons m⁻² s⁻¹) and saturating (150 µmol photons m⁻² s^–1) irradiances for 8 days. Then, NO₃⁻ and NH₄⁺ uptake rates were measured under limiting and saturating irradiances at six concentrations ranging from 2–64 μM. NO₃⁻ uptake did not follow saturating uptake kinetics at both irradiances suggesting multiple uptake mechanisms. NH₄⁺ uptake saturated at concentrations <32 µM under limiting but not under saturating irradiance. Saturating irradiance resulted in greater maximum uptake rates of both NO₃⁻ and NH₄⁺. There was no evidence that irradiance regulated NO₃⁻ reduction by nitrate reductase. Also illustrated is the importance of measuring nitrate reductase activity on fresh material, as freezing in liquid nitrogen and storage at −80°C for 7 days caused a 65% decline in activity. Photosynthetic pigments, soluble tissue nitrogen and % total tissue nitrogen were all higher in limiting irradiance. In this first study of the nitrogen physiology of a non-CCM seaweed, we show that light regulates NO₃⁻ and NH₄⁺ uptake but not NO₃⁻ assimilation.

KEYWORDS:

• Ammonium
• Hemineura frondosa
• irradiance
• nitrate
• nitrate reductase
• nitrogen uptake
• Rhodophyta

Introduction

Macroalgae are an intrinsic component of temperate coastal marine ecosystems and their growth is primarily regulated by the availability of light and nitrogen (Chapman & Craigie, 1977; Harrison & Hurd, 2001). Light regulates metabolic activities including photosynthesis and nutrient uptake (D’Elia & DeBoer, 1978; Udayakumar et al., 1981), while nitrogen is both an essential and limiting nutrient for growth (Phillips & Hurd, 2003; Hurd et al., 2014). The effects of light and nitrogen on macroalgal growth are tightly coupled and neither factor can facilitate primary production without the other (Lapointe et al., 1984; Turpin, 1991; Hurd et al., 2014). Both influence the biochemical composition of macroalgae (Figueroa et al., 1995, 1997; Marinho-Soriano, 2012), including light harvesting pigments – chlorophylls and red macroalgal accessory pigments (phycobilins) which are nitrogen rich – amino acids and inorganic soluble tissue nitrogen storage ‘pools’ (Figueroa et al., 1997; Barufi et al., 2011).

There are two main sources of dissolved inorganic nitrogen available for macroalgae: nitrate (NO₃⁻) and ammonium (NH₄⁺) (Smit, 2002; Ochoa-Izaguirre & Soto-Jiménez, 2015). NO₃⁻ is taken up across the plasma membrane via active transport with hydrogen ions (H⁺) (Syrett, 1981; Glass, 2003; Roleda & Hurd, 2019). In the cytoplasm it is reduced by the enzyme nitrate reductase (NR) to nitrite (NO₂⁻) which uses NAD(P)H as an electron donor (Marquardt et al., 2010). NO₂⁻ is subsequently reduced to NH₄⁺ by nitrite reductase (NiR) in the chloroplast before entering the glutamine synthetase-glutamine oxoglutarate aminotransferase (GS-GOGAT) pathway for assimilation into amino acids (Hipkin et al., 1983; Chow et al., 2007). In contrast, NH₄⁺ is typically taken up by passive transport and directly assimilated into amino acids via GS-GOGAT, so has a much-reduced energetic cost compared with NO₃⁻ uptake; thus many macroalgae have a strong preference for NH₄⁺ over NO₃⁻ as a nitrogen source (D’Elia & DeBoer, 1978; Raven et al., 1992; Nishihara et al., 2005; Abreu et al., 2011; Pritchard et al., 2015).

The mechanisms of NO₃⁻ and NH₄⁺ uptake can be elucidated, in part, by examining the kinetic pattern of uptake at a range of concentrations. Active transport is indicated by a hyperbolic curve in which uptake rate reaches a maximum rate at a critical concentration i.e. uptake is ‘saturated’ indicating involvement of a saturable transmembrane transporter (Harrison & Druehl, 1982). Passive diffusion is indicated by an uptake rate which increases linearly with external nutrient concentration (Hurd et al., 2014). A curve that shows evidence of saturating uptake kinetics at low concentrations and linear components at higher concentrations indicates that both active and passive uptake mechanisms are operating (Hurd et al., 2014); or that a combination of two or more active transport systems with high- and low-affinities for the nutrient are operating (Martínez & Rico, 2004). The preference and acquisition mechanism(s) for each dissolved inorganic nitrogen source are species dependent (Smit, 2002; Ochoa-Izaguirre & Soto-Jiménez, 2015).

Macroalgae can store NO₃⁻ in the cell vacuole and cytoplasm when there is a surplus of NO₃⁻ in the marine environment or insufficient light energy in the environment for NO₃⁻ assimilation (Davison & Stewart, 1984; Raven et al., 1992; Young et al., 2007). A large store of NO₃⁻ can inhibit further NO₃⁻ uptake and result in increased NR activity, with the rate of depletion of the NO₃⁻ pool being proportional to an increase in growth rate (Thomas & Harrison, 1985; Hwang et al., 1987; McGlathery et al., 1996; Lartigue & Sherman, 2005). When sufficient light energy is available, NO₃⁻ may be assimilated to NH₄⁺ which is subsequently assimilated into organic nitrogen such as pigments (chlorophyll, phycobilins), free amino acids (e.g. cirtulline, arginine) and proteins (Syrett, 1981; Harrison & Hurd, 2001). In red macroalgae, the synthesis of accessory pigments (phycobilins) has a relatively high requirement for nitrogen (Chopin et al., 1995). Their primary function is light harvesting, but they are sometimes considered nitrogen stores because when nitrogen supply is limiting, but light levels are optimal, the pigments are catabolized and used in growth (Lapointe, 1981; Marinho-Soriano, 2012). Under low light, phycobilin content can increase substantially to harvest more light energy but sufficient nitrogen supply is required for this process (Vergara & Niell, 1993; Barufi et al., 2011; Ak & Yücesan, 2012; Marinho-Soriano, 2012). In contrast to NO₃⁻, NH₄⁺ is not usually stored in large, unassimilated inorganic pools and is efficiently assimilated into amino acids, which in turn maintain the concentration gradient allowing NH₄⁺ to continue to diffuse into the cell when external NH₄⁺ is high (Fujita et al. 1988; Harrison & Druehl 1982; Raven et al. 1992).

NR activity is known to be up-regulated by the increased availability of NO₃⁻ in the surrounding medium (Davison & Stewart, 1984; Thomas & Harrison, 1987, 1988; Gordillo et al., 2006; Cabello-Pasini et al., 2011), and is also regulated by light levels (Corzo & Niell, 1992; Figueroa, 1996). Chow et al. (2004) found that NR activity in Gracilaria chilensis (Rhodophyta) increased when light pulses were given during dark periods, and suggested that NR was regulated mainly by light and not by a biological clock. Additionally, Chow et al. (2013) found a direct dependence of NR activity on PSII and PSI electron transport, but there is little information on the effect of light levels i.e. photon irradiance (µmol photons m⁻² s⁻¹) on the enzyme’s capacity to reduce NO₃⁻ in macroalgae. Saturating light can promote greater NO₃⁻ uptake rates compared with limiting light; this may be facilitated by an increase in NR enzyme activity due to the greater energy available (Wheeler, 1982; Hurd et al., 2014).

Tasmania has a globally unique macroalgal flora with an unusually high proportion (up to 90% of some sites) of macroalgae that solely use carbon dioxide (CO₂) as a source of dissolved inorganic carbon, and not energetically expensive bicarbonate; they do not operate a CO₂ concentrating mechanism (CCM) and are termed non-CCM seaweeds (Cornwall et al., 2015). These macroalgae can grow in both low and high light environments but become increasingly dominant with depth, suggesting a mechanism for ‘preserving energy’ at low light levels (Raven & Hurd, 2012; Raven et al., 2014; Cornwall et al., 2015). However, the nitrogen physiology of non-CCM species has not been examined to date, and we predict NH₄⁺ to be preferred as it requires little energy for uptake and assimilation and could thus be another mechanism for preserving energy under low lightlevels (Raven et al., 1992). Pritchard et al. (2015) found that low-light adapted Anotrichium crinitum had a strong preference for NH₄⁺, however for non-CCM red macroalgae it is not understood how uptake and assimilation of NO₃⁻ and NH₄⁺ is regulated by light. The objective of this study was to determine the effect of limiting and saturating irradiances on NO₃⁻ and NH₄⁺ uptake, soluble tissue NO₃⁻ and NH₄⁺ storage pools and NR activity of the non-CCM, red macroalga Hemineura frondosa (J.D.Hooker & Harvey) Harvey.

We hypothesized that for H. frondosa acclimated to limiting and saturating irradiance, (1) NO₃⁻ would exhibit saturating uptake kinetics indicative of active uptake, whilst (2) NH₄⁺ would exhibit linear uptake with increasing concentration indicative of passive diffusion. (3) Pre-experimental acclimation to saturating irradiance would promote greater NO₃⁻ uptake rates and this increase in uptake would be facilitated by higher NR activity compared with limiting irradiance and (4) acclimation to limiting irradiance would promote larger internal storage pools of NO₃⁻ compared with saturating irradiance.

Materials and methods

Macroalgae collection and experimental conditions

For the main experiment, 20 individuals of H. frondosa (30–40 cm² each) were collected by hand using SCUBA from Blackmans Bay (43°01′S, 147°33′E), south-eastern Tasmania, Australia, on 17 July 2017. Another 10 specimens were sampled for a subsequent NR experiment on 25 August 2017. Specimens were found growing in the subtidal on rock substratum at 2–6 m depth and were transported to the laboratory 20 min away in a cool box (25 litres, polypropylene/polyurethane). On this day, the irradiance at the collection depths, measured using a HOBO MX2202 (Mx temperature/light data logger), was between ~10–50 μmol photons m⁻² s⁻¹ at 10:00–11:00 AM. In the laboratory, all visible epiphytes were gently removed using tweezers and wiping with tissue paper. All experiments were conducted in a walk-in temperature-controlled room set at ambient temperature of collection, 12°C ± 0.4°C, with overhead irradiance provided by cool-white fluorescent tubes. A pulse-amplitude modulated (PAM) chlorophyll fluorescence meter (Walz Diving-PAM Underwater Fluorometer) was used to create rapid light curves (RLC), from which relative electron transport rate (rETR) were determined in sub-saturating and saturating irradiance of H. frondosa specimens. These irradiance levels were used as the ‘saturating irradiance’ and ‘limiting irradiance’ for experimental acclimation.

Seawater was available on tap from a 10 000 litre storage facility at the Institute for Marine and Antarctic Studies, University of Tasmania, and is collected bi-monthly from South Bruny Island, Tasmania (43°30′S, 147°13′E). Seawater was filtered to 1 μm and UV-sterilized prior to experiments (Emperor Aquatics Smart HO UV sterilizer, 025050-2, 50 W lamp, cartridge filter). All glassware was sterilized in 2% Decon 90^TM and 10% v/v hydrochloric acid solution (Merck 37%) prior to use. Seawater samples for nutrient analyses from the experiments outlined below were filtered using glass microfibre filter paper (0.7 µM, Whatman, GF/F), stored in 12 ml polyethylene nutrient tubes (LabServ®) at −20°C until analysis. Concentrations of N(NO₃⁻ + NO₂⁻) and NH₄⁺ in seawater were analysed using a QuickChem® 8000 Automated Ion Analyser (LaChat Instruments) following methods outlined in Diamond (2008) and Liao (2008).

Pre-experimental acclimation to limiting and saturating irradiance

Each of 20 individual algae were cut into ten 4 cm² pieces: pieces of each alga were placed into the same beaker with 500 ml of filtered seawater (0.7 uM, cartridge filter) and an air stone to maintain water motion. Ten of these beakers were placed under a limiting irradiance (30 µmol photons m⁻² s⁻¹) and the other 10 under a saturating irradiance (150 µmol photons m⁻² s⁻¹). The algae were acclimated for 8 days to daily seawater enrichment by adding 30 µM of NO₃⁻ as NaNO₃ (EMSURE® ACS,ISO,Reag. Ph Eur) and seawater was changed every 3 days.

Experimental design

After the 8 days’ acclimation, the 10 seaweed pieces within each of the 20 beakers were removed and used in experiments and assays as follows. Six pieces of algae from one beaker were used immediately in the NO₃⁻ and NH₄⁺ uptake experiments (see below). A seventh piece was used to determine pigment content and an eighth piece for NR activity: both these were immediately blotted dry, weighed (0.1 g ± 0.01) (± 0.01 mg, ME-T Analytical Balance), frozen with liquid nitrogen and stored at −80°C for 7 days. A ninth piece of alga was used to determine soluble tissue nitrogen and was immediately blotted dry, weighed (0.25 g ± 0.01), placed in a 50 ml boiling tube (borosilicate) with 20 ml of distilled water (ultrapure) and refrigerated overnight prior to boiling extraction. Finally, one piece of algae was randomly selected from 8 of the 20 flasks (i.e. four pieces of algae from each irradiance acclimation) for C:N analysis and were oven dried and weighed (5 mg ± 0.01) (see below).

NO₃⁻ and NH₄⁺ uptake kinetics

Multiple flask uptake experiments were run to determine the maximum uptake rate (V_max) and half saturation constant (K_m) of both NH₄⁺ and NO₃⁻ for H. frondosa which had been acclimated to either limiting or saturating irradiance (Harrison & Druehl, 1982; Hurd & Dring, 1990). Uptake experiments were conducted under the same irradiances as the 8 day acclimation period and consisted of the following treatments: 1. Limiting irradiance, NH₄⁺ uptake; 2. Saturating irradiance, NH₄⁺ uptake; 3. Limiting irradiance, NO₃⁻ uptake; 4. Saturating irradiance, NO₃⁻ uptake. There were five replicates for each of the four experimental treatments; each experimental replicate consisted of six seaweed pieces (see above) for which nutrient uptake was measured at each of six initial concentrations. Experiments were conducted in 250 ml conical flasks each containing 200 ml filtered seawater. Stock solutions of NO₃⁻ as NaNO₃ (EMSURE® ACS,ISO,Reag. Ph Eur) and NH₄⁺ as NH₄Cl (EMSURE® ACS,ISO,Reag. Ph Eur) were added to seawater in flasks to give six initial experimental concentrations of 2, 4, 8, 16, 32 and 64 µM in addition to the background concentrations. Four flasks with no algae acted as controls, two containing filtered seawater and two containing seawater plus 64 µM of both nitrogen sources.

At the start of the experiment, one piece of algae was added to each of 120 flasks (five replicate algae × six cut pieces for each alga × four experimental treatments). Conical flasks were then randomly positioned on shaker tables set at 100 rpm under limiting and saturating irradiances corresponding to the prior acclimation. Initial seawater samples (10 ml) were taken 2 h after the algal pieces were added to each flask and final samples were taken at 3 h: this time period was chosen because our preliminary time-course experiment revealed a 2 h lag phase in NO₃⁻ uptake. At the end of the experiment, algae were removed, and a final seawater sample taken. Algae were photographed for surface area which was calculated using image processing program ImageJ, and nitrogen uptake rates were standardized to surface area.

Soluble tissue NO₃⁻ and NH₄⁺ pools

Soluble tissue NO₃⁻ and NH₄⁺ content were analysed to determine unassimilated inorganic NO₃⁻ and NH₄⁺ within the alga cells using a boiling extraction method (Hurd et al., 1996). Boiling tubes (borosilicate, 50 ml) were filled with 20 ml of distilled water (ultrapure) and 0.25 ± 0.1 g pieces of alga tissue were added. Boiling tubes were removed after overnight refrigeration, capped in aluminium foil and placed in a boiling water bath (~100°C) for 40 min. Tubes were left to cool before the solution was filtered (Whatman, GF/F) and stored at −20°C. This process was repeated twice to ensure all soluble nitrogen was removed from the algal tissue. Each extract was analysed for concentrations of NO₃⁻ and NH₄^+. Results were standardized to wet weight using the following formula:

$\frac{\left({\mathrm{N}}_1+{\mathrm{N}}_2\right)\times 0.02}{\mathrm{W}\mathrm{W}}$

where N₁, N₂ are the NO₃⁻ or NH₄⁺ concentrations (µM) in the supernatant after the first and second extractions, 0.02 is the volume of liquid in each boiling tube (l) and WW is the wet weight of the algae (g).

Photosynthetic pigment content and C:N ratio

Phycoerythrin (PE) and chlorophyll contents were analysed to study the acclimation of H. frondosa to limiting and saturating irradiances. Pigment contents were determined using preserved algal material (0.1 ± 0.01 g) following methods outlined in Pritchard et al. (2013). Frozen tissue was ground using liquid nitrogen over ice in a mortar and pestle under dim light and phycobilins were extracted overnight in 15 ml centrifuge tubes (Greiner) using 5 ml of ice-cold 0.1 M phosphate buffer (pH 6.8, sodium phosphate dibasic and monobasic ≥ 99%, Sigma Life Sciences). The absorbance of the supernatant was measured at 564, 592, 455, 618 and 645 nm using a scanning spectrophotometer (Halo RB-10, Dynamica) and phycobilin contents were calculated using equations and coefficients by Beer & Eshel (1985). Remaining pellets were re-suspended in 5 ml of 100% ethanol (99.8%, Merck) in 15 ml centrifuge tubes, vortexed and chlorophyll was extracted overnight at 4°C. The absorbance of the supernatant was measured at 629, 649 and 665 nm before chlorophyll contents were calculated using equations from Ritchie (2006). Carbon and nitrogen content and C:N ratio were analysed to determine total nitrogen contents under saturating and limiting irradiances. Contents were analysed using a NA1500 elemental analyser coupled to a Thermo Scientific Delta V Plus via a Conflo IV. Tissue carbon and nitrogen content were expressed as a percentage of the alga’s dry weight and C:N ratios were indicated based on their atomic weights.

Nitrate reductase activity

During our preliminary experiments to optimize NR extraction, we achieved a NR activity of 36.55 (± 3.65) μmol NO₃⁻ reduced min⁻¹ g⁻¹ WW on fresh algal material. However, due to logistical limitations, for the experiment we froze samples in liquid nitrogen which were subsequently stored at −80°C for 7 days prior to measuring NR activity. The resulting activity was ~65% lower than that measured on fresh material. Therefore, to confirm our NR results (see later), we repeated the 1 week acclimation as described above between 27 August–4 September 2017, but this time measured the NR activity on fresh material immediately following the acclimation. In this second NR experiment, 10 individual algae were each cut into four pieces (each ~4 cm² as in the first experiment). Of these four pieces, two were acclimated under saturating and two under limiting irradiance. For the pieces acclimated under saturating irradiance, one was used for NR assay and the second for inorganic NO₃⁻ and NH₄⁺ pools to ensure nitrogen physiology of H. frondosa was comparable to the previous acclimation experiment. Similarly, under limiting irradiance, one piece of alga was used for NR assay and the second for inorganic NO₃⁻ and NH₄⁺ pools.

NR assays were run to analyse the rate of nitrate assimilation and followed the protocol by Pritchard (2013) who optimized the brown macroalgae methods of Hurd et al. (1996) and Young et al. (2005) for red macroalgae. Frozen algal pieces (0.1 g ± 0.01) were ground to a fine powder under liquid nitrogen in an ice-cold mortar and pestle. Algal powder was immediately transferred to a micro-centrifuge tube (1.5 ml, Eppendorf) and 1.5 ml of ice-cold NR buffer was added. NR buffer consisted of 200 mM phosphate extraction buffer (pH 7.9, sodium phosphate dibasic and monobasic ≥ 99%, Sigma Life Sciences), 5 mM EDTA (≥ 99.4%, chem-supply), 1% Triton^TM × 100 (v/v, Sigma Life Sciences), 0.3% polyvinylpyrrolidone (PVP, ≥ 99%, Fluka Analytical), 3% bovine serum albumin (BSA, Bovogen Biologicals) and 2 mM dithiothreitol (DTT, ≥ 97%, BIO-RAD). Solution was homogenized and left on ice for no longer than 30 min before assays began. NR assays were conducted in microcentrifuge tubes in a total volume of 1 ml with 200 µl of homogenate. Reactions were started with the addition of 100 μl 10 mM stock KNO₃ (≥ 98%, LabChem) solution and run for 40 min before being stopped using 500 µl of 50 mM zinc acetate (≥ 98%, Chem-supply) and suspension in a boiling water for 5 min. Homogenate was centrifuged (Centurion Scientific) at 13 000 g for 5 min and 1 ml of the supernatant was removed for colour reactions. Subsequent reactions indicated the presence of NO₂⁻ and included the addition of 20 µl 825 mM phenazine methosulphate (PMS, ≥ 99%, Sigma-Aldrich), 500 µl sulphanilamide (> 99%, Scharlau) and 500 µl N-(1-napthyl) ethylenediamine dihydrochloride (NEDD, > 98.0%, chem-supply). Concentrations of NO₂⁻ were determined spectrophotometrically at 540 nm and NR activity is reported as μmol NO₃⁻ reduced min⁻¹ gWW⁻¹. To account for any internal NO₂⁻ present in the algae a blank reaction was conducted by adding zinc acetate (< 98%, Chem-supply) prior to the addition of KNO₃ for each replicate.

Data analysis

Statistical analysis and curve fitting to nitrogen uptake data were conducted using R version 2.2 (R Development Core Team, 2012). A Shapiro–Wilk test was conducted to check the normality of the data and regressions of best fit were plotted using the graphing software package SigmaPlot (Systat Software Inc). Where possible, the Michaelis–Menten model was fitted to the uptake data to determine maximum uptake rate (V_max) and half saturation constant (K_m).

$\mathrm{V}={\mathrm{V}}_{\mathrm{m}\mathrm{a}\mathrm{x}}\frac{\mathrm{S}}{{\mathrm{K}}_{\mathrm{m}}+\mathrm{S}}$

where V is the uptake rate, V_max is the maximum uptake rate, S is the concentration of the limiting nutrient and K_m is the half saturation constant. The function was used as a single model for all the data due to variability, excluding outliers. In cases where the data did not follow a rectangular hyperbola, the regression of best fit was plotted. For our data, the regression best fit was plotted using the following equation:

$\mathrm{V}={\mathrm{V}}_{\mathrm{m}\mathrm{a}\mathrm{x}}\frac{\mathrm{S}}{{\mathrm{K}}_{\mathrm{m}}+\mathrm{S}}+\mathrm{C}\ast \mathrm{S}$

where V is the uptake rate, V_max is the maximum uptake rate, S is the concentration of the limiting nutrient, K_m is the half saturation constant and C is the linear trend. Again, the function was used as a single model for all the data due to variability, excluding outliers. To compare the uptake rates in the limiting- and saturating- irradiance acclimation, the relationship between uptake and concentration was determined by running an analysis of covariance (ANCOVA) on the following linear models:

Nitrate uptake rates:

$\begin{array}{rl}& \mathrm{u}\mathrm{p}\mathrm{d}\mathrm{a}\mathrm{t}\mathrm{e}=\alpha +{\beta }_1^{0.25}\left(\mathrm{p}\mathrm{o}\mathrm{l}\mathrm{y}\left(\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\right)\right)\\ & +{\beta }_2\left(\mathrm{t}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{t}\right)+{\epsilon }_{\mathrm{i},}\mathrm{d}\mathrm{a}\mathrm{t}\mathrm{a}=\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{e}\end{array}$

$\epsilon \sim \mathrm{N}\left(\mu ,{\upsilon }^2\right)$

Ammonium uptake rates:

$\begin{array}{rl}& \mathrm{u}\mathrm{p}\mathrm{t}\mathrm{a}\mathrm{k}\mathrm{e}=\alpha +{\beta }_1^{0.25}\left(\mathrm{l}\mathrm{o}\mathrm{g}\left(\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\right)\right)\\ & +{\beta }_2\left(\mathrm{t}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{t}\right)+{\epsilon }_{\mathrm{i},}\mathrm{d}\mathrm{a}\mathrm{t}\mathrm{a}=\mathrm{a}\mathrm{m}\mathrm{m}\mathrm{o}\mathrm{n}\mathrm{i}\mathrm{u}\mathrm{m}\end{array}$

$\epsilon \sim \mathrm{N}\left(\mu ,{\upsilon }^2\right)$

Where α is the intercept of the linear model, β are the uptake regression coefficients, treat is a categorical variable that represents irradiance and ε is the model residuals. ANCOVA test assumptions were assessed using diagnostic plots of model residuals and data were transformed where necessary using values of λ from Box–Cox plots. A t-test or Mann–Whitney U test (depending on normality of the data) was used to compare responses of soluble tissue, pigment content and NR enzyme activity for saturated and limited irradiance acclimated thalli.

Results

NO₃⁻ and NH₄⁺ uptake kinetics

For H. frondosa at both limiting and saturating irradiance, the regression of best fit for the uptake of NO₃⁻ was non-linear (saturating irradiance: adjusted R² = 0.77, limiting irradiance: adjusted R² = 0.53). At concentrations below 16 µM, the NO₃⁻ uptake rates appeared to saturate; however at higher concentrations, 16 µM and above, uptake rates further increased linearly (Fig. 1–2). NH₄⁺ uptake rates were best described by a Michaelis-Menten function (saturating irradiance: adjusted R² = 0.47, K_m = 152.93 µM and V_max = 0.12 µmol cm⁻¹ h⁻¹ and limiting irradiance: adjusted R² = 0.25, K_m = 16.17 µM and V_max = 0.01 µmol cm⁻¹ whereby uptake rates saturate with increasin g NH₄⁺ concentration (Fig. 3-4). The conversion factor between wet weight and surface area was 1 cm² = 10.67 g WW⁻¹.

Fig. 1–4 Uptake rates (µmol cm⁻² h⁻¹) for Heminura frondosa as a function of increasing NO₃^– concentration (µM) for algae acclimated to either (1) saturating irradiance (150 µmol photons m⁻² s⁻¹) and (2) limiting irradiance (30 µmol photons m⁻² s⁻¹) and NH₄⁺ under (3) saturating irradiance (150 µmol photons m^–2 s^–1) and (4) limiting irradiance (30 µmol photons m⁻² s⁻¹). Solid lines indicate regression of best fit. Note that the y-axis range for graphs 1 and 2 are different to those of 3 and 4. Each symbol represents an individual macroalgal replicate (n = 5)

Irradiance acclimation had a significant effect on the uptake rates of NO₃⁻ (two-way ANCOVA, p < 0.001), with higher uptake rates at all concentrations of NO₃⁻ under saturating irradiance than limiting irradiance (Fig. 1–2). Similarly, NH₄⁺, uptake rates were higher in thalli under saturating irradiance when compared with the limiting irradiance acclimation (two-way ANCOVA, p = 0.0208) (Fig. 3–4).

Soluble tissue NO₃⁻ and NH₄⁺ pools

Soluble tissue NO₃⁻ pools were two-times higher in thalli acclimated to limiting than saturating irradiance (Wilcoxon rank sum test, p < 0.05, Fig. 5), whereas there was no effect of irradiance on soluble tissue NH₄⁺ pools (Wilcoxon rank sum test, p = 0.90).

Fig. 5. Soluble tissue NO₃⁻ and NH₄⁺ concentrations (µmol g WW⁻¹) from Heminura frondosa individuals acclimated to saturating (150 µmol photons m⁻² s⁻¹) and limiting (30 µmol photons m⁻² s⁻¹) irradiance. Bars represent mean ± SE, n =10, * = p < 0.05 (Wilcoxon rank sum test)

Photosynthetic pigment contents

Hemineura frondosa acclimated in a limiting irradiance had 30% more PE than those acclimated under saturating irradiance (Wilcoxon rank sum test, p < 0.05; Fig. 6). Similarly, Chl a content was 35% higher under limiting irradiance compared with saturating irradiance (Wilcoxon rank sum test, p < 0.05; Fig. 7).

Fig. 6–7. Photosynthetic pigment content of Heminura frondosa (mg g WW⁻¹) acclimated to saturating (150 µmol photons m⁻² s⁻¹) and limiting (30 µmol photons m⁻² s⁻¹) irradiance (6) PE and (7) Chl a. Bars represent mean ± SE, n =10, * = p < 0.05 (Wilcoxon rank sum test)

Nitrate reductase assay

NR activity was similar (average of 27.2 ± 1.46 μmol NO₃⁻ reduced min⁻¹) for individuals acclimated to saturating and limiting irradiance (two tailed t-test, p = 0.55; Fig. 8).

Fig. 8. NR activity (μmol NO₃⁻ reduced min^–1 g WW^–1) from Heminura frondosa acclimated under saturating (150 µmol photons m⁻² s⁻¹) and limiting (30 µmol photons m⁻² s⁻¹) irradiance. Bars represent mean ± SE, n = 10

Total tissue C, N and C:N ratio

Total tissue carbon (%) was not significantly affected by irradiance acclimation (two tailed t-test, p = 0.84; Fig. 9) whereas total tissue nitrogen (%) was 20% higher in thalli acclimated to limiting than saturating irradiance (two tailed t-test, p < 0.01; Fig. 10). As a result, the C:N ratio was 25% lower for the limiting compared with saturating irradiance acclimation (two-tailed t-test, p < 0.05; Fig. 11).

Fig. 9–11. Tissue carbon (9) and nitrogen content (10) expressed as %DW, and C:N ratio (11), for Heminura frondosa acclimated to saturating (150 µmol photons m⁻² s⁻¹) and limiting (30 µmol photons m⁻² s⁻¹) irradiance. Bars represent mean ± SE, n = 4, * = p < 0.05 (Wilcoxon rank sum test)

Discussion

The patterns of NO₃⁻ and NH₄⁺ uptake in H. frondosa were influenced by prior irradiance acclimation indicating that light plays a significant role in inorganic nitrogen uptake and utilization. The first hypothesis, that NO₃⁻ would exhibit saturating uptake kinetics, indicative of active uptake, was partly supported: uptake rates under both saturating and limiting irradiance appeared to saturate between 2 and 16 µM, but was linear at higher concentrations. This uptake pattern is similar to NO₃⁻ uptake observed in Palmaria palmata (Rhodophyta; Martínez & Rico, 2004), Gracilaria pacifica (Rhodophyta; Thomas et al., 1987), Ulva rigida (Chlorophyta; Lavery & McComb, 1991), Laminaria saccharina and Laminaria digitata (Phaeophyceae; Conolly & Drew, 1985) and has been explained by active uptake mechanism(s) at lower concentrations and passive diffusion at higher concentrations (see fig. 6.2d in Hurd et al., 2014). However NO₃⁻ uptake is typically via active transport rather than diffusive uptake (Hurd et al., 2014). NO₃⁻ transporters in algae and higher plants are encoded by multigene families which are differentially affected by external concentrations of NO₃⁻ (Runcie et al., 2003; Kakinuma et al., 2008). In terrestrial plants, high affinity transport systems are indicated by low K_m values and saturable uptake rates, whilst low affinity systems have a relatively high K_m, and at high nutrient concentrations they have a linear relationship with uptake rates (Glass, 2003). This pattern is similar to that observed for H. frondosa, and it is most likely that both high- and low-affinity NO₃⁻ transport systems are used for NO₃⁻ uptake, rather than a diffusive component. This would be advantageous under fluctuating ambient NO₃⁻ concentrations, dependent on the requirements of the algal tissue (Galván & Fernández, 2001). Our work highlights that for macroalgae, further research to identify the underpinning cellular and molecular regulatory mechanisms of NO₃⁻ uptake is needed.

The second hypothesis that NH₄⁺ uptake would increase linearly with concentration, indicating uptake by passive diffusion, was partly supported at saturating, but not limiting, irradiance. Although NH₄⁺ uptake rates under saturating irradiance appear to increase with NH₄⁺ concentration, a rectangular hyperbola was the best fit curve suggesting that had we used higher concentrations, uptake rates would have saturated. This is reflected in V_max which was very high (0.12 µmol cm⁻¹ h⁻¹), and the K_m of 152.93 µM, which suggests saturation would occur at ~ 300 µM. Under limiting irradiance, NH₄⁺ uptake saturated at 32.34 µM with a K_m of 16.17 µM which was within the range of other marine Rhodophytes (range = 1.60–76.45 µM) although the V_max of 0.01 µmol cm⁻¹ h⁻¹ was more than 10× lower than other subtidal Rhodophytes: Gracilaria foliifera, Neoagardhiella baileyi (D’Elia & DeBoer, 1978), Anotrichium crinitum (Pritchard et al., 2015) and Palmaria palmata (Martínez & Rico, 2004). We hypothesized that NH₄⁺ uptake would exhibit non-saturating kinetics indicative of passive diffusion since H. frondosa has physiological characteristics of a species adapted to low-light environments (Raven et al., 1992; Cornwall et al., 2015). However, saturable uptake kinetics usually indicate active or mediated uptake mechanisms which are energy dependent (Harrison & Druehl, 1982; Harrison & Hurd, 2001) and assimilation rate may have exerted control over the pattern of uptake rate with concentration (Rees et al., 1998). Either one or both of these mechanisms are likely employed by H. frondosa as saturating irradiance promoted increased rates of NH₄⁺ uptake and suggests an irradiance-regulated mechanism, however without mechanistic study using inhibitors, it is impossible to determine which uptake mechanim is used (Roleda & Hurd, 2019). To our knowledge, ours is the only study to examine the effect of light on NH₄⁺ uptake rates by a non-CCM macroalga.

We hypothesized that acclimation to saturating irradiance would promote greater uptake of NO₃⁻ and that this would be facilitated by an increase in NR activity. This hypothesis was partially supported; NO₃⁻ uptake was greater under saturating irradiance, but NR activity was unaffected, suggesting that NO₃⁻ uptake and NR activity are uncoupled for this species. Furthermore, acclimation to limiting irradiance resulted in larger internal stores of NO₃⁻, compared with saturating irradiance. This supported our hypothesis and may suggest that the differences in NO₃⁻ uptake rates of H. frondosa under limiting and saturating irradiances are instead facilitated by the preferential reduction of inorganic nitrogen pools under limiting irradiance before NO₃⁻ is actively taken out of the external medium. This method of nitrogen ‘recycling’ has been described for seagrass and was linked to a reduction of the plant’s dependence on external nitrogen sources (Borum et al., 1989). The preferential reduction of stored inorganic nitrogen contributed 40–69% of total nitrogen gained by Posidonia oceanica and 60% of that gained by Zostera marina (Borum et al., 1989; Alcoverro et al., 2000). To our knowledge, this reduced dependence on external nitrogen sources due to recycling of internal stores has not previously been described for macroalgae and clearly requires further work to test if it is the case.

Storage of unassimilated NH₄⁺ in H. frondosa was unaffected by irradiance but content was high (5.36 μmol g WW⁻¹ ± 0.93 SD, saturating irradiance, tissue nitrogen 2.8%) in comparison to other subtidal Rhodophytes, Gracilaria tikvahiae (3.80 μmol g WW⁻¹, tissue nitrogen 2.0%; Bird et al., 1982) and Anotrichium crinitum (1.8 μmol g WW⁻¹ ± 0.61 SD; Pritchard et al., 2015). The unassimilated stores of NH₄⁺ may have decreased NH₄⁺ uptake by H. frondosa as evidenced by a saturation of the uptake rates under limiting irradiance (McGlathery et al., 1996). High internal concentrations of unassimilated NO₃⁻ and NH₄⁺ limit the uptake capacity of macroalgae and may have resulted in the relatively low uptake rates observed in this study for both NO₃⁻ and NH₄⁺, as nutrient-limited algae generally show higher nutrient uptake rates than nutrient-replete algae (Kregting et al., 2008).

Phycobilins are the main light-harvesting accessory pigments of the Rhodophyta, and for some species can be a source of organic nitrogen which can be catabolized during periods of low nitrogen concentrations in ambient seawater (Martínez & Rico, 2002; Kim et al., 2007; Barufi et al., 2011). Phycobilins may be a major nitrogen store, but under these experimental conditions there is no evidence that the stores were used as the algae were nitrogen sufficient during the acclimation period. The algae responded to the limiting irradiance by increasing both PE and Chl a content, which would enhance the light harvesting capacity i.e. shade adaptation. Phycocyanin content was negligible (0.015 ± 0.005 mg g WW⁻¹) indicating that it is not an important phycobilin for H. frondosa. PE and Chl a contents were similar to other Rhodophyta studies (Sampath-Wiley et al., 2008; Ribeiro et al., 2013; Wu, 2016).

An important finding of this study was that NR activity was strongly, negatively affected by freezing samples in liquid nitrogen and storing at −80°C for 2 weeks. Although we achieved the same result, i.e. no effect of light on NR activity, the activity in fresh material was ~65% greater than that of frozen material. This technique of freezing biological material for subsequent enzyme assays is common practice (Geiger et al., 1998; Chow et al., 2004, 2007; Chow & De Oliveira, 2008; Pritchard, 2013; Fernández et al., 2015), but our study illustrates the importance of testing the effect of freezing and storage prior to experiments.

We have shown that irradiance plays a significant role in both NO₃⁻ and NH₄⁺ uptake, as well as utilization for H. frondosa. Genomic characterization of NO₃⁻ and NH₄⁺ transporters and their regulation by external factors would enhance our knowledge of inorganic nitrogen uptake in macroalgae in the future. The majority of studies on nitrogen uptake kinetics of red macroalgae are for species of commercial interest e.g. Porphyra/Pyropia spp. (Thomas & Harrison, 1985; Kraemer et al., 2004; Kang et al., 2014), Gracilaria spp. (Bird et al., 1982; Lapointe et al., 1984; Thomas et al., 1987; Naldi & Wheeler, 1999, 2002; Smit, 2002; Barufi et al., 2011), Palmaria palmata (Martínez & Rico, 2004; Corey et al., 2013; Grote, 2016) and Chondrus crispus (Amat & Braud, 1990; Corey et al., 2013). The limited knowledge of nitrogen uptake kinetics for non-commercial red macroalgae growing in natural systems highlights a key gap in the current phycological literature, along with the need for an improved understanding of the biochemical and molecular processes regulating inorganic nitrogen uptake and assimilation.

Author contributions

E. Paine: experimental design, conducted experiments and data analyses, drafting and editing of manuscript; M. Schmid: experimental design, laboratory assistance, editing manuscript; A. Revill: analysis of tissue carbon and nitrogen content; C. Hurd: original concept, experimental design, editing manuscript

Acknowledgements

We wish to thank Damon Britton, Dr Fanny Noisette, Harrison Vermont, Abbie Smith, Olivia Johnson and Joanna Smart for their help in the laboratory and field, Drs Erica Young and John Berges for critical feedback on the manuscript and Dr Daniel Pritchard for assistance with nitrate reductase assays.

Disclosure statement

No potential conflict of interest was reported by the authors.

Additional information

Funding

Matthias Schmid was funded through the Deutsche Forschungsgemeinschaft (DFG, grant ID: SCHM 3335/1-1).