Effects of temperature and nitrate on phosphomonoesterase activities between carbon source and sink tissues in Zostera marina L.

Journal of Experimental Marine Biology and Ecology 342 (2007) 313 – 324 www.elsevier.com/locate/jembe

Effects of temperature and nitrate on phosphomonoesterase activities between carbon source and sink tissues in Zostera marina L. Brant W. Touchette a,⁎, JoAnn M. Burkholder b b

a Center for Environmental Studies and Department of Biology, Campus Box 2625, Elon University, Elon, NC 27244, USA Center for Applied Aquatic Ecology, North Carolina State University, 620 Hutton Street, Suite 104, Raleigh, NC 27606, USA

Received 6 September 2006; received in revised form 9 October 2006; accepted 10 November 2006

Abstract Inorganic phosphorus (Pi) is important in the regulation of many carbon and nitrogen metabolic processes of plants. In this study, we examined alterations of phosphomonoesterase activity (PA; both alkaline and acid) in a submersed marine angiosperm, Zostera marina, grown in Pi non-limiting conditions under elevated temperature and/or nitrate enrichment. Control plants (ambient water-column NO−3 b 2.5 μM, with weekly mean water temperatures between 26.5–27.0 °C based on a 20-yr data set in a local embayment) were compared to treated plants that were exposed to increased water-column nitrate (8 μM NO−3 above ambient, pulsed daily at 0900 h), and/or increased temperature (ca. 3 °C above weekly means) over eight weeks in late summer–fall. Under both nitrate regimes, increased temperature resulted in periodic increased leaf and root-rhizome tissue carbon content, and increased acid and alkaline PA activities (AcPAs and AlPAs, respectively). There was a positive correlation between AlPA and AcPA activities and sucrose synthase activities in belowground structures, and a negative correlation between AlPA activities and sucrose concentrations. There were also periodic changes in PA partitioning between carbon source and sink tissues. In hightemperature and high-nitrate treatments, AcPAs significantly increased in leaves relative to activities in root-rhizome tissues (up to 12-fold higher in aboveground than belowground tissues in as little as 3 weeks after initiation of treatments). These responses were not observed in control plants, which maintained comparable AcPA activities in above- and belowground tissues. In addition, AlPA activity was significantly higher in leaf than in root-rhizome tissues of plants in high-temperature (weeks 3 and 6) and high temperature combined with high nitrate treatments (week 8), relative to AlPA activities in control plants. The observed changes in PAs were not related to Pi growth limitation, and may allow Z. marina to alter its carbon metabolism during periods of increased carbon demand/mobilization. This response would make it possible for Z. marina to meet short-term P requirements to maximize carbon production/allocation. Such a mechanism could help to explain the variability in PA activities that has been observed for many plant species during periods when environmental Pi exceeds requirements for optimal growth. © 2006 Elsevier B.V. All rights reserved. Keywords: Carbon metabolism; Nitrate enrichment; Phosphatase; Sucrose synthase; Seagrass; Temperature; Zostera marina

1. Introduction Phosphorus acquisition is important for plant growth, development, and survival. Phosphomonoesterases (PAs, ⁎ Corresponding author. Tel.: +1 336 278 6185; fax: +1 336 278 6258. E-mail address: [email protected] (B.W. Touchette). 0022-0981/$ - see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.jembe.2006.11.005

also referred to as phosphomonoester-hydrolases or phosphatases) are involved in P acquisition as catalysts for the release of orthophosphate (Pi) from organic compounds, especially during periods of increased P demand (McLachlan, 1980b; Barrett et al., 1998). These enzymes tend to be substrate non-specific, allowing the release of Pi from a broad range of P-containing

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compounds. The ecological role of PAs, especially alkaline phosphomonoesterases (AlPA; EC 3.1.3.1), has been linked to P deprivation in algae and higher plants (McLachlan, 1980b; Burkholder and Wetzel, 1990; Perez and Romero, 1993; Lapointe et al., 1994). As Pi becomes less available, plants respond by enhancing PA activity, thereby increasing the potential for P utilization from environmentally derived organic compounds, and/or recycling P from internal organic compounds (Vincent and Crowder, 1995; Dore and Priscu, 2001). Typically for aquatic plants under Pi limitation, PAs may be released extracellularly into the localized rhizosphere and the surrounding environment (Healey and Hendzel, 1979; Smith and Kalff, 1981; Lapointe et al., 1994), and/or may accumulate internally in both apoplastic and symplastic tissues (Jansson et al., 1988; Thaker et al., 1996). Phosphomonoesterases are also involved in metabolic processes including hydrolysis, transport of materials across membranes, synthesis of P-containing compounds via transphosphorylation, formation of the primary cell wall, wall lignification, and polysaccharide synthesis (McComb et al., 1979; Georgieva, 1980; Jeanmaire et al., 1985; Jansson et al., 1988; Thaker et al., 1996). Acid PAs (AcPAs; EC 3.1.3.2) have sometimes been described as less responsive to available environmental Pi than alkaline PAs (AlPAs; Wynne, 1977; Jansson et al., 1988). Therefore, it has been suggested that AcPAs are continuously expressed and may be involved in internal P metabolism, whereas AlPAs, with greater external function, are more responsive to environmental Pi levels (Vincent and Crowder, 1995; Invers et al., 1995). The primary role of P in plant metabolism has been well documented. It is a major constituent in nucleic acids and phospholipids; it is also involved in energy transfer (e.g. ATP and NADPH), intermediates of metabolic pathways (e.g. glucose-6-phosphate), and regulation of enzyme activity through protein phosphorylation, including enzymes involved in carbon and nitrogen metabolism (Marschner, 1995). P i availability also has been shown to affect photosynthetic rates. Under low Pi, photosynthesis can decrease by as much as 50% (Dietz and Foyer, 1986; Rao et al., 1989; Usuda and Shimogawara, 1991). This decrease may result from lower activation of ribulose-1,5bisphosphate carboxylase-oxygenase (Rubisco) and/or decreased regeneration of ribulose-1,5-bisphosphate (Brooks, 1986; Rao et al., 1989). Lower levels of internal Pi can develop following accumulation of phosphorylated intermediates of starch and sucrose synthesis/degradation (e.g. glucose-6-phosphate and

fructose-1,6-bisphosphate; Sharkey, 1985; Rao et al., 1989). Low ATP levels, which can develop under low Pi conditions, can limit the active uptake of nitrate through plant membranes. This limitation may occur by restricting the influx of nitrate ions against an electrical potential gradient, or by limiting synthesis of the membrane transport system specific for nitrate (Rufty et al., 1993). Nitrate reduction also can be controlled by available Pi through allosteric regulation via protein phosphorylation of nitrate reductase (NR) — as NR becomes phosphorylated via a NR kinase, the rate of nitrate reduction is substantially decreased (Huber et al., 1992). Phosphate translocators facilitate the exchange of Pi from the cytosol and triose phosphates (triose-P; e.g. glyceraldehyde-3-phosphate and dihydroxyacetone phosphate) from the chloroplast (Marschner, 1995). The triose-P that enters into the cytosol can then be utilized for the formation of more complex carbohydrates. Therefore, cytosolic Pi is necessary for the release of triose-P from the chloroplast. When Pi becomes relatively low in the cytosol, triose-P can accumulate in the chloroplast, enhancing starch production (Marschner, 1995). Because of the importance of Pi in plant metabolism, during times of increased metabolic Pi demand (e.g. periods of increased carbon demand), plants may need to adjust their Pi sequestering efficiency and/or internal P recycling, even when Pi is not intrinsically growth-limiting (Touchette and Burkholder, 1998). The purpose of this study was to test the hypothesis that short-term physiological P deficiency, indicated by elevated PA activity, can develop during periods of increased N and C metabolism in the dominant north temperate seagrass species, Zostera marina L. (eelgrass). We examined the physiological Pi demands in Z. marina under changing conditions of carbon allocation/partitioning, achieved by increasing nitrate uptake (to impose a carbon “drain” from nitrate reduction and carbon skeletons required for amino acid synthesis), increasing temperature (thereby increasing respiration as well as enhancing carbohydrate accumulation; Biebl and McRoy, 1971; Touchette and Burkholder, 2001), or increasing nitrate and temperature in combination. The submersed seagrass species selected for study can become N- or C-limited under some conditions, including internal C-limitation during periods of inorganic N assimilation and reduction (Borum et al., 1989; Burkholder et al., 1992; van Lent et al., 1995; Zimmerman et al., 1997; Touchette et al., 2003), making it suitable for examination of metabolic Pi demands unrelated to direct Pi limitation.

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2. Materials and methods 2.1. Study area The research was conducted in North Carolina, U.S.A., which is the southernmost extension for Z. marina along the western Atlantic Coast (Thayer et al., 1984). In this warm temperate area, growth of Z. marina is stunted apparently by high temperature stress (Den Hartog, 1970). Temperatures above 30 °C are common in shallow water embayments during late spring to early autumn (National Oceanic and Atmospheric Administration [NOAA], 1994; Motes et al., 1998). In more northern geographic regions, temperatures above 30 °C have been shown to adversely affect the physiology of this species by increasing respiration and impairing enzyme activities (Lambers, 1985; Marsh et al., 1986; Zimmerman et al., 1989). 2.2. Experimental system Transplanted Z. marina shoots were grown in the experimental mesocosm system described in Burkholder et al. (1992). Briefly, the system consisted of 12 fiberglass mesocosms (2.0 m diameter by 1.0 m height, with a 60 cm water column depth). The mesocosms had a raised base to accommodate the small plants (mean length 40 cm) while minimizing wall shading effects. The substratum (thickness ∼ 30 cm) consisted of natural marine sediments and sand mixed in a 3:1 ratio by volume. Prior to transplanting, the mesocosms were allowed to acclimate in running seawater for four months to establish natural chemical and biological gradients in the water column and sediment. Each mesocosm was individually plumbed with circulating seawater on both intake and outflow lines, so that there was no treatment cross-contamination. The continuous water circulation prevented anoxic conditions from establishing in the water column during dark periods. Temperatures were maintained using a chiller system that consisted of three 4.5 t condensing units, each capable of a maximum of 60000 BTU h- 1 cooling capacity. The chiller units had electronically controlled thermal expansion valves that regulated freon coolant to each titanium heat exchanger, which provided independent temperature control for each tank to within + 0.5 °C. Light reduction associated with tidal fluctuations was simulated by using neutral density shades that decreased photosynthetically active radiation (PAR) by ca. 30%. To simulate changing light over tidal cycles, shades were placed over each tank for 3 h on a rotating schedule (0900–1200 h for 3 d followed by 1200–1500 h for 3 d,

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then 3 d without shading). Under shaded conditions, Z. marina plants still were above reported light saturation for photosynthesis in field populations (N350 μmol m− 2 s− 1; Dennison and Alberte, 1985; Dennison, 1987). 2.3. Experimental design Plants were collected in mid-April from natural populations in an oliglotrophic estuarine system, Middle Marsh near Beaufort, NC (Coleman and Burkholder, 1995). They were cleaned of epiphytes and sediments, transplanted into each mesocosm at an initial density of 650–700 shoots m− 2, and allowed to acclimate for 4 months. The experiment was conducted over a twomonth period during late summer–fall (26 August–21 October). The experiment followed a split-plot design, with condensing units defined as the plot variant (4 mesocosms per condensing unit). Control mesocosms (n = 3) were maintained at ambient nitrate + nitrite (hereafter referred to a nitrate, b2.5 μM) and mean weekly temperatures, based on 20-yr weekly means (26.5–27.0 °C; NOAA, 1994). The experimental treatments consisted of increased nitrate (HN; n = 3), increased temperature (HT; n = 3), and a combined condition of increased nitrate and temperature (HTN; n = 3). Nitrate-enriched treatments were imposed by increasing the ambient water-column nitrate concentrations by 8.0 μM NO3−. Nitrate was pulsed daily as sodium nitrate at 0900 h. Temperature treatments were imposed by increasing mean weekly environmental temperature by 3–4 °C. All tanks were flushed with seawater between 1700 and 1800 h at a volume exchange rate of 10% d− 1 to simulate the low natural flushing characteristic of many shallow lagoons. 2.4. Environmental analyses Minimum and maximum water temperatures were tracked daily in each mesocosm. Salinity, dissolved oxygen (DO) and pH were monitored weekly using a Hydrolab H2O multiprobe and transmitter with SVR3-DL datalogger (Hydrolab Inc., Austin, TX, USA). Photosynthetically active radiation (PAR) was measured at the water surface and at the canopy level using a Li-Cor LI1000 data logger with a 4π light sensor (Li-Cor Biosciences, Lincoln, NE, USA). Water-column nutrients were collected in duplicate during weeks 3, 4, 6, and 8, except for ammonium which was assayed during weeks 3, 4 and 8. Ammonium (NH4+) was analyzed by the Solórzano procedure (Parsons et al., 1985; practical quantitation limit 0.67 μM NH4+), modified for automation

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Table 1 Physiochemical variables including temperature (Temp., °C), pH, salinity, dissolved oxygen (DO, mg L− 1), photosynthetically active irradiance (PAR, μE m− 2 s− 1), ammonium (NH+4 , μM), nitrate (NO−3 , μM), and total phosphorus (TP, μM) PAR

NH+4

NO−3

TP

6.6 ± 0.4 9.6 ± 0.2 6.0 ± 0.2 6.3 ± 0.4

1770 ± 30 2000 ± 50 1770 ± 60 1670 ± 40

1.8 ± 0.08 3.6 ± 0.36 ND 0.6 ± 0.17

1.1 ± 0.15 2.1 ± 0.39 2.1 ± 0.04 0.7 ± 0.42

0.32 ± 0.07 0.43 ± 0.09 0.42 ± 0.07 0.38 ± 0.08

32.1 ± 0.4 34.5 ± 0.4 36.5 ± 0.3 36.8 ± 0.2

6.0 ± 0.2 8.9 ± 0.1 5.7 ± 0.1 5.0 ± 0.6

1790 ± 20 2030 ± 95 1660 ± 95 1660 ± 70

2.6 ± 0.79 3.7 ± 0.31 ND 0.6 ± 0.06

2.1 ± 0.20 1.3 ± 0.64 1.9 ± 0.39 1.4 ± 0.23

0.37 ± 0.07 0.26 ± 0.01 0.29 ± 0.01 0.37 ± 0.09

8.1 ± 0.08 8.1 ± 0.03 8.1 ± 0.03 8.2 ± 0.04

31.3 ± 0.3 32.7 ± 0.3 35.5 ± 0.1 34.9 ± 0.1

7.0 ± 0.5 9.2 ± 0.2 6.0 ± 0.2 6.5 ± 0.1

1730 ± 15 2050 ± 20 1570 ± 55 1750 ± 10

1.6 ± 0.31 5.4 ± 1.36 ND 1.1 ± 0.21

1.0 ± 0.16 0.7 ± 0.44 2.4 ± 0.24 1.6 ± 0.31

0.26 ± 0.01 0.40 ± 0.04 0.39 ± 0.01 0.38 ± 0.04

8.2 ± 0.13 8.3 ± 0.20 8.2 ± 0.19 8.2 ± 0.20

32.3 ± 0.1 34.3 ± 0.2 36.1 ± 0.6 36.4 ± 0.1

7.5 ± 1.0 10.2 ± 0.9 6.7 ± 0.6 6.6 ± 1.0

1750 ± 20 2010 ± 10 1560 ± 80 1690 ± 30

1.4 ± 0.05 3.1 ± 0.34 ND 1.0 ± 0.22

1.4 ± 0.59 1.7 ± 0.47 2.0 ± 0.55 1.1 ± 0.19

0.26 ± 0.01 0.29 ± 0.04 0.25 ± 0.03 0.32 ± 0.01

Treatment (week)

Temp.

pH

Salinity

Control Week-3 Week-4 Week-6 Week-8

26.6 ± 0.1 26.5 ± 0.1 26.4 ± 0.2 26.4 ± 0.1

8.1 ± 0.10 8.2 ± 0.11 8.0 ± 0.06 8.1 ± 0.05

31.4 ± 0.2 32.6 ± 0.1 35.4 ± 0.2 34.5 ± 0.2

HT Week-3 Week-4 Week-6 Week-8

29.5 ± 0.5 29.2 ± 0.5 31.8 ± 0.2 29.2 ± 0.2

8.0 ± 0.06 8.1 ± 0.03 8.0 ± 0.08 7.8 ± 0.12

HN Week-3 Week-4 Week-6 Week-8

26.4 ± 0.2 26.6 ± 0.1 26.8 ± 0.2 26.6 ± 0.1

HTN Week-3 Week-4 Week-6 Week-8

29.2 ± 0.5 29.4 ± 0.1 31.4 ± 0.2 29.4 ± 0.1

DO

Temperature, pH, salinity, and DO were obtained between 1100 and 1130 h, PAR was recorded between 1200 and 1300 h, and nutrients were collected between 0800 and 0900 h (approximately 1 h prior to nitrate additions on the collection date). Data are given as means ± 1 SE. Note that week-6 NH+4 was not determined (ND).

using a Technicon Tracks 800 autoanalyzer (Technicon Instruments, Chicago, IL, USA). Water samples for total P (TP) analysis were frozen at −20 °C until analysis, using a variance of EPA method 365.1 (U.S. EPA, 1992, 1993; practical quantitation limit [American Public Health Association et al., 1992] 0.11 μM PO4− 3). Samples for nitrate analysis were frozen and analyzed within two months, using a variance of EPA method 353.4 (U.S. EPA, 1992; practical quantitation limit 0.10 μM NO3−). 2.5. Plant tissue analyses Above- and belowground tissues of Z. marina were thoroughly washed with seawater prior to enzyme and tissue analyses to remove epiphytic and microrhizal flora that could potentially have confounded measurements. Previous studies have shown negligible microbial interference of PA activity by sparse epiphytic biofilms, relative to bulk tissue activities (Ridge and Rovira, 1971; McLachlan, 1980a; Kroehler and Linkins, 1988). All tissue samples used for enzyme analysis were collected between 1500 and 1700 h to allow time for the treated plants to respond to increased water-column nitrate conditions prior to water exchange. Five enzymes

were assayed, including enzymes involved in the metabolism of carbon (sucrose-P synthase [SPS; EC 2.4.1.14] in leaf tissue, and sucrose synthase [SS; EC 2.4.1.13] in the root-rhizome complex), nitrogen (glutamine synthetase [GS; EC 6.3.1.2] in both shoot and root-rhizome complex tissues), and phosphorous (acid phosphatase [AcPA; EC 3.1.3.2] and alkaline phosphatase [AlPA; EC 3.1.3.1], in both shoot and rootrhizome complex tissues). Glutamine synthetase (GS) activity was assayed using the in vitro technique of Weissman (1976), adapted for use with Z. marina following Pregnall et al. (1987). GS was extracted from shoot tissues and the root-rhizome complex at 4 °C using an extraction buffer solution (50 mM Tris– HCl, 10 mM 2-mercaptoethanol, 1 mM EDTA, 2% PVP; pH 7.5). Activity in the extract was determined by incubating samples in an assay buffer (100 mM Tris– HCl, 30 mM MgSO4, 80 mM glutamic acid, 10 mM ATP, 15 mM hydroxylamine; pH 8.0) at 35 °C over a 45-min. time course. The reaction was terminated by adding a color reagent consisting of 0.37 M ferric chloride and 0.2 M TCA (Lea, 1988). Samples were centrifuged to remove precipitated protein, and absorbance was measured spectrophotometrically at 540 nm. GS was assayed on weeks 3 and 8.

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Fig. 1. Carbon (% DW) and sucrose (mg g− 1 FW) levels in Zostera marina leaf and root-rhizome tissues from controls and treatments as high temperature (HT), high nitrate (HN), and high temperature + nitrate (HTN). Samples were collected in week 4 and week 14 (plants continued to receive experimental treatments 6 weeks beyond PA measurements). Data are given as means ± 1 (SE); significant differences are indicated by asterisks (⁎) and interactions were noted by I (in this case, the addition of NO−3 to elevated temperatures resulted in lower sucrose levels; p = 0.05, n = 3).

Sucrose synthase (SS) and sucrose-P synthase (SPS) activities were assayed following procedures described by Buczynski et al. (1993) and Zimmerman et al. (1995). The enzymes were extracted from Z. marina tissues using a mortar and pestle (4 °C) in an extraction buffer (50 mM Hepes, 15 mM MgCl2, 2% PEG-20, 0.02% Triton-X100, 10 mM DTT, 1 mM EDTA, 1% PVP, 20 mM Ascorbate; pH 7.2). The extract was centrifuged and the supernatant was desalted using a G-25 Sephadex spin column maintained at 4 °C. SS activity was assayed after incubating samples at 37 °C for 20 min. in an assay buffer (10 mM UDP-glucose, 17 mM fructose, and 50 mM glucose-6-P; pH 7.5). The reaction was terminated by adding a 30% KOH solution. The quantity of sucrose produced was determined using an anthrone technique (Van Handel, 1968) described by Huber et al. (1991). SPS was measured similarly, but with 17 mM

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fructose-6-P rather than 17 mM fructose. These enzymes, like GS, were assayed on weeks 3 and 8. Shoot and root-rhizome tissue samples were also analyzed for sucrose concentrations using the anthrone technique of Yemm and Willis (1954) and Van Handel (1968). Samples were assayed on week 4 and week 14 (i.e. 6 weeks beyond PA measurements to observe any amplification in carbon differences). This procedure involved extracting carbohydrates from tissue in 70% hot ethanol. The extract was centrifuged, and the supernatant was collected and allowed to dry in vacuo. The samples were resuspended in hot DI-water prior to analysis. Phosphomonoesterases (PA) have different pH optima for maximum hydrolyzing capacity. As previously noted, this characteristic has led to a common division among functional groups, as AcPAs (pH optimum ca. 5.0) and AlPAs (pH optimum ca. 9.0; Matavulj et al., 1990). Thus, minor activities of one PA group were expected when assaying for another. Phosphomonoesterases were assayed weekly to biweekly following Berman et al. (1990) and Matavulj et al. (1990). Tissues were ground in 0.05 M Tris–HCl buffer (4 °C, pH 8.0). The extract was centrifuged and PAs in the supernatant were assayed for activity at 35 °C, by adding the extract to a solution containing p-nitrophenyl phosphate and a 0.05 M acetate buffer (pH 5.0, for AcPA) or 0.05 M Tris buffer with 1 mM MgCl2 (pH 9, for AlPA). Assays were terminated by adding 0.2N NaOH. The reaction mixture was centrifuged and the optical density of the supernatant was measured spectrophotometrically at 410 nm. 2.6. Data analysis Statistical analyses were completed using the Statistical Analysis System (SAS; SAS Institute, Inc., 1997). Correlation analyses (Pearson) and repeated measures ANOVA (General Linear Models with Least Squares Means post hoc analysis) were performed where appropriate, with days in treatment as the repeated-measures factor. All evaluations were considered significant at p-values less than 0.05. 3. Results Throughout the 8-wk study period, water temperatures remained fairly constant (26.4–26.8 °C and 29.2–31.8 °C in ambient [control and HN] and high temperature treatments [HT and HTN], respectively; Table 1). Other physiochemical variables remained comparable among the different treatments, including pH, dissolved oxygen, light, and nutrients (NH4+ , NO3− ,

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Table 2 Correlations between Zostera marina leaf phosphatase activity (AcPA, AlPA) and environmental variables throughout the experiment, for controls and treatments as high temperature (HT), high nitrate (HN), and combined high temperature and nitrate (HTN) Treatment

Correlation coefficients (PAs and environmental variables) AlPA

TP

NH+4

NO−3

Temp

pH

Salinity

DO

Control Leaf AcPA AlPA

0.769 −

0.426 0.286

− 0.445 − 0.517

−0.215 −0.273

− 0.033 − 0.069

− 0.400 − 0.571

0.271 0.184

− 0.394 − 0.432

HT-Leaf AcPA AlPA

0.890 –

− 0.320 − 0.291

− 0.063 0.008

0.252 0.291

0.580 0.586

− 0.282 − 0.327

0.271 0.180

− 0.394 − 0.420

HN-Leaf AcPA AlPA

0.885 –

− 0.369 − 0.237

0.142 0.039

0.153 0.273

0.327 0.397

− 0.373 − 0.459

− 0.032 − 0.024

− 0.236 − 0.257

HTN-Leaf AcPA AlPA

0.842 –

0.318 0.308

0.338 0.073

0.188 −0.030

0.108 0.340

− 0.454 − 0.479

0.578 0.383

− 0.131 − 0.367

Environmental variables (n = 12) include water-column total phosphorus, ammonium, and nitrate, mean weekly temperature, pH, salinity, and dissolved oxygen (DO). Significant correlations are in bold ( p b 0.05) or bold with italics ( p b 0.01).

1.4 mg g− 1 FW in HTN; Fig. 1). Plants in the HT treatment periodically had higher content of both sucrose (after 14 weeks) and carbon (after 4 weeks) in comparison to controls ( p = 0.01 for sucrose; p = 0.04 for % C in leaf tissue; Fig. 1). There were no significant correlations among PA activities, tissue phosphate levels (P content varied from 0.25 to 0.31%, data not shown), and water-column TP concentrations (Tables 2 and 3), suggesting that plant growth had not been P-limited (Perez and

and TP; Table 1). In contrast, salinity was typically 1 to 2 units greater in the high temperature treatments in comparison to ambient temperature treatments (Table 1). While high temperatures promoted significant increases in sucrose levels after 14 weeks, interactions with nitrate enrichment in HTN treatments significantly decreased sucrose concentrations in both above- and belowground tissues (e.g. leaf tissue sucrose levels in HT plants were 8.7 ± 1.1 mg g− 1 FW, compared to 5.0 ±

Table 3 Correlations between Zostera marina belowground phosphatase activity (AcPA, and AlPA) and environmental variables throughout the experiment, for controls and treatments as high temperature (HT), high nitrate (HN), and combined high temperature and nitrate (HTN) Treatment

Correlation coefficients (PAs and environmental variables) NH+4

NO−3

Temp

pH

0.415 0.289

− 0.439 − 0.765

− 0.166 − 0.360

−0.320 −0.320

− 0.361 − 0.361

0.532 0.602

− 0.175 −0.627

− 0.301 − 0.277

− 0.179 − 0.148

− 0.029 0.026

0.084 0.164

− 0.266 − 0.268

0.476 0.489

− 0.099 − 0.107

HN-belowground AcPA − 0.115 AlPA –

0.401 − 0.450

0.008 − 0.151

0.483 − 0.141

−0.244 0.414

− 0.050 − 0.182

0.484 −0.432

− 0.147 − 0.273

HTN-belowground AcPA 0.927 AlPA –

− 0.526 − 0.424

0.226 0.301

− 0.237 − 0.192

−0.400 −0.364

0.553 0.357

0.039 −0.047

0.572 0.483

AlPA Control-belowground AcPA 0.787 AlPA – HT-belowground AcPA AlPA

0.989 –

TP

Salinity

DO

Environmental variables (n = 12) include water-column total phosphorus, ammonium, nitrate, mean weekly temperature, pH, salinity, and dissolved oxygen (DO). Significant correlations are in bold ( p b 0.05) or bold with italics ( p b 0.01).

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Fig. 2. The effect of temperature on the activity of acid- and alkaline PAs (μmol p-nitrophenyl g FW− 1 min− 1) in Zostera marina leaf tissue at ambient [low] nitrate. Data are given as means ± 1 SE (n ≥ 8). Different letters above error bars denote significant differences between PA activities at the different temperatures.

Romero, 1993). Throughout the study, PA activity was seldom correlated with other environmental parameters such as salinity, water-column NH4+ or water-column NO3− (Tables 2 and 3). Moreover, there were no significant correlations between PAs and shoot SPS activity (r = 0.28 and 0.13 for acid- and alkaline PAs, respectively; data not shown). Nevertheless, PA activities apparently were adjusted during exposure to increased water-column NO3− and/or

Fig. 3. The effect of environmental pH on alkaline PA in Zostera marina leaf tissue. Data are given as means ± 1 SE (n ≥ 4). Different letters above error bars denote significant differences in PA activity at the different pH values.

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Fig. 4. Relationship between the activities of acid- and alkaline PAs and sucrose synthase (SS) in root-rhizome tissues of control Zostera marina.

increased temperatures. Under ambient NO3− conditions, only water temperature and pH were significantly correlated with shoot AcPA and AlPA activities (Table 2). As temperature increased, PA activities generally increased (Fig. 2). The higher Pi demand at elevated temperatures may help to explain the low phosphate levels that were measured in the water column of high-temperature treatments relative to

Fig. 5. Relationship between the activities of acid- and alkaline PAs and glutamine synthetase (GS) in control Zostera marina root-rhizome tissues.

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Fig. 6. Leaf-(A and C) and root-rhizome (B and D) phosphomonoesterase activities (AcPA, A and B; AlPA, C and D) in Zostera marina throughout the experiment in controls and treatments as high temperature (HT), high nitrate (HN), and combined high temperature + nitrate (HTN). Data are given as means ± 1 SE (n = 3); there were no significant differences between controls and treatments.

ambient-temperature mesocosms (e.g. at week 6, TP in controls were 0.42 ± 0.07 μM, compared to 0.29 ± 0.01 and 0.25 ± 0.03 μM TP in the HT and HTN treatments, respectively; Table 1). AlPAs in these alkaline marine waters (pH 7.6–8.6) were significantly correlated with water-column pH via a quadratic interaction, with a pH optimum at 7.8–8.2 (Fig. 3). As expected, this trend was not observed for AcPAs (data not shown). In the root-rhizome complex, PA activities of unenriched plants were significantly correlated with both GS and SS activities. PA activities were positively correlated with SS activity (Fig. 4), and inversely correlated with GS activity (Fig. 5). In contrast, in nitrate-enriched plants, AlPA activities in the root-rhizome complex were negatively correlated with tissue sucrose concentrations (r = −0.73, p = 0.018; data not shown). One of the more interesting findings from this experiment was the apparent partitioning of PA between above- and belowground tissues on some dates. There were no significant alterations in PA activity, relative to controls, in treatments when above- and belowground tissues were considered separately (Fig. 6). However, there were significant interactions between treatments for PA activity when considering tissue partitioning ( p = 0.006 for AcPA control versus N-enriched plants;

and p = 0.011 for AlPA in control versus high temperature plants; repeated-measures ANOVA). The data indicate significant adjustments in PA partitioning within Z. marina under both water-column nitrate enrichment and increased temperature (Fig. 7). On most dates, control plants had shoot: root-rhizome AcPA ratios b1 (mean values ranged between 0.62 and 0.97), indicating that the PA activity in these plants was more localized in belowground structures, or at least comparable in above- and belowground tissues. In contrast, under elevated water-column nitrate and/or increased temperature, the shoot: root-rhizome PA balance shifted in favor of carbon source tissues, with AcPA activities up to 12-fold higher than in belowground tissues on some dates (AcPA ratio of 1.97 ± 0.38, 1.55 ± 0.30, and 2.27 ± 0.59 for HN, HT, and HTN, respectively; Fig. 7). Correlation analyses between PA ratios versus environmental and physiological parameters indicated that both sucrose levels and water-column TP concentrations may have been associated with this response. For HT plants, there was a significant positive correlation between the AlPA ratio and belowground-tissue sucrose levels (r = 0.95; p = 0.003). In HN plants, there was a significant negative correlation between the AcPA ratio and water-column TP concentrations (r = − 0.62,

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Fig. 7. Ratios (log scale) between shoot and root-rhizome phosphomonoesterase activities in Zostera marina throughout the experiment in controls and treatments as high temperature (HT), high nitrate (HN), and combined high temperature + nitrate (HTN). Data are given as means ± 1 SE (n = 3). Significant differences from controls are indicated by an asterisk (⁎).

p = 0.03), and between the AcPA ratio and leaf sucrose levels (r = − 0.91; p = 0.003). 4. Discussion This study indicates the importance of PAs in supplying Pi for Z. marina metabolism at elevated temperatures and water-column nitrate. The response may, in part, be necessary because of the increased carbon demand during periods of nitrate assimilation. Apparent depletions in stored carbon were considered to have been associated with increased energy expenditures from elevated nitrate assimilation/ reduction (Turpin, 1991), and with increased carbon skeletons necessary for synthesis of amino acids and proteins (Banziger et al., 1994; Rabb and Terry, 1995). When water-column nitrate was low, temperature appeared to be a major influence on PA activity, with temperature positively correlated with PA activity. In this seagrass, high temperature also has been associated with significant increases in cellular respiration and, therefore, increased carbon consumption (Biebl and McRoy, 1971; Touchette and Burkholder, 2000a). Similar temperature interactions with PA activity have been observed for various other plant species (e.g., Bhadula et al., 1986; Kroehler and Linkins, 1988; Juma and Tabatabai, 1988). Under most conditions, PA activity generally increases with increasing temperatures (Murthy et al., 1990; Biswas and Cundiff, 1991).

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We also documented positive correlations between SS and PA activities. SS activity is considered to indicate sucrose mobilization from sink tissues during periods of high carbon demand (e.g. increased respiration; Koch and Nolte, 1995). An increase in PA during periods of higher SS activity may assist in providing the Pi necessary to enhance carbon metabolism/mobilization. Chitarra and Lajolo (1981) showed increased PA activity in developing fruits of banana (Musa acuminate Colla x Musa balbisiana Colla) during ripening periods when starch was metabolized to reducing (mostly glucose and fructose) and non-reducing sugars (mostly sucrose). Basra et al. (1989) observed enhanced AcPA initially in germinating maize (Zea mays ssp. Mays) seedlings, and suggested that this enzyme could be important in stimulating germinative metabolism. Thaker et al. (1996) found increased AcPA activity during the secondary thickening phase of cotton (Gossypium hirsutum) cell walls, and suggested that AcPA may play an important role in carbohydrate allocation. Such studies indicate an important role of PAs in plant carbon metabolism by providing the Pi needed for metabolic regulation/processes. The results from this study suggest that there may be an increased metabolic demand for Pi in carbon source tissues as shoots are exposed to higher nitrate and/or increasing temperatures. The response to water-column nitrate may be related to the specific role of shoots in reducing nitrate (Roth and Pregnall, 1988; Burkholder et al., 1992; Touchette and Burkholder, 2001; Alexandre et al., 2004). That is, belowground tissues in this plant apparently may not be as involved in the reduction of nitrate to nitrite. Similar observations were made on Z. noltii, where in vivo nitrate reduction rates were as much as 40 fold higher in leaves relative roots (Alexandre et al., 2004). As Z. marina occurs in nitrogen-limited waters, rapid nitrate reduction in aboveground tissues would be favored over nitrate storage (Touchette and Burkholder, 2000b). Furthermore, the typically hypoxic sediments surrounding belowground structures would favor ammonium over nitrate as the predominant form of inorganic N, thus further diminishing the role of belowground structures in nitrate reduction (Burkholder et al., 1992). Increased cellular Pi has been shown to inhibit SPS and nitrate reductase (NR; EC 1.6.6.1) activities in other plants during light–dark modulation (Huber et al., 1992). We did not sample Z. marina during dark periods, so any interference of potentially increased Pi on SPS regulation in Z. marina cannot be discerned. A direct inhibitory response of Pi would be unlikely during photosynthetic periods, due to the kinetic properties of specific regulatory enzymes such as SPS kinase. This enzyme

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is inhibited/regulated by glucose-6 P levels (Huber and Huber, 1991) that, in turn, are altered during photosynthetic periods rather than directly via Pi availability. Low sucrose levels in root-rhizome tissues were correlated with an increase in AlPA activity in belowground tissues relative to activities in the leaves. If PAs are responsive to C levels/availability, then as sucrose levels in belowground tissues increase, AlPA activity would be expected to decrease in these carbon sink tissues relative to AlPA activities in shoot tissues. Such changes again would support the premise of an increased demand for Pi sequestering during periods of carbon deficit, especially in belowground storage tissues. Invers et al. (1995) reported lower AlPA activities in roots of Posidonia oceanica relative to AlPA activities in leaf tissue. Given that Posidonia tends to accumulate high levels of soluble carbohydrates in belowground tissues (up to ∼ 400 mg g− 1 DW; Touchette and Burkholder, 2000b), the observed low AlPA activity may reflect the high soluble carbohydrate storage capacity of this species. Interestingly, no AlPA partitioning response was observed for any treatment during week 4, which was marked by unusually high irradiance and likely high carbon fixation rates, indicated by light measurements and DO concentrations (mean PAR levels on week 4 ranged from 2000–2500 μE m− 2 s− 1 compared to 1560– 1790 μE m− 2 s− 1 for the other three sample dates, and mean DO levels on week 4 for all treatments ranged from 9.0 to 10.3 mg L− 1 compared to 4.9 to 7.0 mg L− 1). Under these high light conditions, photosynthesis rates may have been high enough to alleviate the need for carbon mobilization between sink and source tissues, thereby maintaining comparable PA levels for both tissue types. The isozyme AcPA also had lower ratio values for week 4, again suggesting that higher irradiance may play a similar role in influencing PA enzyme partitioning. PA activity may also be regulated on a smaller scale, aside from its association with growth-limiting Pi conditions. Sustained, sometimes high PA activities have been observed in plants even when growth is N-limited. Such small-scale adjustments in PA activity and, therefore, P i mobilization may be necessary to enable Z. marina to take advantage of short temporal spikes of nutrients (e.g. short-term nitrate NO3− enrichment from storm water runoff in marine coastal habitats). The PA response could enhance carbon metabolism to help sustain increased carbon flow, which would benefit nitrate uptake processes by supplying energy and/or carbon skeletons for amino acid synthesis and other metabolic processes during nitrogen assimilation. Additionally, as storm

water runoff is often coupled with elevated turbidities (e.g. Mallin et al., 1999), increases in PA activity could also help mobilize storage carbohydrates during periods of high internal carbon demands associated with light limiting conditions. 5. Conclusions The research suggests that PA activity can be influenced by the carbon and nitrogen status of Z. marina. Under low water-column nitrate conditions, shoot PA activity may be affected by environmental temperature and pH. In the root-rhizome tissues, AlPA activity was strongly correlated with SS activity and inversely correlated with GS activity. As water-column nitrate increased, temperature and pH responses apparently were no longer significant in shoot tissues, suggesting an alteration in PA regulation. In the rootrhizome complex, there was a significant interaction between sucrose levels and AlPA activity under elevated water-column nitrate conditions. There also were significant tissue alterations in PA between above- and belowground tissues under increased temperature and/or nitrate. In these treatment conditions, PA activity appeared to shift from the root-rhizome to predominance in the leaf tissue, especially for AcPA. Moreover, this partitioning response was significant between AlPA ratios and sucrose levels in belowground tissues. We hypothesize that alterations in PA activities may be a mechanism that provides the Pi needed for metabolic regulation during short time intervals when this seagrass experiences increased carbon demand. The associated increase in cellular Pi would enable Z. marina to allocate/partition carbohydrates for uptake of inorganic nitrogen compounds, or to enhance carbon metabolism during periods of increased temperature. Acknowledgments Funding support for this research was provided by the North Carolina Sea Grant College Program, the North Carolina Agricultural Research Service, the College of Agriculture and Life Sciences, the Department of Plant Biology at North Carolina State University, and the North Carolina General Assembly. D. Briley, T. Brister, J. Compton, M. Crawford, N. Deamer, E. Fensin, H. Glasgow, E. Allen, M. Larsen, M. McCally, M. Mallin, G. Morgan, J. Springer, and H. Williams assisted in transplanting Z. marina. Plant and environmental sample collections were assisted by T. Brister, G. Burrows, J. Colton, and S. McCarthy. J. Compton, E. Allen, A. Hamilton, and J. Manning. A. Penny assisted in

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