Primary production, nutrient assimilation and microzooplankton grazing along a hypersaline gradient

FEMS Microbiology Ecology 39 (2002) 245^257

www.fems-microbiology.org

Primary production, nutrient assimilation and microzooplankton grazing along a hypersaline gradient Ian Joint b

a;


, Peter Henriksen b , Kristine Garde c , Bo Riemann

b

a NERC Plymouth Marine Laboratory, Prospect Place, The Hoe, Plymouth PL1 3DH, UK Department of Marine Ecology, National Environmental Research Institute, Roskilde DK-4000, Denmark c DHI Water and Environment, Agern Alle¤ 11, H,rsholm DK-2970, Denmark

Received 5 November 2001; received in revised form 9 January 2002 ; accepted 10 January 2002 First published online 15 February 2002

Abstract As part of an investigation of the relationship between diversity and productivity, measurements were made in a solar saltern of carbon fixation, nitrate and ammonium uptake and microzooplankton grazing at salt concentrations ranging from 4 to 37%. Elevated photosynthetic pigment concentrations were present in ponds of intermediate (5^11%) and high ( s 32%) salinity but rates of primary production and nutrient uptake were generally reduced at the highest salinity. Maximum primary production was measured at 8% salinity and chlorophyll-specific carbon fixation also maximised at this salinity. Ammonium was the dominant nitrogen source throughout the salinity gradient; turnover times of ammonium were from 2 to 14 days. Nitrate turnover times were very long (V100 days) at salinities 6 22% but at 37% salinity, nitrate was taken up rapidly by the microbial assemblage in the light and turnover times for the ambient nitrate concentrations in the 37%-salinity pond were between 6 and 12 days. There were large changes in C:N uptake ratio. At salinities 6 11%, the C:N uptake ratio was higher than the Redfield ratio. However, at s 22% salinity, the C:N uptake ratio was V1. That is, much more nitrate and ammonium were taken up than would be expected from the observed carbon-fixation rates. Although primary production declined with decreasing phytoplankton diversity along the salinity gradient, there was no clear relationship between heterotrophic activity and microbial biodiversity. ; 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : Solar saltern; Primary production; Nutrient uptake; Microzooplankton grazing

1. Introduction The relationship between biodiversity and productivity is part of a lively debate in terrestrial ecology. It has been suggested that an overall increase in the total number of species present in a biotope is associated with increased community productivity [1]. However, this hypothesis has been challenged by Grime [2] who argued that manipulation experiments may alter conditions and result in changes in assemblage productivity which are not necessarily related to general biodiversity. Lehman and Tilman [3] have proposed models which suggest that greater diversity increases the stability of the entire community and also

* Corresponding author. Tel : +44 (1752) 633100; Fax: +44 (1752) 633101. E-mail address : [email protected] (I. Joint).

increases productivity. However, proof of these hypotheses remains di⁄cult within terrestrial ecosystems. Terrestrial environments are characterised by high standing stocks of primary producers and relatively low generation time. In these circumstances, it is di⁄cult to establish a direct link between biodiversity and productivity because the response time of the primary producers tends to be slow. In contrast, marine systems can be characterised as low biomass, high-turnover systems. That is, the dominant primary producers are microalgae which have generation times of the order of 1 day, but which rarely form signi¢cant accumulations of biomass. These microbial-dominated marine ecosystems o¡er an opportunity to investigate the relationship between biodiversity and productivity in highly productive systems. However, manipulation of marine ecosystems is less easy than terrestrial systems. Marine assemblages are subject to continuous dispersion within the aquatic environment, making it di⁄cult to follow a cohort through several generations ^

0168-6496 / 02 / $22.00 ; 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII : S 0 1 6 8 - 6 4 9 6 ( 0 2 ) 0 0 1 7 8 - 2

FEMSEC 1330 6-5-02

246

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

even when that may only take a few days. Attempts to circumvent mixing and dispersion have largely involved mesocosm studies but these can be criticised because containment of an assemblage may cause artefacts ; in particular, reduced turbulence results in faster sinking rates. The walls of a mesocosm may also develop atypical assemblages, which can have undue in£uence on the bulk of the contained water. Solar salterns o¡er an alternative experimental system with a strong gradient in biodiversity of primary and secondary producers. At one end of the gradient is seawater, which contains a wide diversity of phytoplankton, zooplankton and heterotrophic microbes. At the other end of the gradient, at extreme salinities where salt crystallises from solution, phytoplankton biodiversity is generally reduced to a single organism, Dunaliella salina, and there are no mesozooplankton. The microbial community-structure changes from that in the sea, where V75% of the prokaryotes are bacteria and the rest are archaea of the marine clade, to high salinities which are dominated by extremophile haloarchaea. At intermediate salinities, mesozooplankton diversity is restricted to one species, Artemia salina, but this is absent from the crystalliser ponds with saturating salt solutions. The other major group of grazers is the microzooplankton, but less is known about the biodiversity of microzooplankton in these systems. Therefore, the salinity range maintained within the solar salterns controls biodiversity and o¡ers an environment to test the relationship between biodiversity and production within a marine system of low biomass and high productivity. The hypersaline environment studied was a solar saltern operated for the commercial production of sea salt at Bras del Port, Santa Pola, Alicante, Spain (38‡12PN, 0‡36PW). This saltern has been the subject of a number of microbiological studies of extremophile bacteria and archaea [4^7] and microbial food webs [8,9] but productivity data are lacking. The experiments took place between 17 and 28 May, 1999. This paper concentrates on aspects of production and we describe the measurement of primary productivity, nutrient uptake and microzooplankton grazing. The measurements were part of a larger study of microbial biodiversity along the salinity gradient, based on 16S rRNA sequence data [10].

2. Materials and methods 2.1. Study site The salterns comprise a large number of ponds, which are managed to maximise salt production throughout the year. Seawater is periodically transferred from one pond to another as evaporation progresses so that salt precipitation and harvesting takes place in a restricted number of ponds. There are over 100 ponds in the saltern complex

but only eight were sampled in this study; those had salinities of 4, 5.4, 8, 11, 15, 22.4, 31.6 and 37%. Samples were taken between 18 and 27 May, 1999. Salinities were determined in the ¢eld with a hand-held refractometer; samples with salinities s 13% were diluted with distilled water to bring the salt concentration within the range of the refractometer [8]. Each pond is shallow ( 6 1 m depth) and samples were taken from the surface with an all-plastic sampler, taking care not to disturb the sediment surface. Sampling was done from the saltern edges, avoiding the corners of the ponds where wind-driven detritus tended to accumulate. 2.2. Pigment analysis Phytoplankton pigments were analysed according to the method of Wright et al. [11] with minor modi¢cations as described in Schlu«ter and Havskum [12]. Water samples were kept in the dark and ¢ltered, generally within 3 h of sampling. At salinities 922%, samples of 100^300 ml were ¢ltered onto 25-mm Advantec GF 75 glass-¢bre ¢lters (Toyo Roshi Kaisha, Japan) and placed immediately in liquid nitrogen. Due to the high biomass and viscosity at salinities s 22%, ¢ltering was extremely slow and the ¢ltered volumes were reduced to 10^20 ml. Filters were transferred to 3 ml acetone, sonicated on ice for 15 min, and left for extraction of pigments for 24 h at 4‡C. Pigment extracts were ¢ltered (0.2 Wm) into high-performance liquid chromatography (HPLC)-vials and water was added (300 Wl water to 1 ml pigment extract). Pigments were analysed with a Shimazu LC 10A HPLC system with a Supercosil C18 column (250U4.6 mm, 5 Wm). Pigments were identi¢ed from retention times and absorption spectra identical to those of authentic standards, and quanti¢ed against standards purchased from the International Agency for 14 C Determination, HPrsholm, Denmark. 2.3. Nitrate and ammonium analysis Nitrate and ammonium concentrations were determined as soon as practicable after sampling, which was always within 3 h of collection. In the period between sampling and analysis, samples were stored in an insulated box in the dark. Nutrient concentrations were determined by colorimetric determination using manual methods based on Brewer and Riley [13] for nitrate, Grassho¡ [14] for nitrite and Mantoura and Woodward [15] for ammonium. Analysis was complicated because the high salt concentration interfered with the chemistry. In the analysis of nitrate, there were two distinct problems. The e⁄ciency of reduction of nitrate to nitrite in the copper^cadmium reduction column was greatly reduced at high salinities. Also, there was a large e¡ect of high salt on colour development in both the nitrite and ammonium assays. There was little e¡ect at salt concentrations 6 10% and all samples of higher salinity were diluted with deionised water to

FEMSEC 1330 6-5-02

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

6 10% salinity to overcome the salt e¡ect on colour development. Care was taken to accurately determine blank concentrations and the e⁄ciency of nutrient determination was established by standard additions of nitrate, nitrite or ammonium to replicates of each sample.

247

bonic acid vary with changing ionic strength [17]. In order to estimate the concentration of dissolved CO2 /bicarbonate, total alkalinity measurements were adjusted for the e¡ect of high-salt concentrations by determining values for the empirical factor fH [18] which is related to the apparent ion-activation coe⁄cient of Hþ [19].

2.4. Primary-production determination 2.5. Nitrate and ammonium uptake The rate of carbon ¢xation was estimated from the incorporation of [14 C]bicarbonate. Water samples were collected at dawn from four ponds using an all-plastic sampler. Water samples were transferred into four 60-ml clear, and two black, polycarbonate bottles; all bottles were cleaned following JGOFS protocols [16] to reduce trace metal contamination. Each bottle was inoculated with 370 kBq (10 WCi) NaH14 CO3 and incubated for 24 h under the water surface of the ponds from which samples were taken. Incubation periods of 24 h were used since they include a night period, hence giving estimates closer to net than to gross primary production. Two experiments were also done in the laboratory. One shorttime period incubation (3 h) compared activities in all eight experimental ponds and one long-time period incubation (14.5 h) tested if the availability of nutrients (i.e. ammonium and/or phosphate) was limiting primary production in the ponds of 4, 8, 11 and 37 salinity. Nutrients were added to give ¢nal concentrations of 25.2 Wmol l31 NH4 Cl and 1.57 Wmol l31 KH2 PO4 . In both experiments, duplicate light bottles and a single dark bottle from each pond were incubated in a 30‡C constanttemperature laboratory, close to the temperature of the ponds, with a quartz^halogen light source which gave an irradiance of 275 Wmol m32 s31 at the front surface of each bottle. At the end of the incubation period, each sample was ¢ltered through 0.2-Wm pore-size polycarbonate ¢lters. The ¢lters were fumed in HCl for 30 min and dried in a desiccator for 24 h. 14 C was measured in a liquid scintillation counter (LSC), the e⁄ciency of which was determined with an external standard, channels ratio method. The quantity of 14 C added to the experimental bottles was determined by adding aliquots of the stock 14 C solution to vials containing CO2 -absorbing scintillation cocktail, which were counted in the LSC. The calculation of primary-production rate from uptake of 14 C requires quanti¢cation of the dissolved carbonate system. However, high-salt concentrations have a profound e¡ect on the overall carbonate chemistry. Precipitation of di¡erent elements takes place along the evaporation gradient; calcium carbonate is the ¢rst mineral to precipitate followed by gypsum (CaSO4 W2H2 O) and ¢nally NaCl precipitation occurs when the volume of the solution reaches V10% of the original seawater volume. These chemical changes a¡ect a number of aspects of the carbonate system; CO2 solubility decreases with increasing salinity, and the apparent dissociation constants of car-

Assimilation rates for nitrate and ammonium were determined by inoculating water samples with the stable isotope 15 N. Samples from each pond were distributed into 640-ml clear polycarbonate bottles and 15 N^NO3 3 and 15 31 N^NHþ 4 were added at 0.5 Wmol l . Duplicate samples in clear bottles, and one dark bottle, were incubated in situ with each tracer for 24 h. At the end of the incubation period samples were ¢ltered onto ashed Whatman GF/F ¢lters. The GF/F ¢lters were washed with ¢ltered seawater and stored frozen until analysis in Plymouth. Filters were oven-dried at 50‡C before analysis. Particulate nitrogen and atom percent 15 N were measured by continuous-£ow nitrogen analysis^mass spectrometry (Europa Scienti¢c Ltd., UK) using the techniques described by Owens and Rees [20]. Rates of assimilation were calculated from the equations of Dugdale and Goering [21]. 2.6. Bacterial activity determined by leucine incorporation Bacterial productivity was determined from rates of incorporation of L-[4,5-3 H]leucine (speci¢c activity 148 Ci mmol31 ; Amersham Pharmacia Biotech) using the microcentrifuge method of Smith and Azam [22]. Samples were taken from the 4-, 11-, 22- and 37%-salinity ponds at dawn on 21 May, 1999. Four replicate 1.7-ml aliquots from each pond were inoculate with leucine, and glutaraldehyde, at a ¢nal concentration of 2.5% v/v, was added to an additional sample from each pond as control. [3 H]Leucine was added to each tube to give ¢nal concentrations of 20 nmol l31 and samples were incubated for 1 h in a temperature-controlled block ( S 0.1‡C). An additional seven samples from each pond were incubated with leucine at concentrations between 5 and 50 nmol l31 to establish the concentration of leucine which saturated uptake; this was below 20 nmol l31 in all ponds. Time-course samples showed that incorporation was linear over the standard incubation period of 1 h. At the end of the incubation, samples were transferred to an ice/water bath and ice-cold trichloroacetic acid (TCA) added to give a ¢nal concentration of 5% v/v. After 15^30 min, the samples were centrifuged for 15 min. The supernatant was removed by aspiration, 1 ml 5% ice-cold TCA added and the tubes were shaken for 10 min before re-centrifugation. The supernatant was discarded, scintillation cocktail was added to the centrifuge tubes and counted in a LSC, the e⁄ciency of which was determined by the external standard, channels ratio method.

FEMSEC 1330 6-5-02

248

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

2.7. Microzooplankton grazing Microzooplankton grazing was assessed by the dilution method [23,24]. In ponds with salinities of 4, 8 and 11%, dilution experiments were performed in 1.5-l polycarbonate bottles submerged in situ in the surface water for 24 h. The viscosity of water in the 37% pond precluded ¢ltration of large volumes, and therefore the dilution experiments in this pond were done in 100-ml polycarbonate bottles each containing a total of 75 ml water. Pond water (pre-¢ltered through 200 Wm mesh to remove mesozooplankton) was diluted with 0.2 Wm ¢ltered pond water to provide ¢nal plankton densities of 25, 50, 75 and 100% ambient levels. For each dilution, nutrients were added to triplicate bottles according to Schlu«ter [25]. For each salinity, two additional bottles with undiluted pond water without nutrient addition were included to determine if phytoplankton growth was nutrient-limited. Initial and ¢nal pigment contents in each dilution were determined as described above. Apparent growth rates (k) related to individual pigments were calculated assuming exponential growth: 1 Pt k ¼ Ulnð Þ t P0 where t is time in days and P0 and Pt are the initial and ¢nal pigment concentrations, respectively. Subsequently, the apparent growth rates were plotted as a function of dilution, and microzooplankton grazing rates (m) and potential growth rates (W) of phytoplankton were determined from linear regressions analysis assuming : k ¼ W 3ðmUDi Þ where Di is the dilution factor [26]. Grazer impact on prey concentration was assumed only if there was a signi¢cant (P 6 0.05) negative slope and grazing was assumed to be zero if the slope was not signi¢cant. 2.8. Susceptibility to system perturbation

along the gradient with the most obvious changes occurring at the highest salinity. The ponds with s 30%-salt concentrations, and particularly the crystalliser ponds from which salt is harvested, were bright red due to the abundance of D. salina and pigment-containing halobacteria. The water was not appreciably coloured in the lower-salinity ponds. Fig. 1 shows the chlorophyll-a concentration along the salinity gradient. The highest chlorophyll-a concentrations were measured in the crystalliser ponds and chlorophyll-a concentrations were s 14 Wg l31 . Chlorophyll-a concentrations were low in the ponds with salinities of 15^25%, higher (V8 Wg l31 ) in the ponds with salinities of 5.4^10.2% salt and were 6 4 Wg l31 in the initial pond which was fed directly with seawater from the Mediterranean Sea. 3.2. Nutrient concentrations Nitrate and ammonium concentrations along the salinity gradient are shown in Fig. 2. Data are shown for all ponds which were sampled at the beginning of the experiment on 18 May, but subsequently, emphasis was on ponds with salinities of 4, 8, 22 and 37%. Generally, the concentration of nitrate was higher than that of ammonium. Between salinities of 5.4 and 15%, the nitrate concentration was V3^4 Wmol l31 . On 18 May, the nitrate concentration in the 4%-salinity water was slightly higher at 5.5 Wmol l31 but the highest nitrate concentrations were present at the highest salt concentrations. At 31.6 and 37% salinity, the nitrate concentration was 6.2 Wmol l31 . Ammonium concentrations showed little variation along the salinity range, ranging between 1.5 and 2.4 Wmol l31 in all ponds except the 22% salinity which had a very low-ammonium concentration of 0.4 Wmol l31 . The highest ammonium concentration measured on 18 May was 3.9 Wmol l31 in the 37%-salinity pond. In the pond fed directly from the sea (4% salinity) a low concentration of 1.1 Wmol l31 was recorded. Interestingly, this pond was also used as a

In order to test the fragility or robustness of the assemblage in the 37%-salinity pond, the water was diluted with deionised water to simulate the e¡ect of severe rainfall. The largest storm in recent years would have resulted in dilution of the order of 25%. Four polycarbonate carboys were autoclaved and ¢lled with water from the 37% pond. Two of the carboys had sterile deionised water added to give a ¢nal dilution to 27.75% salinity. All four carboys were placed in situ in the 37%-salinity pond and sampled daily for pigment concentrations and microbial biodiversity.

3. Results 3.1. Phytoplankton biomass There were large variations in phytoplankton biomass

Fig. 1. Chlorophyll-a concentration determined by HPLC analysis, in each of the experimental ponds sampled on 26 May 1999 ; duplicate samples were analysed from all ponds except at 25 and 31.6% salt.

FEMSEC 1330 6-5-02

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

Fig. 2. Nitrate and ammonium concentrations determined along the salinity gradient on 18, 21 and 25 May 1999. (F) Nitrate concentration on 18 May, (8) nitrate on 21 May, (b) nitrate on 25 May, (E) ammonium on 18 May, (7) ammonium on 21 May and (a) ammonium on 25 May.

production facility for ¢sh and a high biomass of ¢sh was present. Nitrate and ammonium were assayed on two subsequent dates during the experiment ^ 21 May and 25 May, 1999. There were some di¡erences in both nitrate and ammonium concentration (Fig. 2) but generally little change in concentration was measured over the period of the experiment. The most signi¢cant di¡erences were a decline in nitrate concentration in the 4%-salinity pond and variability in the 37%-salinity pond, which ranged between 5.4 and 7.3 Wmol l31 . Ammonium generally showed little change in concentration except in the 22%salinity pond. On 18 May, ammonium was 0.4 Wmol l31 but by 21 May this had increased to 2.4 Wmol l31 and by 25 May the ammonium was 7.1 Wmol l31 . The 22%-salinity pond showed the greatest variability in concentration during the experiment.

249

Fig. 3 also shows the chlorophyll-speci¢c rates of carbon ¢xation at each salinity. These are not estimates of PBm , because no measurements of photosynthesis/irradiance characteristics were made. However, the experiments were done at 275 Wmol quanta m32 s31 which is assumed to be below saturation irradiance and so give scope for using chlorophyll-speci¢c rates to compare activities in the different ponds. It is clear that the chlorophyll-speci¢c rates showed a very similar pattern to the carbon ¢xation rate. There was a maximum of chlorophyll-speci¢c carbon ¢xation (PB ) of 4.9 Wg C Wg31 Chla h31 in the sample from the 8%-salinity pond. At the lowest salinity, PB was 2.5 Wg C Wg31 Chla h31 and it was 0.1 Wg C Wg31 Chla h31 at the highest salinities of the crystalliser ponds. Therefore, at these very high salinities, although there was a high standing stock of chlorophyll, there was much less carbon ¢xation per unit of chlorophyll than at any other salinity. Primary-production estimates, based on 24-h in situ incubations, are given in Table 1. Only four ponds were sampled and simultaneous measurements of nitrate and ammonium uptake were made. Table 1 shows signi¢cant variability in primary production on the two dates sampled. On 21 May, the primary-production rate in the 37%-salinity pond was 27 mg C m33 day31 ; 4 days later on 25 May, this had increased to 56 mg C m33 day31 . However, in the less-saline ponds, even greater variation was measured. At 11% salinity, primary-production increased from 37 to 380 mg C m33 day31 and at 4 and 22% salinities, there were three-fold increases in production. The source of this variability is not clear. Continuously measured PAR irradiance showed no large di¡erences in irradiance during the 10-day sampling period. The most likely reason for the observed variability was lateral patchiness within the ponds. As a measure of within-pond heterogeneity of phytoplankton abundance, D. salina was

3.3. Primary production Primary production was measured in two ways. In situ incubations for 24 h were done in four ponds on 21 and 25 May to estimate net primary production and a laboratory experiment on 26 May assessed the relative rates of photosynthesis in all eight ponds under investigation. Fig. 3 shows the range in potential photosynthesis from the latter experiment. It is clear that the potential for photosynthesis was greatest at the lower end of the salinity range. The maximum rate of carbon ¢xation (38.6 Wg C l31 h31 ) was measured in water samples taken from the 8%-salinity pond. Elevated carbon ¢xation was also measured at salinities between 5.4 and 11%. These ponds showed signi¢cantly greater carbon ¢xation than was measured in the lowest-salinity pond (4% salt) where the carbon ¢xation rate was 7.8 Wg C l31 h31 . In spite of the high chlorophyll concentrations in the crystalliser ponds, the rates of carbon ¢xation were very low.

Fig. 3. Carbon ¢xation rate determined by incubation in arti¢cial light in the laboratory for 3 h. (b) Carbon ¢xation (Wg C ¢xed l31 h31 ) and (a) chlorophyll-speci¢c carbon ¢xation (Wg C ¢xed Wg31 Chla h31 ). The error bars indicate 95% con¢dence intervals which were usually less than the size of the symbol.

FEMSEC 1330 6-5-02

250

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

Table 1 Primary production (PP) measured by in situ incubations on 21 and 25 May 1999 Salinity (%)

4 11 22 37

light dark light dark light dark light dark

21 May

21 May

25 May

25 May

Ratio

mean PP ( S S.D.) Wg C l31 day31

light^dark ratio

mean PP ( S S.D.) Wg C l31 day31

light^dark ratio

25/21 May

61.0 47.0 37.3 8.6 5.2 3.7 27.5 20.8

1.31

239.1 123.5 379.6 28.1 17.2 6.0 56.0 23.8

(4.5) (5.8) (2.1) (3.5) (1.5) (0.3) (8.7) (6.8)

4.34 1.44 1.32

(31.9) (10.0) (29.1) (5.2) (1.0) (1.2) (10.9) (8.4)

1.94 13.52 2.87 2.35

3.87 2.63 10.18 3.27 3.28 1.64 2.04 1.15

Means and standard deviations of four samples incubated in the light and two samples incubated in the dark for 24 h beginning at dawn.

counted in four samples collected from the 37% crystalliser on 25 May, one sample being taken from each side of the pond. There was a large variation in cell densities. Cell numbers were 5090, 6629, 6728 and 10500 Dunaliella ml31 on the four sides of the pond. Pigment concentrations were also measured on these samples and Dunaliella cell numbers correlated strongly (r2 = 0.983) with chlorophyll-a concentrations, which ranged from 23.6 to 45.7 Wg Chla l31 . This latter value was more than twice that recorded in the same pond on 18 and 25 May. Lateral heterogeneity and patchiness is a likely explanation for the day-to-day di¡erences in primary-production rates. Dark carbon-¢xation rates were high and, particularly on 21 May, were a signi¢cant proportion of the rates measured in the light. The ratio of light^dark ¢xation was generally V1.3 except in the 11%-salinity pond where dark carbon ¢xation was about one-quarter that in the light. On 25 May, when there was an overall increase in carbon ¢xation in the light, there was some increase in dark ¢xation at all salinities but a much greater increase occurred in CO2 ¢xation in the light. That is, there appeared to be an overall stimulation of photosynthetic carbon ¢xation which was not directly matched with large changes in dark carbon ¢xation, suggesting changes to the autotrophic but not the heterotrophic community. 3.4. Nutrient uptake Uptake of nitrate and ammonium was measured on the same water samples as were used for primary production; 24-h in situ incubations were done on 21 and 25 May and the data are shown in Table 2. At the lowest salinity (4%), although ambient concentrations of nitrate and ammonium were approximately the same, ammonium uptake was much greater than that of nitrate. There was very little nitrate uptake in the dark and ammonium uptake was much higher in the light than the dark bottles. In the 11%-salinity pond, nitrate uptake was very low on both days; ammonium uptake was only 0.3 Wmol N l31 day31 on 21 May but increased to 1.9 Wmol N l31 day31 on 25 May in the light. At 22% salinity, nitrate uptake was again very low and there were similar increases in ammonium

uptake between the experiments on 21 and 25 May. In the crystalliser pond at 37% salinity, nitrate uptake was much higher than in any of the other ponds. However, ammonium was still the major nitrogen source and ammonium uptake in the light was 2.3^6 times greater than nitrate uptake. The ambient concentrations of nitrate and ammonium showed some di¡erences between 21 and 25 May with a generally higher concentration being present on 25 May in all ponds except the 11% salinity. Combining concentration data with measured uptake rates allows calculation of the time required to utilise all of the nitrate or ammonium present in the ponds at the start of the experiment (i.e. the turnover time) and the data are also shown in Table 2. With the low measured nitrate-uptake rates, the turnover time in the 4^22%-salinity ponds was very long. Indeed, the decrease of nitrate concentration of 1.6 Wmol N l31 in the 11%-salinity pond could not be explained by the measured nitrate assimilation on 21 May. There must have been a process other than pelagic nitrate uptake to account for the observed change in nitrate concentration. In the crystalliser pond (37%), nitrate turnover by the pelagic community was clearly much more signi¢cant and turnover times are estimated to be between 6 and 10 days. The turnover of ammonium was even more rapid. Ammonium was clearly the most important nitrogen source throughout the salinity range, with particularly rapid turnover times in the 37%-salinity pond.

Fig. 4. C:N uptake ratios determined from 24-h in situ incubations in ponds of salinity of 4, 11, 22 and 37% on 21 and 25 May 1999.

FEMSEC 1330 6-5-02

5.78 4.86

1.55 0.86

2.02 0.44

12.70 183.97 1.53 6.34 227.26 594.37 0.54 2.14 246.07 432.90 4.59 8.26 9.17 15.45 1.00 1.14

1.97 0.50

Total NO3 +NH4 assimilation Turnover time (days)

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

251

The uptake ratios for carbon and nitrogen are shown in Fig. 4; these are the ratio of carbon assimilation (Wmol C l31 day31 ), in the 24-h in situ incubation in the light, to the uptake of nitrate plus ammonium (Wmol N l31 day31 ) for the same period. In the 4%-salinity pond, the C:N uptake ratio was 4.4^9.8, at 11% salinity, it varied between 9.5 and 16.0 but was much lower at 22% salinity (1.2^0.9) and 37% salinity (0.7^0.8). The C:N uptake ratios were signi¢cantly di¡erent to the Red¢eld ratio in all ponds.

(0.06) (0.01) (0.09) (0.02) (0.01) (0) (0.21) (0.06) (0) (0) (0.10) (0.01) (010) (0.02) (0.65) (0.16) 0.26 0.02 1.75 0.42 0.01 0.00 1.96 0.49 0.01 0.01 1.54 0.86 0.80 0.47 4.98 4.39 3.3 3.3 2.7 2.7 2.5 2.5 1.1 1.1 2.9 2.9 7.1 7.1 7.3 7.3 5.0 5.0 3.15 1.72

0.38 0.23

nitrate nitrate ammonium ammonium nitrate nitrate ammonium ammonium nitrate nitrate ammonium ammonium nitrate nitrate ammonium ammonium

2.7 2.7 2.3 2.3 4.1 4.1 4.0 4.0 2.6 2.6 2.3 2.3 5.4 5.4 2.3 2.3

0.12 0.02 1.12 0.59 0.03 0.03 0.30 0.10 0.02 0.02 0.35 0.21 0.94 0.06 2.21 1.66

(0.02) (0) (0.08) (0.03) (0.01) (0.01) (0.04) (0) (0) (0) (0.01) (0.01) (0.75) (0.47) (0.10) (0.13)

0.33 0.12

1.24 0.61

22.36 125.41 2.06 3.89 142.13 159.32 13.42 41.00 120.10 143.54 6.49 10.79 5.78 84.19 1.04 1.39

Ambient concentration (Wmol N l31 ) Total NO3 +NH4 assimilation Turnover time (days)

37

22

11

L D L D L D L D L D L D L D L D 4

Ambient concentration (Wmol N l31 )

Mean assimilation rate ( S S.D.) Wmol N l31 day31

25 May 21 May Light^dark

Nutrient

We were interested to discover which, if any, nutrient might be limiting primary productivity in these systems. Fig. 5 shows the result of adding ammonium and/or phosphate on carbon ¢xation. No statistically signi¢cant stimulations of carbon ¢xation were observed at any salinity. At 8% there was a slight stimulation by added phosphate and ammonium plus phosphate but not by ammonium alone, which suggests that there may have been a tendency towards phosphate limitation in this pond. However, again the di¡erences were not signi¢cant. It appears that photosynthetic carbon ¢xation is not limited by nutrient supply at any salinity. The same conclusion was obtained from the dilution experiments incubated in situ (see Section 3.7). Here, apparent growth rates of phytoplankton in the undiluted pond water with added nutrient were not di¡erent from those incubated without nutrients. 3.6. Prokaryote-community activity The activity of the prokaryote community of bacteria and archaea was determined from the incorporation of [3 H]leucine. One experiment was done on 21 May, 1999 on the same water samples that were used to measure carbon ¢xation and nitrate and ammonium uptake (Fig. 6). The highest rate of leucine incorporation was measured in the pond with 4% salt and the lowest uptake was at 37% salt, which was 21% that in the 4% pond. Interestingly, the prokaryote standing stock showed the opposite trend with highest numbers in the 37% pond (3.4U107 ) and lowest (1.2U107 ) at seawater salinity. There was no correlation between leucine uptake and carbon ¢xation in the dark (Table 1). Under some circumstances, dark carbon ¢xation can be a proxy for heterotrophic activity but in this case, it is probable that there was some chemolithotrophic carbon ¢xation which occurred in the dark. 3.7. Microzooplankton herbivory

Salinity (%)

Table 2 Nitrate and ammonium uptake rates estimated from the incorporation of

15

N in 24-h in situ incubations on 21 May and 25 May 1999

Mean assimilation rate ( S S.D.) Wmol N l31 day31

3.5. Nutrient limitation

Microzooplankton grazing was assessed by measuring pigment concentration in water samples in which the grazing pressure of microzooplankton was reduced by dilution [24]. Table 3 shows estimates of microzooplankton grazing determined on di¡erent pigments. The results from these experiments were very variable. Grazing rates on, and

FEMSEC 1330 6-5-02

252

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

growth rates of, phytoplankton can only be estimated where a statistically signi¢cant linear regression is obtained between dilution and the reciprocal of logarithmtransformed change in pigment concentration over the incubation period. In many cases, no statistically signi¢cant relationship was found. Although experiments were done at 4, 8, 11 and 37% salinities, the greatest number of statistically signi¢cant relationships were obtained at 4% salinity and none were found in the crystalliser (37% salinity) pond. Di¡erent pigments resulted in di¡erent estimates of grazing because the variations in pigment concentration were not proportional to one another. Grazing rates determined for individual pigments showed selective grazing on speci¢c groups of organisms and variations between ponds (Table 3). In the 4% pond, almost identical and very high grazing rates (1.9 day31 ) were found for chlorophyll b and violaxanthin, two pigments characteristic of the prasinophytes that do not contain prasinoxanthin [27]. The grazing rates on chlorophyll c (0.51 day31 ) and fucoxanthin (0.55 day31 ), the pigment combination characteristic of diatoms and chrysophyte £agellates [27], were very similar; the latter only signi¢cant at the 10% level, however. In the pond with salinity of 8%, the grazing rate of chlorophyll c (0.30 day31 ) resembled that for alloxanthin (0.33 day31 ), which could be due to grazing of cryptophytes or other alloxanthin-containing organisms. In addition, signi¢cant grazing rates were determined for chlorophyll b, lutein and zeaxanthin (0.46, 0.22 and 0.26 day31 , respectively), indicating grazing on chlorophytes and possibly also cyanobacteria. At a salinity of 11%, the con¢dence limits on the grazing coe⁄cients were broad. The only signi¢cant regressions were those for fucoxanthin and chlorophyll c, suggesting that there was grazing on diatoms or other fucoxanthin-containing organisms. The changes in pigment concentrations in the crystalliser pond suggest that the apparent phytoplankton growth rate was zero and

Fig. 6. Prokaryote production determined on 21 May, 1999 by the incorporation of [3 H]leucine ; data are mean uptake rates and 95% con¢dence intervals are indicated.

there was no indication of microzooplankton grazing. These estimates of no growth contrast with the 14 C-uptake rates (Section 3.3) which suggest that, although primary production was very low, there was measurable carbon ¢xation. At 4% salinity, the grazing rate based on chlorophyll a and chlorophyll c was about half of the estimated growth rate. However, analysis of the same samples for chlorophyll b indicated that grazing and phytoplankton growth were approximately in balance. Potential growth rate was not determined in the 8%-salinity pond due to prolonged storage in the dark between sampling and ¢ltration. Grazing rates in this pond were approximately half those measured in the 4%-salinity pond. Fewer signi¢cant data were obtained at 11% salinity ; the major pigments, chlorophyll a and b, did not provide any estimates of grazing rate but the estimates from changes in chlorophyll c suggest that the magnitude of microzooplankton grazing was many times the potential phytoplankton growth rate. 3.8. Susceptibility to perturbation

Fig. 5. E¡ect of added nutrients on carbon ¢xation rates. Ammonium chloride was added as nitrogen source to give a ¢nal concentration of 25.2 Wmol N l31 and potassium phosphate was added to give 1.57 Wmol P l31 . Samples were incubated at constant light in the laboratory for 14.5 h.

Although the phytoplankton biomass in the crystalliser ponds was high, it was not clear if this was a stable assemblage and how susceptible it might be to perturbation. Water from the crystalliser pond was diluted with sterile water to 27.8% salinity to investigate how diversity might be a¡ected. Analysis of the bacterial and archaeal community by DGGE showed no changes at all in the assemblage structure (unpublished data), suggesting a very robust community structure. Analysis of the pigment composition also showed little change. Fig. 7 shows the concentrations of chlorophyll a and L-carotene in duplicate samples diluted with sterile distilled water and compared with duplicate, undiluted samples incubated under identical conditions. The undiluted samples showed very little variation over a 7-day period in either chlorophyll a or L-carotene content. Similarly, the diluted samples recovered their pigment content quickly and after 3 days,

FEMSEC 1330 6-5-02

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

253

Table 3 Determination of grazing rate (herbivory) and potential growth rate at 4, 8 and 11% salinity Grazing rate (day31 )

Lower 95%

Upper 95%

Potential growth rate (day31 )

Lower 95%

Upper 95%

6.82 0.60 1.46 0.12 0.43 0.16 0.15 1.74 0.21 0.14 0.47

0.541* 1.920*** 0.517**

0.032 1.425 0.206

1.050 2.415 0.829

0.993*** 1.785*** 1.115*** 0.751* 1.840***

0.633 1.435 0.894 0.150 1.350

1.352 2.135 1.335 1.352 2.329

1.905***

1.406

2.405

0.660** 2.198*** 0.840*** 1.088**

0.374 1.845 0.543 0.613

0.946 2.551 1.138 1.562

16.91 1.69 3.68 0.57 0.29 0.18 6.87 0.91 1.06 0.48 0.61 0.78

0.260*** 0.458* 0.303**

0.142 0.050 0.143

0.377 0.866 0.463

0.333***

0.189

0.478

0.220*** 0.255***

0.119 0.140

0.322 0.371

0.673*

0.005

1.341

0.330*

0.048

0.613

Initial pigment concentration in undiluted water (Wg l31 ) 25^26 May, 4% salinity Chlorophyll a Chlorophyll b Chlorophyll c Peridinin Fucoxanthin Neoxanthin Prasinoxanthin Alloxanthin Violaxanthin K-Carotene L-Carotene 26^27 May, 8% salinity Chlorophyll a Chlorophyll b Chlorophyll c Peridinin Fucoxanthin Neoxanthin Alloxanthin Violaxanthin Lutein Zeaxanthin K-Carotene L-Carotene 19^20 May, 11% salinity Chlorophyll a Chlorophyll b Chlorophyll c Fucoxanthin Zeaxanthin L-Carotene

1.98 0.19 0.17 0.36 0.18 0.09

1.426* 0.717**

0.285 0.320

2.566 1.113

Initial pigment concentrations in undiluted water are given to illustrate the pigment-based diversity of the phytoplankton community. Grazing and growth rates with upper and lower limits of 95% con¢dence intervals are given when signi¢cant. Due to prolonged storage in the dark between sampling and ¢ltration of samples from the 8% salinity pond, potential growth rates were not determined. *P 6 0.05; **P 6 0.01; ***P 6 0.005.

L-carotene concentrations were the same as in the undiluted samples. Chlorophyll a showed slightly more variation but, within 4 days, concentrations were not signi¢cantly di¡erent from the undiluted samples. So, in terms of both phytoplankton pigments and microbial diversity, these assemblages are robust and are not sensitive to major changes in salinity.

4. Discussion The salterns give a visual impression of a high-biomass system, particularly at the highest salinities in the crystalliser ponds where the water is highly pigmented due to the high biomass of D. salina. However, although the pigment concentrations were high, the rates of production and nutrient uptake were low at high salinity and most activity occurred at lower salinities. The data presented in this paper relate only to the water

column, but the ponds were shallow and have a high benthic^pelagic ratio. It is possible that, at least in the lower salinity ponds, some of the changes that were observed from day to day could have been the result of benthic activity. In particular, nutrient concentrations in the water could have been a¡ected by benthic microbial activity, and benthic grazers may have removed phytoplankton from the water column. However, the e¡ect of the benthos would have been considerably reduced at higher salinities. A hard crust developed on the sediment surface at s 15% salinity due to the deposition of gypsum; at the highest salinities, salt was crystallising, forming a layer of salt many centimetres deep. Benthic activity was probably greatly reduced under these conditions and con¢ned to micro-organisms. However, little is known about the activity of benthic mats in these systems [8] and it is not possible to determine the contribution of benthic processes to the observed changes in nutrients and pelagic biomass.

FEMSEC 1330 6-5-02

254

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

Fig. 7. E¡ect on pigment concentration of dilution of samples from the crystalliser pond (37% salinity). On the ¢rst day, water samples were diluted with sterile distilled water, contained within a 20-l sterile polycarbonate bottle which was incubated in situ in the crystalliser pond for 7 days. Samples were taken at V10:00 h each day for analysis of (a) chlorophyll a and (b) L-carotene.

4.1. Plankton production and grazing The data on carbon ¢xation rates show that high biomass did not equate with high productivity. The highest rates of primary production were measured at 8% salinity (Fig. 3) and there was also a signi¢cant enhancement of chlorophyll-speci¢c carbon ¢xation at this salinity. In the process of doubling the salt concentration from that of seawater, not only did chlorophyll concentration increase but more carbon ¢xation occurred per unit of chlorophyll. It is not clear why the chlorophyll-speci¢c ¢xation rate changed. It may have been a characteristic of the changing £ora or the light regime in the di¡erent ponds. The maximum phytoplankton diversity was found at 8% salinity, and the species composition changed from dominance by the autotrophic ciliate Myrionecta rubra ( = Mesodinium rubrum) in the 4% pond, through di¡erent £agellates, diatoms and some cyanobacteria at salinities of 8 and 11%, to total dominance by Dunaliella at salinities s 22% (M. Estrada, pers. comm.). There were di¡erences in water turbidity and penetration of light into the ponds which might have a¡ected chlorophyll content per cell. The di¡use attenuation coe⁄cients (k) were 0.97 (4% salt), 4.34 (8% salt), 2.03 (11% salt) and 1.09 m31 (37% salt). Even with these high attenuation coe⁄cients, it is unlikely that the phytoplankton in these ponds were signi¢cantly light-limited. The ponds were shallow and light would have penetrated to the sediment in all ponds. There was su⁄cient wind mixing and heating convection to turnover the shallow water column and there was no evidence of strati¢cation. So, it seems unlikely that the assemblages at 8%

salinity were light-limited. A change in chlorophyll-speci¢c carbon ¢xation may also result from nutrient e¡ects. The variability in many parameters between 21 and 25 May was unexpected. With no transfer of water between the ponds and only evaporation as the major factor in environmental change, it was assumed that each pond would show a high degree of temporal homogeneity. However, there were signi¢cant variations from day-to-day in nutrients, pigments and nutrient-uptake rates, with a 4^10fold increase in carbon ¢xation. Some of the carbon ¢xation was undoubtedly by organisms other than phytoplankton. Certainly the high dark ¢xation rates suggest that there was signi¢cant carbon ¢xation by non-photosynthetic bacteria such as chemolithotrophs, or that there were very high rates of anaplerotic CO2 ¢xation. It is also possible that some of the observed variability was not due to biological processes but were features of the ponds. These were large open spaces and changes in wind direction could have resulted in aggregation in di¡erent parts of the pond as was found for D. salina in the crystallisers. Samples were always taken from the same position in the pond, so heterogeneity within the ponds might explain the apparent variations in nutrient and chlorophyll concentration. However, there were also biological e¡ects. The variations in C:N uptake ratio (Fig. 4) suggest that there were changes in the community activity which could not be explained by assemblage heterogeneity. Clearly, a more extensive sampling programme is required to investigate how signi¢cant assemblage heterogeneity might be as an explanation for the observed variability. The measurements of nitrate and ammonium uptake (Table 2) showed that ammonium is the most important nitrogen source throughout the salinity gradient. The ambient ammonium concentration was not high in any pond and was about one order of magnitude less than that measured by Pedros-Alio¤ et al. [8] in the same saltern system in July 1993. It is likely that ammonium might accumulate during the year and, later in the season, they might have higher ammonium concentrations. However, there would have to be a change in the production/consumption ratio for that accumulation to occur. In May 1999, there was rapid ammonium turnover and the ambient concentration would have been totally consumed within a few days in all ponds. There must have been considerable heterotrophic activity to maintain this supply of ammonium. The experiments involving added nutrients found no evidence for ammonium or phosphate limitation (Fig. 5), indicating that there must be rapid recycling of nutrients across the salinity range and a general balance between supply and demand by the microbial £ora. Microzooplankton grazing could be an important source of nutrient recycling in the system [28]. Using chlorophyll a as an indicator of total phytoplankton biomass, we found that microzooplankton grazing could account for daily losses of up to 42% of phytoplankton standing stock (grazing rate of 0.541 day31 ) in the 4% pond. How-

FEMSEC 1330 6-5-02

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

ever, determination of grazing in the 11- and 37%-salinity ponds was problematic and statistically signi¢cant grazing rates (based on chlorophyll a) [29] were not found often in these dilution experiments. In the extreme environment of the crystalliser, this may indicate the absence of grazers, while in the 11% pond it might be explained by a nonsteady-state system in which the overall biomass was relatively stable but selective grazing a¡ected the community structure. It seems likely that the protozoa were very selective grazers. This can be inferred from the non-signi¢cant grazing rates on dino£agellates (characterised by peridinin) in all ponds, on fucoxanthin-containing organisms in the pond of 8% salinity and on alloxanthin-containing cells in the 4% pond. It is likely that selective grazing occurred on speci¢c groups of organisms along the salinity gradient, thus potentially in£uencing community diversity and nutrient re-cycling as a direct consequence of microzooplankton activity. 4.2. Nutrient cycling Nitrate utilisation was low in all but the highest-salinity pond. At 4% salinity, the measured nitrate uptake would have taken 12^23 days to deplete the ambient nitrate concentration; at 11 and 22% salinity, the estimated turnover time of nitrate was several hundred days. So at these salinities, nitrate was an insigni¢cant nitrogen source and pelagic activity should have caused little change in nitrate concentration. However, variations were observed between 21 and 25 May, particularly at 11% salinity where the nitrate concentration declined from 4.1 to 2.5 Wmol N l31 . This change was real, it could not have resulted from the activity of phytoplankton or bacteria within the water column and was most likely the result of benthic activity, which was not measured in this study. The highest nitrate-uptake rates measured in these experiments occurred in the crystalliser ponds at 37% salinity. The measured nitrate uptake would result in complete depletion of nitrate within 5^9 days and raises the interesting question as to the source of the nitrate. With ambient nitrate concentrations of between 5 and 7 Wmol N l31 (Fig. 2 and Table 2) and nitrate-uptake rates of V1 Wmol N l31 day31 , there must be a signi¢cant source of nitrate to the crystalliser ponds. But, this source is not obvious. It might result from the activity on nitrifying bacteria, utilising ammonium within the crystalliser. However, there is little evidence for nitrifying bacteria with such hypersaline systems. Indeed, Oren [30] argued that, since nitrifying bacteria gain so little energy from oxidation of ammonium and nitrite, that the additional energetic costs associated with halophilism might be too great and that one might not expect nitrifying bacteria to exist in saturated salt solutions. There are few reports of the presence of nitrifying bacteria in hypersaline environments. Ammonium oxidation has been measured in Mono Lake, CA, USA, a hypersaline, alkaline lake, by

255

Joye et al. [31] and the diversity of ammonium oxidising bacteria from the same lake is reported by Ward et al. [32]. However, the salinities involved were considerably less than this in this solar saltern. It is clear from the results presented here that an investigation would be warranted into the source of nitrate in the crystallisers and whether microbial oxidation of ammonium is possible in this hypersaline environment. Certainly, the microbial diversity study carried out by Benlloch et al. [10] as part of this study found a number of 16S sequences in the 22% pond which were closely related to the nitrite oxidiser, Nitrococcus mobilis. Of course, 16S phylogeny does not relate to function, although the nitrifying bacteria do tend to be a closely clustered group, so it is possible that the nitrifying bacteria were present, at least in the 22%-salt pond. It is clear from Fig. 4 that the uptake rates of carbon, nitrate, and ammonium were very variable on 21 and 25 May and there was no consistency in uptake ratio. In particular, daily estimates of molar uptake ratios did not conform to the Red¢eld ratio of 106C:16N [33]. The Red¢eld ratio is based on the carbon, nitrogen and phosphorus content of plankton, but the uptake ratio of carbon and nitrogen over time-scales less than the generation time of an organisms need not be the same as the Red¢eld ratio. However, when incubation experiments are long enough to encompass the generation time, a phytoplankton cell will assimilate carbon and nutrients in proportion to its elemental composition [34]. In other words, there can be uncoupling of nutrient uptake from cell growth, but when generation times are approximately the same as the incubation times for an experiment, the C:N uptake ratio often approximates to the Red¢eld ratio [35,36]. In the case of the lower salinity (4 and 11%) ponds, carbon and nitrogen uptake ratios in the light were not very di¡erent from the Red¢eld ratio of 6.6. At 4% salinity, C:N uptake ratios were 4.4 and 9.8, suggesting that there was excess nitrogen uptake on 21 May but less nitrogen assimilation on 25 May than would have been expected if the plankton composition had the Red¢eld ratio. This suggested that there may have been luxury consumption of nutrients, with the cells assimilating more nitrogen than they require for anabolism. At 11% salinity, the C:N uptake ratios in the light were 9.5 and 16; interestingly, the C:N uptake ratio in the dark was 6 and 5, again suggesting that some of the observed photosynthetic carbon ¢xation occurred when the phytoplankton cells already contained su⁄cient nitrogen for cell division. At the higher salinities, the C:N uptake ratios were extremely low. At 22% salinity, the C:N uptake ratio was V1 and at 37% salinity was as lower as V0.8; very similar ratios were measured in the incubations done in the dark. That is, at these high salinities, the nitrate and ammonium uptake was much higher than would be expected for the amount of photosynthetic carbon ¢xation. Why should this occur? The phytoplankton which dominated at these high salinities was D. salina. In order to be able to

FEMSEC 1330 6-5-02

256

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

grow in saturating salt solutions, Dunaliella must produce large quantities of organic osmolytes, which require very large amounts of metabolic energy. Glycerol is the major osmolyte of Dunaliella and Brown [37] suggests that the intercellular concentration of glycerol may be as high as 7.8 mol l31 . Since glycerol (C3 H8 O3 ) contains no nitrogen, it might be expected that, in order to synthesise su⁄cient glycerol to overcome the water stress, Dunaliella might have a high C:N uptake ratio. In fact, the opposite is the case and much more ammonium and nitrate is taken up (in both the light and dark). This observation has not been made before. However, we cannot assume that the nitrate and ammonium is taken up only by the primary producers and the prokaryote assemblage may be responsible for much of this nitrogen assimilation. The prokaryotes could have a high nitrogen demand for two reasons. Since high concentrations of glycerol are produced as compatible solute by Dunaliella, this could only be metabolised by heterotrophic organisms if nitrogen is also taken up. Alternatively, the organic solute utilised by prokaryotes to maintain osmotic balance in saturating salt may have a high nitrogen content. Oren [30] states that the main osmotic solute produced by heterotrophic halophilic bacteria is ectoine (1,4,5,6-tetrahydro-2-methyl-4-pyrimidine carboxylic acid) with a C:N ratio of 3. These questions can only be resolved by a further study which aims to distinguish between eukaryote and prokaryote ammonium and nitrate uptake. 4.3. Productivity and microbial diversity It is clear from this study that there were substantial changes along the salinity gradient in primary production, prokaryote production, nutrient assimilation and grazing. In particular, photosynthetic carbon ¢xation was greatly reduced at the highest salt concentrations. This decrease occurred at the same time as a great decrease in biodiversity of the primary producer which was e¡ectively reduced to one alga ^ Dunaliella ^ in the saturated salt pond. So, productivity appears to be linked to diversity of primary producers. However, there was much less variation in leucine uptake (a measure of prokaryote production) and in the uptake of nitrate and ammonium. As part of the same study, Benlloch et al. [10] found that the prokaryote diversity at salinities s 30 was as great as the diversity at seawater salinities. This was particularly true of the archaeal diversity, which was high at 4% and 30% salt but low at intermediate salinities. In this case, the archaea in the 30% salt were typical extremophile archaea ^ haloarchaea ^ but at 4% salt the archaeal assemblage consisted mostly of the group III marine archaea, whose function is unknown. There is clearly no simple relationship between productivity and diversity. Although, it could be argued that reduced diversity of the primary producers was linked to

declining primary production, this is certainly not true of the prokaryotic community. Hence, there is a need to relate community activity to the functional diversity of the microbial assemblage, which clearly changes with the stress imposed on the system by high salt but which is not closely related to prokaryote production.

Acknowledgements This study is partly funded by the European Union in the framework of the Mast 3 Programme, Contract number MAS3-CT97-0154 (MIDAS project), by the Plymouth Marine Laboratory, a component of the Natural Environment Research Council. We wish to acknowledge the invaluable assistance given to us by Dr Susana Benlloch during the experiments in Alicante in May 1999 and we thank Andy Rees for mass spectrometric analysis of the 15 N samples. We also thank Mr Miguel Cuervo-Arango for permission to sample the salterns at Santa Pola. Merete Allerup, Susanne Hemmingsen and Winnie Martinsen are acknowledged for technical assistance.

References [1] Naeem, S., Thompson, L.J., Lawler, S.P., Lawton, J.H. and Wood¢n, R.M. (1994) Declining biodiversity can alter the performance of ecosystems. Nature 368, 734^737. [2] Grime, J.P. (1997) Ecology ^ Biodiversity and ecosystem function : the debate deepens. Science 277, 1260^1261. [3] Lehman, C.L. and Tilmann, D. (2000) Biodiversity, stability, and productivity in competitive communities. Am. Nat. 156, 534^552. [4] Rodr|¤guez-Valera, F. (1988) Characteristics and microbial ecology of hypersaline environments. In: Halophilic Bacteria, Vol. 1 (Rodr|¤guez-Valera, F., Ed.), pp. 30. CRC Press, Boca Raton, FL. [5] Benlloch, S., Mart|¤nez-Murcia, A.J. and Rodr|¤guez-Valera, F. (1995) Sequencing of bacterial and archaeal 16S rRNA genes directly ampli¢ed from a hypersaline environment. Syst. Appl. Microbiol. 18, 574^581. [6] Benlloch, S., Acinas, S.G., Martinez-Murcia, A.J. and RodriguezValera, F. (1996) Description of prokaryotic biodiversity along the salinity gradient of a multipond solar saltern by direct PCR ampli¢cation of 16S rDNA. Hydrobiologia 329, 19^31. [7] Anton, J., Rossello-Mora, R., Rodriguez-Valera, F. and Amann, R. (2000) Extremely halophilic Bacteria in crystallizer ponds from solar salterns. Appl. Environ. Microbiol. 66, 3052^3057. [8] Pedro¤s-Alio¤, C., Caldero¤n-Paz, J.I., MacLean, M.H., Medina, G., Marrase¤, C., Gasol, J.M. and Guixa-Boixereu, N. (2000a) The microbial food web along salinity gradients. FEMS Microbiol. Ecol. 32, 143^155. [9] Pedro¤s-Alio¤, C., Caldero¤n-Paz, J.I. and Gasol, J.M. (2000b) Comparative analysis shows that bacterivory, not viral lysis, controls the abundance of heterotrophic prokaryotic plankton. FEMS Microbiol. Ecol. 32, 157^165. [10] Benlloch, S., Lo¤pez-Lo¤pez, A., Yvreafis, L., Goddard, V., Casamayor, E.O., Daae, F.L., Smerdon, G., Massana, R., Joint, I., Thingstad, F., Pedro¤s-Alio¤, C. and Rodr|¤guez-Valera, F. (2002) Prokaryotic diversity throughout the salinity gradient of a solar saltern. FEMS Microbiol. Ecol., in preparation. [11] Wright, S.W., Je¡rey, S.W., Mantoura, R.F.C., Llewellyn, C.A., Bjornland, T., Repeta, D. and Welschmeyer, N. (1991) Improved

FEMSEC 1330 6-5-02

I. Joint et al. / FEMS Microbiology Ecology 39 (2002) 245^257

[12]

[13] [14] [15]

[16]

[17]

[18] [19]

[20]

[21]

[22]

[23] [24]

HPLC method for the analysis of chlorophylls and carotenoids from marine phytoplankton. Mar. Ecol. Prog. Ser. 77, 183^196. Schlu«ter, L. and Havskum, H. (1997) Phytoplankton pigments in relation to carbon content in phytoplankton communities. Mar. Ecol. Prog. Ser. 155, 55^65. Brewer, P.G. and Riley, J.P. (1965) The automatic determination of nitrate in seawater. Deep-Sea Res. 12, 765^772. Grassho¡, K. (1976) Methods of Seawater Analysis. Verlag Chemie, Weinheim. Mantoura, R.F.C. and Woodward, E.M.S. (1983) Optimisation of the indophenol blue method for the automated determination of ammonia in estuarine waters. Estuarine Coast. Shelf Sci. 17, 219^224. IOC, Paris (France) (1994) Protocols for the Joint Global Ocean Flux Study (JGOFS) Core Measurements. Manuals Guides IOC 29, 126 pp. Lazar, B., Starinsky, A., Katz, A. and Sass, E. (1983) The carbonate system in hypersaline solutions: alkalinity and CaCO3 solubility of evaporated seawater. Limnol. Oceanogr. 28, 978^986. Burke, C.M. and Atkinson, M.J. (1988) Measurement of total alkalinity in hypersaline waters: values of fH . Mar. Chem. 25, 49^55. Whit¢eld, M. (1979) Activity coe⁄cients in natural waters. In: Activity Coe⁄cients in Electrolyte Solutions, Vol. II (Pytkowicz, R.M., Ed.), pp. 153^299. CRC Press, Boca Raton, FL. Owens, N.J.P. and Rees, A.P. (1989) Determination of nitrogen-15 at submicrogram levels of nitrogen using automated continuous-£ow isotope ratio mass spectrometry. Analyst 114, 1655^1657. Dugdale, R.C. and Goering, J.J. (1967) Uptake of new and regenerated forms of nitrogen in primary productivity. Limnol. Oceanogr. 12, 196^206. Smith, D.C. and Azam, F. (1992) A simple economical method for measuring bacterial protein synthesis rates in sea water using 3 Hleucine. Mar. Microb. Foodwebs 6, 107^114. Landry, M.R. and Hassett, R.P. (1982) Estimating the grazing impact of marine micro-zooplankton. Mar. Biol. 67, 283^288. Landry, M.R. (1993) Estimating rates of growth and grazing mortality of photoautotrophic plankton by dilution. In: Handbook of Methods in Aquatic Microbial Ecology (Kemp, P.F., Sherr, B.F. Sherr, E.B. and Cole, J.J., Eds.), pp. 715^722. Lewis Publishers, London.

257

[25] Schlu«ter, L. (1998) The in£uence of nutrient addition on growth rates of phytoplankton groups, and microzooplankton grazing rates in a mesocosm experiment. J. Exp. Mar. Biol. Ecol. 228, 53^71. [26] Landry, M.R., Kirshtein, J. and Constantinou, J. (1995) A re¢ned dilution technique for measuring the community grazing impact of microzooplankton, with experimental tests in the central equatorial Paci¢c. Mar. Ecol. Prog. Ser. 120, 53^63. [27] Je¡rey, S.W., Mantoura, R.F.C. and Wright, S.W. (1997) Phytoplankton Pigments in Oceanography. UNESCO Publishing, Paris. [28] Goldman, J.C., Caron, D.A., Andersen, O.K. and Dennett, M.R. (1985) Nutrient cycling in a micro£agellate food chain: I. Nitrogen dynamics. Mar. Ecol. Prog. Ser. 24, 231^242. [29] Murrell, M.C. and Hollibaugh, J.T. (1998) Microzooplankton grazing in northern San Francisco Bay measured by the dilution method. Aquat. Microb. Ecol. 15, 53^63. [30] Oren, A. (1999) Bioenergetic aspects of halophilism. Microb. Mol. Biol. Rev. 63, 334^349. [31] Joye, S.B., Connell, T.L., Miller, L.G., Oremland, R.S. and Jellison, R.S. (1999) Oxidation of ammonia and methane in an alkaline, saline lake. Limnol. Oceanogr. 44, 178^188. [32] Ward, B.B., Martino, D.P., Diaz, M.C. and Joye, S.B. (2000) Analysis of ammonia-oxidizing bacteria from hypersaline Mono Lake, California, on the basis of 16S rRNA sequences. Appl. Environ. Microbiol. 66, 2873^2881. [33] Red¢eld, A.C., Ketchum, B.H. and Richards, F.A. (1963) The in£uence of organisms on the composition of sea water. In: The Sea, Vol. 2 (Hill, M.N., Ed.), pp. 26^77. Wiley, New York. [34] Thingstad, T.F., Skjoldal, E.F. and Bohne, R.A. (1993) Phosphorus cycling and algal-bacterial competition in Sandsfjord, western Norway. Mar. Ecol. Prog. Ser. 99, 239^259. [35] Rees, A.P., Joint, I. and Donald, K.M. (1999) Early spring bloom phytoplankton-nutrient dynamics at the Celtic Sea Shelf Break. Deep-Sea Res. I 46, 483^510. [36] Donald, K.M., Joint, I., Rees, A.P., Woodward, E.M.S. and Savidge, G. (2001) Uptake of carbon, nitrogen and phosphorus by phytoplankton along the 20‡W meridian the NE Atlantic between 56.5‡N and 37‡N. Deep-Sea Res. II 48, 873^897. [37] Brown, A.D. (1990) Microbial Water Stress Physiology; Principles and Perspectives. John Wiley and Sons, Chichester.

FEMSEC 1330 6-5-02