Nitrogen fixation by Baltic cyanobacteria is adapted to the prevailing photon flux density

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1 RESEARCH New Phytol. (), 147, Nitrogen fixation by Baltic cyanobacteria is adapted to the prevailing photon flux density A. M. EVANS, J. R. GALLON*, A. JONES, M. STAAL, L.J.STAL, M. VILLBRANDT AND T. J. WALTON Biochemistry Research Group, School of Biological Sciences, University of Wales, Swansea, Singleton Park, Swansea SA2 8PP, UK Netherlands Institute for Ecology, Centre for Estuarine and Coastal Ecology, PO Box 14, NL-44 AC Yerseke, The Netherlands University of Bremen, Faculty 2, Leobener StrasseNW2, D-2839 Bremen, Germany Received 2 August 1999; accepted March SUMMARY N fixation, measured as acetylene reduction, was studied in laboratory cultures and in natural assemblages (both as a mixed population and as individually picked colonies) of the heterocystous cyanobacteria Aphanizomenon sp. and Nodularia spp. from the Baltic Sea. During a diurnal cycle of alternating light and darkness, these organisms reduced acetylene predominantly during the period of illumination, although considerable activity was also observed during the dark period. In both laboratory cultures and natural populations N fixation was saturated below a photon flux density of 6 µm s. In cyanobacterial blooms in the Baltic Sea, nitrogenase activity was mostly confined to the surface layers. Samples collected from greater depths did not possess the same capacity for acetylene reduction as samples from the surface itself, even when incubated at the photon flux density prevailing in surface waters. This suggests that, with respect to N fixation, Baltic cyanobacteria are adapted to the intensity of illumination that they are currently experiencing. Key words: Baltic cyanobacteria, N fixation, Aphanizomenon, Nodularia. INTRODUCTION Most microorganisms depend on a source of combined nitrogen (N) such as ammonium (NH +)or nitrate (NO ) in order to synthesize proteins, nucleic acids and other nitrogenous cell constituents. However, in most environments such sources of N are in short supply, largely as a result of the activity of denitrifying bacteria. Although N is abundant in the atmosphere, where it occurs as N, relatively few organisms (diazotrophs) are able to use it in this form (Gallon, 1992). Diazotrophs possess the enzyme nitrogenase which catalyses the reduction, or fixation, of N to NH +. However, because the two N atoms in N are linked by a stable triple bond, N fixation involves diazotrophs in a considerable energy commitment (as ATP hydrolysis and the generation of low-potential reducing equivalents) that must be met metabolically. Many cyanobacteria can fix N. As phototrophs, cyanobacteria are well equipped to supply nitro- *Author for correspondence (fax ; j.r.gallonswansea.ac.uk). genase with ATP and low-potential electrons that are derived ultimately from photosynthesis (Fay, 1992). However, cyanobacterial photosynthesis also generates O, a potent inhibitor of N fixation (Gallon, 1992). Many N -fixing cyanobacteria therefore separate the O -sensitive process of N fixation from oxygenic photosynthesis by confining nitrogenase in specialized cells known as heterocysts. Heterocysts lack some of the components of photosystem II (Tel-Or & Stewart, 1977) and consequently do not evolve O. The envelope of these cells also forms a diffusion barrier that limits entry of atmospheric O (Walsby, 198). However, because they are defective in photosynthesis, heterocysts depend upon adjacent, undifferentiated vegetative cells for a supply of fixed carbon in order to support heterotrophic metabolism, including N fixation. Among cyanobacteria, heterocystous species are generally considered best equipped for growth with N as an N source (Stal et al., 1994). They are widely distributed in nature, occurring in terrestrial and freshwater environments (Fay, 1983). Because growth of N -fixing cyanobacteria is not limited by

2 286 RESEARCH A. M. Evans et al. availability of N, dense blooms of heterocystous cyanobacteria can develop in lakes, especially when there is an abundant supply of phosphorus (Howarth et al., 1988). Although the marine environment is generally considered to be chronically N-depleted, heterocystous cyanobacteria are almost completely absent (Paerl, 199). By contrast, the major freeliving diazotroph in the open ocean is the nonheterocystous cyanobacterium Trichodesmium (Carpenter & Capone, 1992). Notable exceptions are two large, brackish seas, the Baltic Sea in northern Europe (Kahru et al., 1994; Leppa nen et al., 199; Walsby et al., 199; Kononen et al., 1996; Lehtimaki et al., 1997; Stal & Walsby, 1998) and the Peel Harvey estuary in Western Australia (Huber, 1986). The Baltic Sea is an enclosed, shallow sea (average depth 6 m) with a surface area of 374 km (Leppa nen et al., 1988). It is subdivided into a number of areas: the Gulf of Bothnia, the Gulf of Finland, the Gulf of Riga, the Baltic proper, the Gotland Sea, the Middle Bank, the Bornholm Sea and the Kiel Bight. It is connected via narrow and shallow channels to the Kattegat and Skagerak, that lead to the North Sea, and tides are virtually absent. Salinity in the Baltic Sea ranges from almost freshwater (1 4 practical salinity units, PSU) in the eastern and northern parts, to 7 8 PSU in the south. The Baltic Sea is characterized by dense blooms of cyanobacteria that occur during a short period in summer (Kahru et al., 1994). These are composed mainly of the heterocystous cyanobacteria Nodularia spp., Aphanizomenon sp. and Anabaena sp. (Walsby et al., 199; Wasmund, 1997). Because they contain gas vesicles, these cyanobacteria form colonies or larger aggregates that float to the water surface where they accumulate as dense scums (Walsby et al., 199, 1997). Despite the fact that these cyanobacterial blooms occur locally and for relatively short periods (sometimes only weeks), rough estimates suggest that they fix between and 13 tonnes of N yr (Larsson et al., 198), equivalent to 1 2% of the total N input into the Baltic Sea (Leppa nen et al., 1988). However, data are incomplete and do not allow a precise estimation of the amount of N fixation in the Baltic Sea. The main aim of the work described here was to investigate the diurnal patterns and vertical depth variation of N fixation in individually picked colonies and mixed colony samples of natural populations of Aphanizomenon sp. and Nodularia spp. from the Baltic Sea. In particular, the patterns observed in situ have been compared with those seen in laboratory cultures incubated under 12 h light:12 h darkness. The responses of N fixation to photon flux density have also been studied in laboratory cultures and natural populations of Baltic cyanobacteria. An understanding of the influence that such environmental factors have on N fixation in cyanobacterial blooms is essential for an accurate assessment of the role of cyanobacteria in N cycling in the Baltic Sea. MATERIALS AND METHODS Sample collection Samples were collected in July and August during research cruises aboard the R.V. Alkor (Institut fu r Meereskunde an der Universita t Kiel, Kiel, Germany) during at drift stations in the southern Middle Bank and Bornholm Sea regions, where dense cyanobacterial blooms dominated by either Aphanizomenon sp. or Nodularia spp. were observed. Phytoplankton samples were collected in a -µm mesh plankton net (Hydrobios, Kiel, Germany) taken from the upper mixed layer (usually m in depth) and resuspended in 2 ml of seawater. Alternatively, when samples from a particular depth were required, collection was by means of a conductivity, temperature and depth (CTD)- rosette sampler (General Oceanics, Miami, FL, USA) followed by filtration through a -µm plankton net in order to concentrate the plankton suspension by a factor of. The CTD provided continuous measurement of temperature, salinity, in vivo fluorescence of chlorophyll, oxygen and ph. A quantameter and spectroadiometer were employed to determine light distribution with depth. At each drift station the attenuation coefficient was calculated so that the amount of photosynthetically active radiation penetrating to a particular depth could be established. Hydrography and chemical analyses The weather was relatively calm and sunny during the cruise periods, with minimum mixing of surface water layers. Generally the thermocline was found at a depth of 3 m, with the halocline at m. The largest concentration of chlorophyll a, corresponding to the N -fixing cyanobacterial colonies, was found in the upper m. Nutrient analyses were Table 1. Sampling conditions on 31 July 1 August 1993, 11 August 199, 1 16 August 199 and 19 August 1996 at drift stations in the Southern Middle Bank and Bornholm Sea Date Position Sample time (hours) N : E N 9: E N 8: E N 1: E Surface temperature (C)

3 RESEARCH N fixation by Baltic cyanobacteria 287 performed according to the methods listed in Grasshoff (1976). Growth and maintenance of laboratory cultures Aphanizomenon (Morren) sp., originally isolated from the Gulf of Finland, was kindly provided by Dr Kaarina Sivonen, University of Helsinki, Finland. Morphological and fine structure studies of Aphanizomenon collected from the Gotland Deep region of the Baltic Sea have shown that Baltic Aphanizomenon is considerably different from species isolated from freshwater environments, notably Aphanizomenon flos-aquae (L.) Ralfs ex Born. et Flah. (Janson et al., 1994). The strain used in this study is therefore referred to as Aphanizomenon sp. Cultures of Nodularia (Mertens) SN34D and Nodularia SN1A were kindly provided by Drs Paul Hayes and Gary Barker, School of Biological Sciences, University of Bristol, UK. They were obtained from clonal isolates of single colonies of Nodularia spumigena Mertens ex Born. et Flah. from the Baltic Sea. Stock cultures of Aphanizomenon sp. were maintained in ASM-1 medium free of combined N (Gallon et al., 1978) whereas Nodularia SN34D and Nodularia SN1A were grown in N-free MN medium (Van Baalen, 1962), modified according to Gallon et al. (1991). Cultures were maintained with shaking at 17C and illuminated at 3 µmol m s under 12 h light:12 h darkness. For experimental analysis, large volumes (up to 1 l) were grown in bottles vigorously aerated with sterile air at.3 l min l culture. Unless stated otherwise, all laboratory experiments were carried out with cultures in mid-exponential growth phase and incubated at C under 12 h light (9 µmol m s ) and 12 h darkness. Sample incubation For assay of N fixation, 2 3 ml of laboratory culture or mixed population bloom were transferred directly into 8.9-ml Teflon-sealed screw-top vials or -ml crimp-top vials. Alternatively, colonies of Aphanizomenon sp. and Nodularia spp. ( per assay vial) were individually picked from natural populations with disposable plastic inoculating loops and placed into vials containing 3 ml of filtered (Whatman GFF, 47 mm) seawater. For samples collected with the plankton net, typically 1 h elapsed between sample collection and initiation of the experiment. The additional need to concentrate samples collected from the CTD rosette sampler meant that 2 h usually elapsed before initiation of the experiment in these cases. During this period, samples were incubated on board under conditions simulating in situ illumination and temperature. For experimental analysis, assay vials were incubated in a water bath on deck using neutral density filters where appropriate in order to provide in situ conditions of illumination. For incubation in the dark, vials were wrapped in several layers of aluminium foil. The incubation temperature corresponded to that of surface seawater (Table 1). Measurement of nitrogenase activity N fixation was measured using the acetylene reduction assay (Gallon et al., 1993). During the first cruise (1993), acetylene and ethylene were measured by gas chromatography using a Chrompack CP9 gas chromatograph equipped with a flame ionization detector (FID) and a 2.3 mm wide bore column of Poraplot Q (Chrompack, Bergen op Zoom, The Netherlands). The temperature settings were, 9 and C for oven, injector and detector, respectively. N was used as carrier gas at a flow of 1 ml min. The flow rates of H and air were and ml min, respectively. Acetylene was used as an internal standard (Stal, 1988). Cyanobacterial samples (2 ml) were transferred into -ml Chrompack glass vials and, after sealing with an aluminium crimp and Teflon seal, 2 ml of acetylene was injected. This gave a concentration of acetylene in the gas phase of 2% (by vol). At the end of the incubation period a gas sample was taken using a -ml disposable syringe and subsequently transferred into a -ml evacuated Chrompack vial for storage until analysis in the laboratory. Analysis was carried out within 1 2 wk after the cruise. No significant loss of gas occurred during this period. Small losses were automatically corrected for by the use of acetylene as an internal standard. During 199 and 1996, N fixation was measured on board by addition of 1 ml of acetylene to the gas phase (giving 14 % vv). After 6 min incubation, a 1-ml sample was removed and ethylene production measured using either a Pye Unicam GCD gas chromatograph (FID, Philips PU48 computing integrator) or a Shimadzu GC-8A gas chromatograph (FID, Shimadzu C-R6A computing integrator), both equipped with a 2 m3 mm (ID) stainless steel column packed with Poropak N at 1C. High-purity N was used as the carrier gas, and the amount of ethylene produced was by reference to a standard consisting of nmol of ethylene. Determination of chlorophyll a, protein and carbohydrate At the end of the incubation period the sample was filtered (Whatman GFC, 2 mm) and extracted overnight with 96% (vv) ethanol in the dark at ambient temperature. Absorption was measured at 66 nm and an absorbance coefficient of 72.3 ml mg cm was used to calculate the concentration of chlorophyll a.

4 288 RESEARCH A. M. Evans et al. Protein was measured by the Folin method as modified by Bailey (1962). Samples from laboratory cultures were passed through a French pressure cell at 138 MPa and centrifuged for 2 min at 13 g in order to sediment the cell debris. The protein content of.1 ml of supernatant was then measured. Samples from natural populations were filtered onto GFC filters (Whatman) which were immediately frozen and transported back to the laboratory in dry ice. Filters were boiled in 3 ml 2% (wv) Na CO in.1 M NaOH and, following centrifugation,.1 ml was removed for measurement of protein. In the case of individually sorted colonies of Nodularia spp. it proved impossible to express nitrogenase activity in terms of either chlorophyll or protein as a biomass marker, as the amount per assay ( colonies) was below the limits of detection in each case. Nitrogenase activity is therefore expressed as nmol h per colonies for these samples. Total carbohydrate was measured by the method of Hodge & Hofreiter (1962), whereas intracellular glucan was measured as described by Ernst et al (1984). Immunogold localization of nitrogenase Transmission electron microscopy (TEM)-immunogold labelling of the Fe protein of nitrogenase was generously performed by Professor Birgitta Bergman and collaborators, Department of Botany, University of Stockholm, Sweden. Samples (. l) were concentrated to. ml packed volume by centrifugation and placed in 2.% (wv) glutaraldehyde in sodium cacodylate buffer ( mm Na(CH ) AsO 3H O, 4.1 mm HCl ph 7.2) for 1 h. Each sample was washed twice for a minimum of min in 1 vol sodium cacodylate buffer and 2 vol Palade buffer (7. mm sodium acetate, 2 mm HCl, 6 mm sodium barbital). The pellet was then mixed with 2% (vv) Oxoid agar dissolved in deionized water and kept at C in % (vv) ethanol for 1 min, after which it was serially dehydrated by placing in % (vv) ethanol for 1 min followed by 2 4 h in 7% ethanol. Samples were then sent to the University of Stockholm where they were dehydrated in absolute ethanol and embedded in LR White resin (Taab, Reading, UK). Ultra-thin sections were placed on coated gold grids and treated with antisera (Stal & Bergman, 199). The primary antiserum (raised in rabbits against the Fe protein of nitrogenase purified from Rhodospirillum rubrum) was a gift from Dr Stefan Nordlund, Department of Biochemistry, University of Stockholm, and was used at a dilution of 1:3. The secondary antibody was goat anti-rabbit IgG conjugated to nm gold spheres and diluted 1:. Specimens were viewed in a Zeiss electron microscope operating at 8 kv, at a magnification of A set of four micrographs was obtained from each sample. RESULTS Effect of light and photosynthesis on nitrogenase activity In July 1993, natural populations of cyanobacteria from the Bornholm Sea showed substantial nitrogenase activity (measured as acetylene reduction) when assayed during the day. After incubation for 1 h in the presence of µm 3,-(3,4-dichlorophenyl)- 1,1-dimethylurea (DCMU), an inhibitor of oxygenic photosynthesis, activity decreased to 7127% (SE, six determinations) of that in uninhibited cells. Nitrogenase activity could also be measured in daytime samples incubated in the dark, though at only 41% (SE, six determinations) of the rate seen in the light. In the short term, therefore, N fixation by Baltic cyanobacteria is not wholly dependent on active photosynthesis. This is consistent with the situation in certain other heterocystous cyanobacteria (Bottomley & Stewart, 1977; Khamees et al., 1987), though not in all strains (Gallon, 198). Nevertheless, it is likely that illumination is needed in order to provide sufficient ATP and reductant for N fixation to proceed at maximum rates, although catabolism of endogenous storage carbohydrate might support a lower rate of N fixation in the dark. Diurnal variation in nitrogenase activity In samples containing a mixed population of cyanobacteria, obtained from the Bornholm Sea (31 July 1 Aug 1993; 19 Aug 1996) and Middle Bank regions ( 11 Aug 199), nitrogenase activity demonstrated a clear light dependency (Fig. 1). The patterns of N fixation obtained in 1993, 199 and 1996 were similar, with highest activities occurring between 8: and 1: hours local time, and lowest activity during the period of darkness. Addition of µm DCMU for 1 h slightly decreased nitrogenase activity in samples taken during the light period, but had no effect on the activity of samples taken and assayed in the dark (Fig. 2a). This implies that the small decrease in nitrogenase activity seen when DCMU is added in the light is related to its known effect on photosynthesis, and not to any direct inhibition of N fixation. Transfer to darkness of samples collected during the period of daylight markedly decreased the nitrogenase activity normally seen, presumably as a result of severe limitations in the supply of ATP and reductant for N fixation (Fig. 2b). Nevertheless, nitrogenase activity in the dark was generally greater in samples collected during the daytime than in those collected at night. A notable exception, however, was at 7: hours, shortly after sunrise, when the rate of acetylene reduction measured in samples incubated in the dark was virtually identical to that seen in the night. This

5 RESEARCH N fixation by Baltic cyanobacteria (a) 3 (a) C2H4 produced (µmol h 1 mg 1 Chla) 3 3 (b) (c) C2H4 produced (µmol h 1 mg 1 Chla) 3 (b) 8: 12: 16: : : 4: 8: 12: Local time (hours) Fig. 2. Effects of (a) µdcmu and (b) darkness on the diurnal variation of nitrogenase activity in Baltic cyanobacteria, measured 31 July 1 August : 12: 16: : : 4: 8: 12: Local time (hours) Fig. 1. Diurnal variation of nitrogenase activity in Baltic cyanobacteria, sampled (a) 31 July 1 August 1993 in the Bornholm Sea, (b) 11 August 199 in the Middle Bank region of the Baltic Sea and (c) 19 August 1996 in the Bornholm Sea. Samples were incubated at the temperature and photon flux density prevailing at the sea surface. Values are the mean of three independent measurements; error bars show SD where this was greater than the size of the symbol. C2H4 produced (µmol h 1 mg 1 Chla) 3 (a) contrasts with other samples taken during the day, and suggests that endogenous pools of carbohydrate rapidly become depleted at night. In consequence, cyanobacteria collected during the main part of the day have more reserves available to support nitrogenase activity in the dark than do samples collected at night or early in the morning. During the period 19 August 1996, samples of the mixed cyanobacterial population were sorted into colonies of Aphanizomenon sp. and Nodularia spp. Acetylene reduction by colonies of Aphanizomenon sp. was maximal during the day, peaking at 17.9 µmol h mg Chla. The mean value of activity at night was. µmol h mg Chla, corresponding to 31% of the maximum daytime rate (Fig. 3a). Unlike Aphanizomenon sp., sorted colonies of Nodularia spp. (Fig. 3b) showed little difference in acetylene reduction between day (average activity.73 nmol h per colonies) and night (average activity.71 C2H4 produced (nmol h 1 per colonies) (b) 8: 12: 16: : : 4: 8: 12: Local time (hours) Fig. 3. Diurnal variation of nitrogenase activity in individually sorted colonies ( per assay) of (a) Aphanizomenon sp. and (b) Nodularia spp. collected from a natural population in the Bornholm Sea 19 August Samples were incubated at the temperature and photoflux density prevailing in surface waters. Values are the mean of three independent measurements; error bars show SD where this was greater than the size of the symbol. Note the different scales and units of nitrogenase activity for the two cyanobacteria.

6 29 RESEARCH A. M. Evans et al. C2H4 produced (µmol h 1 mg 1 Chla) 1 1 (a) (b) D L2 L6 L D2 D6 D L2 Sampling time (h after transfer to the light or dark period) Fig. 4. Pattern of acetylene reduction (N fixation) in laboratory cultures of (a) Aphanizomenon sp. and (b) Nodularia SN34D during incubation at C under 12 h light (9 µmol m s ):12 h dark. Sampling times are represented as hours after transfer to the light or dark period, thus L2 refers to samples taken 2 h after the onset of illumination, while D6 refers to samples taken 6 h into the dark period. Values are the mean of three independent measurements; error bars show SD where this was greater than the size of the symbol. nmol h per colonies). The pattern of activity seen in the mixed population (Fig. 1c) resembled that of Aphanizomenon more closely than that of Nodularia. N fixation by laboratory cultures of Aphanizomenon sp. and Nodularia SN34D When grown under 12 h light:12 h dark, laboratory cultures of Aphanizomenon sp. showed a pattern of acetylene reduction (Fig. 4a) closely resembling that previously reported for heterocystous cyanobacteria (Mullineaux et al., 1981; Khamees et al., 1987). The maximum rate of acetylene reduction, seen in the light phase, was. µmol h mg Chla, similar to that observed in individually picked colonies of Aphanizomenon sp. selected from a natural population (Fig. 3a.). Nitrogenase activity in the dark was c. 33% of that obtained in the light. In laboratory cultures of Nodularia SN34D, rates of nitrogenase activity were also greatest during the light period, but significant activity (c. 68% of that seen in the light) was consistently observed during the period of darkness (Fig. 4b). In this regard, Nodularia SN34D resembles heterocystous cyanobacteria such as Scytonema javanicum that have Carbohydrate (µg µg 1 protein) D L2 L6 L D2 D6 D L2 Sampling time (h after transfer to the light or dark period) Fig.. Fluctuations in the carbohydrate content (relative to that of protein) for laboratory cultures of Aphanizomenon sp. (circles) and Nodularia SN34D (squares) during incubation at C under 12 h light (9 µmol m s ): 12 h dark. Sampling times are represented as hours after transfer to the light or dark period, thus L2 refers to samples taken 2 h after the onset of illumination, while D6 refers to samples taken 6 h into the dark period. Values are the mean of three independent measurements; error bars show SD where this was greater than the size of the symbol. previously been reported to fix N in the dark at rates similar to those seen in the light (Khamees et al., 1987). The pattern of nitrogenase activity in laboratory cultures of Nodularia is similar to that seen in natural populations of Nodularia (Fig. 3b). The fact that both Aphanizomenon and Nodularia continue to fix N during the period of darkness implies that light-independent processes can generate sufficient ATP and reductant to support nitrogenase activity in these cyanobacteria. In laboratory cultures grown under alternating light and darkness, the total carbohydrate content (relative to that of protein) was higher in Nodularia SN34D throughout the diurnal cycle than in Aphanizomenon sp. and showed a greater fluctuation, reaching a maximum at the end of the light period and decreasing at the onset of darkness (Fig. ). TEM was performed on samples of Aphanizomenon removed from laboratory cultures simultaneously with the samples taken for acetylene reduction; they were then immunolabelled with gold in order to reveal the Fe protein of nitrogenase. This procedure showed that the Fe protein was confined to heterocysts, that it was present throughout the cycle of 12 h light and 12 h darkness, and that there was no apparent difference in the intensity of labelling between samples collected from the light phase and those collected during the period of darkness (Fig. 6). Dependence of nitrogenase activity on prevailing irradiance In natural populations of Baltic cyanobacteria, nitrogenase activity increased with increased photon

7 RESEARCH N fixation by Baltic cyanobacteria 291 (a) (b) Fig. 6. Immunogold labelled Fe protein of nitrogenase in heterocysts of laboratory cultures of Aphanizomenon sp. Samples were taken (a) h into the light phase (L) and (b) 6 h into the dark phase (D6). Note that the vegetative cells (left) adjacent to the heterocysts contain only background amounts of immunogold-labelled nitrogenase. Scale bars, 1 µm. flux density up to around 6 µmol m s (Stal & Walsby, 1998). Addition of µm DCMU did not substantially alter this pattern (data not shown). In both cases, nitrogenase activity was inhibited above µmol m s. This implies that the inhibitory effect of intense illumination on N fixation by Baltic cyanobacteria is not related to excessive production of O. Laboratory cultures of Aphanizomenon sp., Nodularia SN34D and Nodularia SN1A responded in a similar way to natural populations (Fig. 7). In all three cultures, nitrogenase activity was saturated at a photon flux density of around 6 µmol m s and consistently inhibited above µmol m s. In order to study the effect of varying photon flux density on nitrogenase activity, samples were collected on 9 August 199 from the surface, and from depths of 3 and m. Unfortunately, cloudy conditions meant that the ambient photon flux density was relatively low, with a mean surface irradiance for the incubation period of 6 µmol m s. During the period of acetylene reduction, neutral density filters were used to simulate photon flux densities corresponding to depths of, 3, 8,, 1 and m (6, 281, 14, 37, and 4 µmol m s, respectively). Even when incubated at surface photon flux densities, samples collected from 3 and m were less effective at N fixation than were samples from the surface itself. Maximum rates of acetylene reduction by samples from 3 and m (.8 µmol h mg Chla and.6 µmol h mg Chla, respectively) were only c. 3% of maximum rates observed

8 292 RESEARCH A. M. Evans et al. C2H4 produced (µmol h 1 mg 1 Chla) (a) (b) (c) Photon flux density (µmol m 2 s 1 ) Fig. 7. Effect of increased illumination on the rate of acetylene reduction (N fixation) by laboratory cultures of (a) Aphanizomenon sp., (b) Nodularia SN34D and (c) Nodularia SN1A. in samples from the surface (1.6 µmol h mg Chla). Moreover, whereas the rate of acetylene reduction in cyanobacteria sampled either from the surface or from 3 m was maximal at 6 µmol m s, the rate in cyanobacteria sampled from m was only slightly stimulated by photon flux densities 37 µmol m s. Colonies sampled at m therefore appeared to have adapted to the intensity of illumination prevailing at that depth. Depth distribution of nitrogenase activity Diurnal variation in the specific activity of nitrogenase in samples collected from a mixed population at different depths in the water column was measured August 199 in the Middle Bank region of the Baltic Sea. Samples were collected at the surface and at depths of 3, 6, 9, 12 and 1 m, and were then incubated on deck at the temperature and photon flux density prevailing in surface waters. Thus, except for the surface sample, the measured rates of N fixation do not represent the rates that would occur in situ. Rather, they reflect the capacity for N fixation in populations at different depths after transfer to the conditions existing at the surface of the water column. Except during the period of darkness (samples taken at 22: and 2: hours), the specific activities measured in samples from below the surface were consistently lower than those measured in samples taken at the surface (Fig. 8), supporting the idea that populations might adapt to the conditions prevailing at the depth at which they find themselves. In samples taken from the top 9 m of the water column, the specific activity of nitrogenase was greatest during the day and very low at night. These data all imply that light is a major factor regulating N fixation by Baltic cyanobacteria. The fact that, above 1 m, significant rates of N fixation could be measured during the night implies that heterotrophic metabolism can support N fixation. This might be particularly important in samples at greater depths where the period of effective illumination could be quite short. The daily integral of the capacity for N fixation derived from these studies shows that maximum specific activity of nitrogenase occurs close to the surface and decreases markedly with depth (Fig. 9). Measurement of the diurnal variation of specific nitrogenase activity with depth was repeated 1 16 August 199 in the Bornholm Sea, except that neutral density filters were employed so that (within %) samples were incubated at the photon flux density corresponding to that at the depth from which they were collected. The diurnal pattern of acetylene reduction was typical of that seen previously (Figs 1, 8). Furthermore, when represented as the amount of acetylene reduced at each depth integrated over the diurnal cycle, the specific activity of nitrogenase activity was again shown to decrease markedly with depth (Fig. 9). During this experiment, no significant difference was observed in temperature, ph or salinity in surface waters during the hours of illumination. However, the concentration of dissolved O reached a maximum in mid-afternoon, presumably as a result of O evolution by photosynthesis (Table 2a). The low concentration of dissolved O seen between 6: and : hours could, similarly, be a consequence of respiratory O consumption during the night. Above 12 m, temperature, ph, salinity and dissolved O did not vary much with depth, but between 12 and 1 m there was a marked decrease in temperature and a decrease in both ph and dissolved O (Table 2b). Intracellular glucan in populations from different depths In a mixed population of cyanobacteria sampled at the surface, the concentration of intracellular glucan, measured at 12: hours, was 38 mg mg Chla. This concentration steadily decreased in samples taken from increasing depth in the water column, falling to 33, 19, 12 and 11 mg mg Chla at 3, 6, 9 and 12 m, respectively.

9 RESEARCH N fixation by Baltic cyanobacteria C2H4 produced (µmol h 1 mg 1 Chla) 1 3 : 14: 18: 22: 2: 6: Local time (hours) : Depth (m) Fig. 8. Diurnal variation in the specific activity of nitrogenase in a natural population of cyanobacteria taken from various depths in the Middle Bank region of the Baltic Sea. Samples were collected at :, 14:, 18: and 22: hours on 12 August 199, and at 2:, 6: and : hours on 13 August 199 by means of a concentration, temperature and depth (CTD)-rosette sampler at the surface, and at depths of 3, 6, 9, 12 and 1 m. The contents (24 l) were filtered throught a -µm net. Samples were incubated for 2 h at the temperature and photon flux density prevailing at the surface. Mean surface photon flux densities during these incubations were 12243, , 9,,, 3193 and 2132 µmol m s, respectively. The amount of Chla present in each assay was.6.3,.8.3,.6.2,.7.3,.7.3,..2 and.6.3 µg, averaged for all depths at :, 14:, 18:, 22:, 2:, 6: and : hours, respectively, and.64.32,.6.29,.6.2,.7.27, and µg, averaged for all sample times at, 3, 6, 9, 12 and 1 m, respectively. Depth (m) 1 1 C 2 H 4 produced (mmol d 1 mg 1 Chla) Fig. 9. Total acetylene reduced by Baltic cyanobacteria at each depth integrated over a 24-h period August 199 (solid line), when samples were incubated at the photo flux density prevailing in surface waters, and 1 16 August 199 (dotted line), when samples were incubated at the light intensity prevailing at the depth from which they were sampled. DISCUSSION In blooms of Aphanizomenon and Nodularia spp. in the Baltic Sea, N fixation shows a characteristic diurnal pattern. Maximum rates of N fixation occur during the day, especially during the morning (Fig. 1), with minimum rates at about midnight. However, even during the period of darkness, nitrogenase activity could be detected in these cyanobacterial assemblages. There is no doubt that Baltic cyanobacteria are capable of fixing N for limited periods in the dark, and these organisms are probably responsible for most, if not all, of the nitrogenase activity that can be measured at night. However, in the absence of unequivocal evidence to the contrary, it remains possible that at least some of the residual nitrogenase activity seen in natural populations at night might be due to heterotrophic diazotrophs. The decline in nitrogenase activity that occurs during the afternoon could be related to fluctuations in the concentration of dissolved O, which reached a maximum mid-afternoon. However, as heterocysts do not produce O and their thick walls form a diffusion barrier against atmospheric O and N (Walsby, 198) this seems unlikely. In any case, inhibition of photosynthetic O evolution by DCMU did not prevent the afternoon decline in nitrogenase activity (Fig. 2a). Moreover, the relatively low concentration of dissolved O measured during the early morning (Table 2a) did not always correlate with high rates of N fixation (Fig. 1). Light might be much more significant than O in regulating nitrogenase activity in Baltic cyanobacteria. Nitrogenase

10 294 RESEARCH A. M. Evans et al. Table 2. Variations in temperature, salinity, dissolved O and ph (a) in surface waters at various times and (b) at : hours at various depths in the Baltic Sea on 1 August 199 Temperature (C) Salinity (PSU) O (ml l ) ph (a) Local time (hours) 6: : : : : (b) Depth (m) PSU, practical salinity units. activity is saturated at the relatively low photon flux density of 6 µmol m s, so the decreased rate of N fixation seen after noon could be a direct result of photoinhibition. For example, 11 August 199 the mean photon flux density between 12: and 18: hours was 933 µmol m s, sufficient to cause inhibition of N fixation (Fig. 7). High rates of nitrogenase activity during the morning, followed by a decrease in rates during the afternoon, have been observed on several occasions in both pelagic and benthic systems (Vanderhoef et al., 197; Carpenter et al., 1978). A midday drop is also often seen in photosynthesis, and this has been attributed to a decrease in quantum efficiency (Kromkamp et al., 199). The physiological basis for such a decrease in quantum efficiency is not yet known but, as a decrease in quantum efficiency is equivalent to a decrease in photon flux density, this phenomenon can either stimulate or inhibit N fixation depending upon whether the prevailing photon flux density is suboptimal or superoptimal for N fixation (Fig. 7). An alternative explanation for the decrease in photosynthesis seen around noon is that CO becomes depleted in the vicinity of the cyanobacterial cells after a period of active photosynthesis, an event that might be exacerbated by a local increase in ph which would further limit the availability of CO. Inhibition of photosynthetic CO fixation would adversely affect N fixation, mainly by restricting the import of carbon into heterocysts. The slightly lower rate of nitrogenase activity seen in the light in the presence of DCMU (Fig. 2a) might reflect a similar decrease in carbon supply to heterocysts. The typical diurnal pattern of N fixation was also seen when nitrogenase activity was assayed in the dark throughout a 24-h period. Samples collected during the day reduced acetylene in the dark at higher rates than were found in samples collected at night. This suggests that products of photosynthesis, accumulated during the light period, can support N fixation for a limited period of time. It is not clear, however, whether this is related to the heterotrophic generation of ATP, reducing power and acceptors for newly fixed N, or whether nitrogenase itself is specifically activated during the day. Illumination is required to obtain maximum rates of N fixation in both natural populations and laboratory cultures of Baltic cyanobacteria; nevertheless, it seems unlikely that oxygenic photosynthesis directly provides ATP and reductant for N fixation. The spatial separation between oxygenic photosynthesis (in vegetative cells) and N fixation (in heterocysts) would preclude such a relationship, and the limited effect of DCMU on nitrogenase activity is entirely consistent with this. DCMU probably acts by inhibiting photosynthesis in vegetative cells and eventually preventing them from exporting fixed carbon to the heterocysts. As long as carbon reserves are available in the vegetative cells or in the heterocysts themselves, inhibition of photosynthesis is unlikely to have a dramatic effect on N fixation. Heterocysts lack the DCMU-sensitive photosystem II, so cannot generate reductant using electrons derived from water. Light energy might, however, be involved in the provision of reductant for N fixation in these cells by driving a photosystem I-mediated electron flow from reduced pyridine nucleotides (generated from the oxidation of imported carbon compounds) to ferredoxin (Murai & Katoh, 197). In addition, heterocysts can use light energy to generate ATP by cyclic photophosphorylation, also catalysed by photosystem I. These processes would be insensitive to DCMU, but would

11 RESEARCH N fixation by Baltic cyanobacteria 29 be inhibited in the dark. This could explain the dramatic decline in nitrogenase activity that is seen when cultures are transferred to darkness. However, the fact that N fixation can be measured in the dark demonstrates that Aphanizomenon and Nodularia spp. can generate the necessary ATP and reductant entirely by light-independent metabolism. Acetylene was reduced during the dark period at significant rates, both in laboratory cultures and in natural populations of Aphanizomenon sp. and Nodularia spp. In the case of Aphanizomenon sp., nitrogenase activity during the dark period amounted to 33 and 31% of the maximum rate seen in the light in laboratory cultures and in colonies sorted from natural populations, respectively (Figs 3a, 4a). Individual colonies of Nodularia spp. sampled from a mixed population were able to reduce acetylene at night at a rate of 97% of the maximum rate seen during the day (Fig. 3b), while laboratory cultures of Nodularia SN34D reduced acetylene during the dark period at 68% of the rate seen in the light (Fig. 4b). The ability of Nodularia to sustain relatively higher rates of nitrogenase activity than Aphanizomenon during periods of darkness might reflect the fact that, in laboratory culture, Nodularia SN34D appeared to be more efficient than Aphanizomenon sp. at accumulating carbohydrate during the light phase (Fig. ). On both occasions when it was examined, N fixation per unit chlorophyll decreased with increased depth in the water column. This finding is in agreement with previous studies on the depth distribution of nitrogenase activity in the Baltic Sea (Rinne et al., 1978; Hu bel & Hu bel, 198) and with the theoretical model of Stal & Walsby (1998). However, this behaviour was observed not only when samples were incubated at the photon flux density prevailing at the sample depth, but also when samples were incubated under the conditions of illumination found at the surface. This implies that populations show at least some degree of adaptation to the depth at which they find themselves, although the nature of this adaptation is unclear. Superimposed upon any adaptive loss of nitrogenase activity, however, would be the influence of the environmental conditions themselves. The variation in the specific activity of nitrogenase with respect to sample depth could not be attributed to fluctuations in temperature (except below 12 m), salinity, dissolved O or ph (Table 2b). Furthermore, it did not correlate with any observed variation in the N: P ratio, which was 8.36 (.92 µm N:.11 µm P), 2.8 (.37 µm N:.13 µm P) and 4. (.4 µm N:.12 µm P) at, and 1 m, respectively. Low N:P ratios generally favour not only the presence of N -fixing cyanobacteria, but also high rates of N fixation (Rinne et al., 1978, 1981; Wallstro m, 1988). However, no increase in the specific activity of nitrogenase was observed in the population at m (Fig. 9). The higher N:P ratio at the surface was presumably due to decomposition of the bloom (Gabrielson & Hamel, 198). The most likely explanation for the observed decrease in N fixation with increasing depth is therefore the attenuation of surface illumination with depth. This decrease in photon flux density with depth could also explain the observed decrease in the concentration of endogenous storage carbohydrate (glucan). The glucan content of batch cultures of the heterocystous cyanobacterium Anabaena variabilis decreases markedly as the culture ages, and also once cultures are transferred to darkness (Ernst et al., 1984). The increased cell density of older cultures results in increased self-shading, thereby decreasing the ability of photosynthesis to generate glucan. When transferred to the dark, older cultures also exhibit both a decreased capacity for acetylene reduction and a lower ability to protect nitrogenase from inactivation by O, although both these effects can be prevented by addition of fructose. This suggests that A. variabilis can use either photosynthesis or an exogenous, metabolizable carbon source in order to sustain maximum rates of nitrogenase activity and efficient protection of the enzyme from inactivation by O. Nevertheless, in the absence of both light and an external source of carbon, stored glucan can support a lower rate of N fixation. However, when glucan reserves are low, the ability of Anabaena variabilis to fix N in the dark is compromised. The parallel decline in storage glucan and in the ability to fix N as Baltic cyanobacteria sink in the water column, and thereby receive less illumination, suggests that they might behave similarly to A. variabilis, although there is no evidence that they can metabolize any exogenous carbon source. Colonies at the surface of the bloom are believed to be older than those deeper in the water column (Reynolds & Walsby, 197). However, in Nodularia nitrogenase activity per heterocyst is greater in a newly developing population than in an older one (Hu bel & Hu bel, 198; Huber, 1986). On August 199, and also 1 16 August 199, the specific activity of nitrogenase in a mixed population of cyanobacterial cells at the surface was much higher than that in subsurface samples, even though the surface sample contained a large proportion of decaying material. Furthermore, samples taken from 3 and m did not possess the same capacity for N fixation, even when incubated under surface conditions. This phenomenon has also been described by Huber (1986) who showed that the nitrogenase activities of heterocysts of N. spumigena, sampled from the bottom and at mid-depth of a marine estuary, were improved when incubated at the surface but were never as great as the activity in heterocysts from samples actually taken from the surface. It was suggested that Nodularia spp. from

12 296 RESEARCH A. M. Evans et al. deeper waters might exhibit a lag in their ability to increase the rate of N fixation in response to changed conditions of illumination. Such adaptation of cyanobacteria to a particular depth implies that the water column in which they were suspended was not well mixed. The mechanism of adaptation is not known at present, but it might be significant that, in many cyanobacteria, the Fe protein of nitrogenase is found in two different forms, only one of which might be catalytically active (Reich & Bo ger, 1989; Du & Gallon, 1993; Bergman et al., 1997). In the marine cyanobacterium Trichodesmium, for example, a larger form of the Fe protein, presumed to be catalytically inactive, is converted to a smaller (assumed active) form within 3 h of illumination (Zehr et al., 1993; Chen et al., 1998). This behaviour might be important in explaining the diurnal fluctuations seen in nitrogenase activity in cyanobacteria from the Baltic Sea; in addition, conversion of nitrogenase to a less active form in deeper waters could also account for the lower rates of N fixation that were observed in samples from deeper water, even when incubated at the photon flux density prevailing at the surface. At present, however, it is not known whether two forms of the Fe protein occur in Aphanizomenon sp. and Nodularia SN34D. The work described here constitutes the first comparison of N fixation between natural populations of Baltic cyanobacteria and cultures grown under laboratory conditions. Despite the fact that both Aphanizomenon sp. and Nodularia spp. are heterocystous, colony-forming, diazotrophic cyanobacteria, they exhibit different patterns of N fixation during a diurnal cycle. This temporal variation in nitrogenase activity, along with the spatial variation observed in populations sampled from different depths, has clear implications for estimation of the total input of fixed N by cyanobacteria in the Baltic Sea. In formulating these estimates due account must be taken of the cyanobacterial species dominating at particular sampling sites. A mathematical simulation of N fixation in the Baltic Sea has already been published (Stal & Walsby, 1998), and is based on correlating the effect of illumination on nitrogenase activity with in situ measurements of the photon flux density at the surface, light attenuation with respect to depth, and cyanobacterial biomass. The data presented in this paper lend practical support to this model, and also point to ways in which it may be refined so that it becomes possible to calculate N input in the Baltic Sea in a reliable way. ACKNOWLEDGEMENTS We thank Drs Rolf Boje and Frank Jochem, Institut fu r Meereskunde an der Universita t Kiel, Kiel, Germany for organizing the cruises and for making available the nutrient and hydrography data. In addition, we thank the crew of R. V. Alkor for their assistance at sea. We also acknowledge the help of Professor Birgitta Bergman, Department of Botany, University of Stockholm, Sweden, who performed the immunogold labelling and allowed us to use her gas chromatograph during the 1996 cruise. We are grateful to Dr Kaarina Sivonen, Department of Applied Chemistry and Microbiology, Division of Microbiology, University of Helsinki, Finland and Dr Paul Hayes, School of Biological Sciences, University of Bristol, UK for the generous provision of laboratory cultures. Thanks are also due to ARISE, University of Amsterdam, The Netherlands, where part of this work was carried out. This study was funded by the European Commission Environment RTD Programme DG XII (Contract EV CT94 44) and forms part of the European Land Ocean Interaction Studies (ELOISE) network. This paper is publication 263 from the NIOO Centre for Estuarine and Coastal Ecology and has ELOISE No REFERENCES Bailey JL Miscellaneous analytical methods: estimation of protein, Folin Ciocalteau reagent. In: Bailey JL, ed. Techniques in protein chemistry. Amsterdam, The Netherlands: Elsevier, Bergman B, Gallon JR, Rai AN, Stal LJ N fixation by non-heterocystous cyanobacteria. FEMS Microbiology Reviews 19: Bottomley PJ, Stewart WDP ATP and nitrogenase activity in nitrogen fixing heterocystous blue-green algae. New Phytologist 12: Carpenter EJ, Capone DG Nitrogen fixation in Trichodesmium blooms. In: Carpenter EJ, Capone DG, Rueter JG, eds. Marine pelagic cyanobacteria: Trichodesmium and other diazotrophs. 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