Structure and development of a benthic marine microbial mat

Structure and development of a benthic marine microbial mat

FEMS Microbiology Ecology 31 (1985) 111-125 Published by Elsevier 111 FEC 0016 Structure and development of a benthic marine microbial mat (Microbi...

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FEMS Microbiology Ecology 31 (1985) 111-125 Published by Elsevier


FEC 0016

Structure and development of a benthic marine microbial mat (Microbial mat; cyanobacteria; purple sulfur bacteria; light; oxygen; sulfide; redox potential)

Lucas J. Stal, Hans van Gemerden * and Wolfgang E. Krumbein Geomicrobiology Division, University of Oldenburg~ Postfach 2503, D- 2900 Oldenbur& F.R.G., and * Department of MicrobioloD,, University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands Received 18 January 1985 Revision received and accepted 8 March 1985

1. SUMMARY Vertically stratified microbial communities of phototrophic bacteria in the upper intertidal zones of the North Sea island of Mellum were investigated. Growth and population dynamics of the cyanobacterial mat were followed over three successive years. It was concluded that the initial colonization of the sandy sediments was by the cyanobacterium Oscillatoria. In well-established mats, however, the dominant organism was Microcoleus chthonoplastes. The observed succession of cyanobacteria during mat development is correlated with nitrogen fixation. Nitrogen fixation is necessary in this low-nutrient environment to ensure colonization by mat-constructing cyanobacteria. Under certain conditions, a red layer of purple sulfur bacteria developed underneath the cyanobacterial mat in which Chromatium and Thiocapsa spp. dominated, but Thiopedia and Ectothiorhodospira spp. have alSO been observed. Measurements of light penetrating the cyanobacterial mat indicated that sufficient light is available for the photosynthetic growth of purple sulfur bacteria. Profiles of oxygen, sulfide and redox potential within the microbial mat were measured using microelectrodes. Maximum oxygen concentrations, measured at a depth of 0.7 mm, reached levels more than twice the normal air

saturation. Dissolved sulfide was not detected by the microelectrodes. Determination of acid-distilled sulfide, however, revealed appreciable amounts of bound sulfide in the mat. Redox profiles measured in the mat led to the conclusion that the upper 10 mm of the sedimentary sequence is in a relatively oxidized state. 2. INTRODUCTION Microbial mats are benthic populations of microorganisms, usually dominated by phototrophic bacteria [1-4]. In most cases, cyanobacteria are the mat-building organisms. Microbial mats are found in, e.g., marine intertidal fiats [2,4], hypersaline environments [5,6] and hot springs [7]. The cyanobacteria cover the sediment surface, and may form a rigid, leathery structure. Depending on the environmental conditions and species composition, these mats may gain a characteristic morphology [3]. Krumbein et al. [1] and Jorgensen et al. [3] described the mat types found in Solar Lake (Sinai). They recognized a 'shallow flat mat', a 'deep flat mat', a 'blister mat', and a 'gelatinous mat'. A classification of microbial mats on the basis of morphology was proposed by Golubic [8,9]. Gerdes et al. [10] described 10 salinity-related microbial mat zones of the Gavish Sabkha (Sinai).

0168-6496/85/$03.30 © 1985 Federation of European Microbiological Societies

112 Microbial mats may develop as vertically stratified communities of microorganisms. These laminated microbial ecosystems are the result of physico-chemical gradients within the mat, and of the physiology of the contributing microflora. These ecosystems are usually considered to be organo-sedimentary structures. They are produced by sediment binding, sediment trapping or precipitation, as a result of the metabolic activity and growth of microorganisms [11]. The lamination of the biologically active layer may be preserved in older, already degraded layers. Because of the similarities, modern microbial mats are considered to be analogous to Precambrian and younger stromatolites [11]. The development of mat systems mainly depends on: (1) grain size of the sediment; (2) light penetration; (3) capillary water rise; (4) sedimentation and erosion rates; and (5) grazing stress [12]. Orstedt [13] was the first to recognize the characteristic red microbial layer below the green cyanobacterial mat. This colourful banding has been named "Farbstreifen-Sandwatt' by other investigators [14,15]. The first comprehensive report on the North Sea microbial mats was by Hoffmann [16]. Laminated microbial mats develop where the Mellum sediments consist of fine, almost pure quartz sand [17]. Recently, we reported the importance of nitrogen fixation in the initial colonization of low-nutrient sediments [18]. The present contribution focuses on factors concerning mat development, species composition and physicochemical conditions.


3.1. Area of investigation The microbial mats investigated in this study are located on the island of Mellum, in the southern North Sea. The island is situated at latitude 59055 ' North, longitude 34044 ' East. Mellum is part of the chain of islands separating the Waddensea from the North Sea. The uninhabited island is very young, both geologically and historically. The transition from a sandbank, completely

covered by water at high tide, to an island, took place once more at the beginning of this century, and the process is still continuing [19]. Microbial mats are found in the upper intertidal zones of the island but are most pronounced on the western shore, where they form a strip below the vegetation border at 1.5-2 m above mean sea level.

3. 2. Sampling sites 3 Sampling sites were chosen, representing different states of mat development. The sampling sites were identical to those mentioned in [18]. Site 1 lies 1.60 m above mean sea level, far from the vegetation border, At this site, a very young community of cyanobacteria is developing. No noticeable iron sulfide precipitation occurs here, as judged by the absence of a black layer. Site 2 lies 1.70 m above mean sea level, and represents a well-developed microbial mat. At this site, the cyanobacterial layer is on average approx. 2 mm thick. Although sulfate reduction takes place, no purple layer of anoxygenic phototrophic bacteria is usually observed. Site 3 lies 1.58 m above mean sea level, near a tidal channel and close to the vegetation border. The cyanobacterial mat forms a very tough and leathery surface layer 0.5-1.0 mm thick. A distinct layer of sulfate-reducing bacteria is present (Laanbroek, personal communication), and a purple layer of anoxygenic phototrophic bacteria is also frequently observed at this site. 3.3. Sampling methods Sampling methods were essentially the same as those described in a previous paper [18]. The upper 1.3 mm of the mat was sampled with a corer (10 mm diameter). Vertical profiles of the mat were obtained by cross-sectioning a 7-mm-deep core (10 mm diameter) in 1-mm slices. Interstitial water was sampled by making small holes in the sediment and collecting seepage water. Sea water and interstitial water were filtered (Glass fiber 13400, Sartorius, F.R.G.) and stored at - 2 0 ° C immediately after sampling. 3.4. Light microscopy The relative frequencies of cyanobacteria in the mat were determined immediately after sampling by phase contrast microscopy. Individual morpho-

113 types were grouped roughly into the 3 abundancy classes 'dominant', 'common' and ' present'.

3.5. Classification of cyanobacteria As far as possible, we followed the provisional generic assignment of Rippka et al. [20]. Where possible and useful we used species names according to Geitler [21]. 3.6. Scanning electron microscopy Samples of the microbial mats from various sites on Mellum were studied by scanning electron microscopy (SEM). Small pieces of a cross-sectioned mat were fixed overnight in 4% glutaraldehyde solution in a phosphate buffer, pH 7.2. The samples were washed in phosphate buffer and double-distilled water. Water was removed in stages in an ethanol series from 10-95% followed by 4 passages through absolute ethanol. The sampies were than critical-point dried, gold-sputtered, and analyzed with a Cambridge Instruments S 180 SEM [22]. 3. 7 Pigment determination Chlorophyll a, pheophytin a, bacteriochlorophyll a and bacteriopheophytin a were determined by the method of Stal et al. [23]. Pigments were extracted with methanol and partitioned with n-hexane. The 2 phases were separated, and absorbance was read at 660 nm and 768 nm in the hexane phase. (Bacterio)chiorophyll was then q u a n t i t a t i v e l y c o n v e r t e d into (bacterio)pheophytin by acidifying the hexane phase with hydrochloric acid, and the absorbance readings were repeated. (Bacterio)chlorophyll and (bactedo)pheophytin were calculated according to Stal et al. [23]. In situ concentrations were calculated, taking into account that 90% of the chlorophyll a and pheophytin a enter the hexane phase, whereas 62% of the bacteriochlorophyll a and 69% of the bacteriopheophytin a appear in this phase. Finally, the amounts of chlorophyll a were corrected for absorption of bacteriochlorophyU a. No other bacteriochlorophylls were present, as was established by scanning the methanol phase [23]. 3.8. Light measurements Light penetrating the cyanobacterial mat was

measured using a Techtum Instruments Quanta spectrometer QSM-2500 (Sweden). A 1.5-mm-thick mat from site 2 was placed on the sensor. Reproducible illumination of the mat was guaranteed by the use of a slide projector.

3.9. Construction of microelectrodes Oxygen and sulfide microelectrodes were constructed as described by Revsbech et al. [24]. Oxygen was measured polarographically [25] using a custom-made nanoampere meter and a calomel reference electrode. A polarization voltage of 0.75 V was applied to the oxygen and reference electrodes. The electrode was calibrated over a range of oxygen concentrations from zero to air saturation. Actual oxygen concentrations were measured with an Orbisphere microprocessor oxygen indicator, model 2609 (Switzerland). Prior to each measurement, a 2-point calibration (zero and air saturation) was made. The sulfide electrode [26] was constructed by melting a 0.1-mm platinum wire in an AR-glass capillary. The tip of the electrode was plated with silver in a solution of 0.05 M AgNO 3 and 0.3 M KCN. The silver electrode tip was coated with silver sulfide by inserting the electrode in an 20% ammonium sulfide solution (Merck, F.R.G.) for 1 rain. Measurements were made with a custommade mV-meter, using a calomel electrode as reference. Calibration of the electrode was carried out in Na2S standard solutions, in a sulfide antioxidant buffer (Orion Research Inc., U.S.A.), composed of 40 g. 1"1 NaOH, 33.5 g. 1-1 Na2EDTA , and 17.5g. 1-1 Na-ascorbate. Reproducible readings were obtainable at concentrations of 10-5 M and higher. Redox microelectrodes were constructed by melting a 0.1-mm platinum wire in an AR-glass capillary [27]. Measurements were made with a custom-made mV-meter, using a calomel reference electrode. The redox readings were controlled by measurements in redox standard solution (Ingold, F.R.G.). pH Was measured using a commercial pH electrode, with a tip diameter of 3 mm (Ingold, F.R.G.). Microelectrode measurements were carried out in a 20 cm long, 7.5 cm wide sediment core, ,covered by 1 cm of sea water. The measurements

114 were started within 30 min of taking the core. Illumination was provided by a slide projector. Microelectrodes were inserted into the sediment using a micromanipulator.




3.10. Determination of sulfide and elemental sulfur Elemental sulfur was estimated by extracting the sediment with methanol, the methanol was partitioned with n-hexane and the absorbance was read photometrically at 260 nm [23,28]. Sulfide was determined colorimetrically after Pachmeyer [29]. Sediment samples were kept in small bottles, completely filled with 2% Zn-acetate. These samples were than acid-distilled prior to the determination of sulfide.




3.11. Determination of nitrite, nitrate, ammonia and salinity Sea water and pore water were filtered on a fiberglass filter (Sartorius 13400, F.R.G.) and analyzed for nitrate, nitrite, ammonia and salinity by the methods described [30].


4.1. Morphology and growth of the mats Sandy sediments of fine grain size in the upper littoral of Mellum are colonized by cyanobacteria. In the course of their development, due partly to the growth and metabolism of the cyanobacteria, rigid mats are formed. Oxygenic photosynthesis of the cyanobacteria provides the main input of organic material to the low-nutrient sandy sediments. This organic material allows the ecosystem to develop. A well-developed laminated microbial ecosystem shows stratified populations of microorganisms (Fig. 1). Cyanobacteria occur in the top layer - - the mat itself - - covering a layer of purple sulfur bacteria and a layer of sulfate-reducing bacteria. The cyanobacterial mat was investigated by scanning electron microscopy (SEM). Fig. 2 shows a freshly colonized sediment, illustrating the uniform grain size. Mainly filamentous cyanobacteria are growing in and on the sediment. Sand grains are entangled in the filaments, an imPortant"process for the development of a siliciclastic mat. Oscillatoria sp. is an important organism

Fig. 1. Model of a laminated microbialsediment ecosystem. in initial sediment colonization [18]. In young mats, Oscillatoria sp. is the principal organism found, although Spirulina sp. and coccoid cyanobacteria can also be found (Fig. 3). After the initial colonization of the sediment by Oscillatoria sp., Microcoleus chthonoplastes, according to Rippka et al, [20] a member of the LPP-B group (Lygnbya, Plectonema, Phormidium), becomes the dominant

Fig. 2. An SEM-photograph of freshly colonized sediment. Filamentous cyanobacteriaare attached to the sand. The uniform grain size of the sediment can be seen.


Fig. 3. An SEM photograph of a typical example of a young mat system. Oscillatoriaslap. are the dominant organisms,but other cyanobacteria, e.g., Spirulina sp., are also present.

organism (Fig. 4a and b). This organism is responsible for the final structure, and forms a tough coherent mat. The occurrence of m a n y trichomes within a c o m m o n sheath is characteristic for this organism. In the Mellum mats, we observed 20-100 trichomes in one sheath. A thick bundle, as shown in Fig. 4b, is subdivided into several compartments. At present, the functional explanation of bundle formation and compartmentation is unknown. This mechanism conceivably protects the organism against dessication during dry periods, and allows the free movement of individual trichomes whilst maintaining a firm anchorage of the bundle. The formation of a common sheath has never been observed in culture. The population dynamics of the cyanobacteria at the 3 sites were followed by observation at regular intervals over the three-year period (Fig. 5). The mats of site 1 were completely destroyed during the winter of 1980/1981. This enabled us to study the recolonization of the sediment. In the early stages, coccoid cyanobacteria such as Gloeocapsa sp., Synechocystis sp. and Synechococcus sp. were observed frequently. The filamentous Spirulina sp. was also present most of the time. Oscillatoria sp., however, was always present, and in almost all cases this organism was dominant. The filamentous LPP-forms were never present

Fig. 4. (a) An SEM photograph of a M. chthonoplastes mat.

This organism forms trichome bundles with a common sheath. (b) Detail of a trichome bundle of M. chthonoplastes.

during the initial colonization. This was in contrast to the older mats at sites 2 and 3. Here, the LPP-B form M. chthonoplastes (LPP-B 1) was always dominant. On m a n y occasions, Oscillatoria sp. was also observed. Other LPP-B forms (Phormidium, LPP-B 2) were observed only rarely, as were Spirulina sp. and Gioeocapsa ~p. It is concluded that Oscillatoria sp. plays an "Lmportant role in initial sediment colonization. The ability of Oscillatoria sp. to fix nitrogen gives this organism its ecological advantage in a low-nutrient environment [31]. Its occurrence in older mats is also






I HI1,1

i:kl rollLlkl r





Site 1



Aug 1980

' Oct





Jun 1981









Aug 1982







Fig. 5. The abundance of cyanobacteria in the mat was roughly classified as either 'dominant' (3), 'common' (2) or 'present' (1). At regular intervals during 3 years, abundancies were estimated by light microscopy of samples from the sites 1, 2 and 3. I LPP-B (M. chthonoplastes); LPP-B 2 (Phormidium sp.); Oscillatoria sp. ( Lyngbya aestuarii ); Spirulina sp. (Spirulina subsalsa ); Gloeocapsa sp. ( Chroococcus turgidus ); Synechocystis sp. ( Merismopedia glauca ); Synechococcus sp. ( Synechococcus elongatus ).

positively correlated with nitrogen-fixing activity [181. The growth of a mat usually starts in May or June and stops in October or November (Fig. 6). The actual starting point and end-point of the growth period depends strongly on the prevailing weather. The initiation of mat growth in spring also depends on the conditions during the preceding winter. Erosion by ice floes or driftwood, or high sedimentation rates, can eventually destroy a mat. An example for this is given in Fig. 6. The severe winter of 1980/1981 completely destroyed the young mat at site 1. The older mats from sites 2 and 3 were also greatly reduced, and only in 1982 did chlorophyll a content approximately regain its 1980 value. The chlorophyll a contents of sites 2 and 3 were about 3 times as high as that of site 1. More than

450 mg chlorophyll a - m -2 was found at sites 2 and 3. This amount is comparable with the standing crop which Bauld [4] measured in the mats of Shark Bay, Australia. Although the mats at sites 2 and 3 had approximately the same chlorophyll a content, their morphologies were different. The mat at site 2 was approx. 1.5-2.0 mm thick, while the mat at site 3 was only 0.5-1.0 mm. This mat, however, was very tough and leathery, and contained little sediment compared to the mat at site 2. In addition, purple sulfur bacteria were often found under the site 3 mat. At site 2, no distinct red layer of purple sulfur bacteria was ever observed.

4.2. Salinity and nitrogen compounds Salinity and the concentrations of nitrate, nitrite and ammonia were determined in pore water from

117 SITE I




SITE 2 ~" t.5C




30C Is(

o e,-




300 150


~98G. . . . . . . . .19.81. . . . .o. .~. . . . . .198"2 .

Fig. 6. Chlorophyll a concentrations at sites 1, 2 and 3. The upper 1.3 m m of the mat was sampled.

the 3 sites, and compared with sea water at low tide. Our measurements revealed pore water salinities which were little higher than that of sea water (Table 1). The sea water salinity showed a slight increase from 2.3% in May to 2.8% in October. Pore water from all stations was relatively constant in salinity, varying from 2.9-3.1%. No data on short-term fluctuations were collected. Theoretically, the salinity of the pore water should show relatively large changes. Rain will dilute pore water, resulting in a low salinity up to almost fresh water conditions [16], whereas extended evaporation periods will raise the salinity. Nitrate was measured in the same water samples used for salinity measurements. Nitrate concentration in sea water was low (2.9-5.6 #g NO3-N • 1-1), and there was a slight increase between May and October (Table 2). Pore water, however, had a nitrate concentration 3 times as high as seawater, probably due to microbial activity. These concentrations are still comparatively low (up to 15.7 #g NO3-N. 1-1). Nitrite concentrations were also very low. In contrast to nitrate, pore water did not contain more nitrite than sea water (Table 3). Ammonium concentrations were much higher than those of nitrate or nitrite (Table 4). Sea water contained 115-244 #g NH3-N.1-1. Pore water concentrations were much higher, particularly at sites 2 and 3, where concentrations of 430-666 #g NH3-N • 1-1 were found. The high concentrations at sites 2 and 3 correlated with the high biomass at these sites. Moreover, the low concentrations at site 1 (127-358 #g N H 3 - N . 1 - 1 ) were also in agreement with the high specific nitrogen-fixing

Table 1

Table 2

Salinity measurements of sea water and interstitial water from the 3 stations.

Nitrate concentrations in sea water or interstitial water from the 3 stations.

Salinity is expressed as promille. (-), No interstitial water present. Measurements were taken from May to October 1982.

Values are in # g NO3-N. 1-1. (_), N o interstitial water present. Measurements were taken from M a y to October 1982.


May July August September October

Sea water

23 26 25 27 28


Interstitial water Station I

Station II

Station III

34 31 31 -

29 30 30 -

25 30 29 -

May July August September October

Sea water

3.7 2.9 4.3 5.3 5.6

Interstitial water Station I

Station II

Station III

8.0 6.8 15.7 -

4.1 12.9 10.0 -

4.0 10.1 6.8 -

118 Table 3

Table 5

Nitrite concentrations in sea water or interstitial water from the 3 stations.

Light penetration through sand.

Values are in # g NO2-N. 1-1. (_), No interstitial water present. Measurements were taken from May to October 1982. M onth

Sea water

M ay July August September October

2.3 2.1 1.7 2.7 2.4

Interstitial water Station I

Station II

Station III

2.0 3.2 0.3 -

3.9 2.1 1.0 -

2.0 2.4 2.2 -

Light was measured as the integration of quanta, m - 2 - s - 1 from 400-750 nm. % Light that penetrates pure sand is shown. % Light that penetrates a cyanobacterial mat of 1.5 mm thickness, was 0.45%. Daylight in October on a cloudy day at 1:00 p.m. had 1320×1017 Q.m-2-s -1. Reproducible light penetration measurements were obtained using a slide projector as light source. D e pt h (mm)

Dry Sand (%)

Wet Sand (%)

1 2

2.55 0.55

10.64 3.36

3 4 5

0.25 0.20 0.15

0.32 0.20 0.11

Table 4 A m m o n i u m concentrations in sea water or interstitial water from the 3 stations. Numbers are in ~g N H a - N . 1 - 1 . (-), No interstitial water present. Measurements were taken from May to October 1982. Month

Sea water

Interstitial water Station I

Station II

Station III

M ay July August September October

244 115 144 185 186

358 232 127 -

430 464 579 -

488 666 579 -

activity at this site [18]. The ammonium concentrations at sites 2 and 3 are presumably not high enough to inhibit nitrogen fixation.

4. 3. Light penetration Cyanobacteria do not usually grow at high light intensities. The optimal light intensity for the growth of cyanobacteria varies from 15-150/xEins t e i n . m - 2 . s -l, approx. 1-10% of daylight illumination. Thus, cyanobacterial populations do not usually occur in direct sunlight. Sometimes, the cyanobacteria can be found under a thin layer of sand. If this is not the case, the uppermost cyanobacteria may be inactive and function as a light-filter. The majority of the cyanobacteria show gliding motility, so that when sedimentation occurs the cyanobacteria move phototactically in the direction of optimal light conditions. This is only possible if the sedimentation rate is not exceedingly high. Light penetration through fine sand is

good, albeit limited. Table 5 shows the results of measurements of light penetration through sand from the intertidal flat which was not colonized by microorganisms. Light penetrated wet sand much better than dry sand, as also observed by Hoffmann [32]. We measured light penetration to a depth of 5 mm of uncolonized sand, about the same depth as reported by Hoffmann [32]. Indeed, when artificial mats of M. chthonoplastes cultures were buried under approx. 3 mm of sand in the laboratory, the cyanobacteria were found at the surface within 2 or 3 h. Similar observations were reported recently by Pentecost [33] who observed gliding rates of 3-7 mm- h -1. Under well-developed cyahobacterial mats, a layer of purple sulfur bacteria often occurs. Fig. 7 shows the distribution of chlorophyll a and bacteriochlorophyll a in a depth profile. It is evident that the maximum of bacteriochlorophyll a lies immediately below the chlorophyll a maximum. Direct photoacoustic spectroscopy of mat samples from the same environment have also shown distinct layers of chlorophyll a and bacteriochlorophyll a [34]. The total light (400-750 nm) penetrating a mat 1.5 mm thick, was estimated as 0.45% of the incident light. However, the spectral composition of the penetrating light is of particular relevance. It was found that light of long wavelength penetrates the mat much better than light of shorter wavelengths (Fig. 8). Fig. 9 compares the spectrum of a methanol/hexane ex-


~2 E

-r-3 I---

t't ,,,t~ r-~

,<2 cu L

Z C)

Z O I--et CIC C~ (d3 e~

t/I Z

5 10 PIGMENT (jJgIsampte)

rr" I--


Fig. 7. Gradients of chlorophyll a (open bars) and bacteriochlorophyll a (solid bars) in a 7-ram-long sediment core of the microbial mat. The core was cross-sectioned into 1' mm slices.

tract of a mat containing cyanobacteria and purple sulfur bacteria and the proportional light penetration through a cyanobacterial mat without purple 400


i/ ¢,w




/ I




Fig. 9. Spectrum of an extract of a mat sample (solid line), containing chlorophyll a and bacteriochlorophy]l a. The proportional light penetration through the cyanobacterial mat is shown by the closed symbols.


I / I


600 WAVELENfiTH (nm)

/ I


sulfur bacteria, at various wavelengths. At the wavelength of maximal absorption of bacteriochlorophyll a, the light penetration through the cyanobacterial mat is relatively high. Further, the in vivo absorption of bacteriochiorophyll a occurs at even longer wavelengths than in the methanol/ hexane extract of Fig. 9. It can therefore be expected that the purple sulfur bacteria obtain considerably more light than the 2.5% of the incident light estimated at 750 nm (Fig. 9). Moreover, purple sulfur bacteria are generally able to grow at light intensities as low as 5-10 /~Einstein. m -2. S - 1 [35].

500 600 700 WAVELENETH (nm) Fig. 8 Spectrum of light penetrating a 1.5-ram-thick cyanobacterial mat (solid line). The mat was illuminated with a slide projector. For comparison, the spectrum of the light source is also shown (dashed line). Curves represent different scales.

4.4. Gradients of oxygen, redox potential, sulfide and sulfur Far red light ( > 700 nm) which penetrates the mat very well (Fig. 8), does not permit oxygenic photosynthesis [36]. This is also evident from the


~2 E D

-7- 3 b---




1_ L




25 50 SULFIDE (~g/samp[e) Fig. 11. Acid-volatile sulfide gradient. The sulfide was measured in 1-mm slices of a 8-mm-long cross-sectioned sediment core. The samples were acid-distilledprior to sulfide determination.




Ct_ i,i

o 5 6

7 m


2 -



I t~


I 8


I 12


I 16




OXYGEN (mg / I ) Fig. 10. Oxygen gradient in the sediment. This gradient was obtained using microelectrodes. The cyanobacterial layer was 1.5 mm.



_1 E E

--2 -1-

o x y g e n g r a d i e n t in Fig. 10. O x y g e n c o n c e n t r a t i o n s d r o p p e d to zero in the lowest region of the mat. M a x i m u m o x y g e n c o n c e n t r a t i o n s in the ill u m i n a t e d m a t were o b s e r v e d at a d e p t h of a p prox. 0.7 m m , p r o b a b l y d u e to o p t i m a l light intensity in this p a r t o f the mat. T h e small decrease in o x y g e n c o n c e n t r a t i o n at the m a t surface is p r o b a b l y a result of p h o t o o x i d a t i o n caused b y high i n c i d e n t light intensity. D u r i n g the night, o x y g e n c o n c e n t r a t i o n s d e c r e a s e d r a p i d l y to zero, a n d the m a t surface b e c a m e a n o x i c ( d a t a n o t shown). A l t h o u g h the s e d i m e n t is always a n o x i c at d e p t h s greater t h a n 1.5 m m , and, d u r i n g the night, even at the m a t surface, we were u n a b l e to d e t e c t sulfide with a sulfide microelectrode. Even after p r o l o n g e d d a r k i n c u b a t i o n , the c o n c e n t r a t i o n of sulfide was b e l o w the limit of detection. T h e sulfide e l e c t r o d e o n l y senses free S 2 - ions, with a limit of

r~ t.iJ


+ 350

* t~00

+ t~S0

REDOX POTENTIAL (mY) Fig. 12. Redox gradient in the sediment measured using microelectrodes. The readings were corrected for the difference in voltage of the standard hydrogen electrode. The open symbols refer to the day situation, the black symbols refer to the night situation.

121 detection of 10-5 M. The concentration of S 2 - is dependent on the total free sulfide concentration ( H 2 S + H S - + S2-), and on the prevailing pH. The p H of the sea water was 8.3. The upper 3 mm of the sediment had a p H of 7.9 in the light and 7.3 after dark incubation. Under these conditions, only small fractions of the total sulfide will exist in the S 2- form. Therefore, no conclusion can be drawn at present on the total free sulfide concentrations in the sediment. The black colour of the sediment indicates that appreciable amounts of sulfide must be present in a bound form (e.g., FeS) (Fig. 11). In addition to oxygen and sulfide, the redox potential in the sediment was determined using microelectrodes (Fig. 12). It is striking that the redox potential of deeper sediment layers is still very high. After dark incubation, the redox potential decreased only slightly in the upper 3 mm, and showed the same pattern as the redox gradient in the illuminated core below this depth. We conclude that although oxygen is absent, the system remains in a relatively oxidized state. The pattern of elemental sulfur in a cross-sec-

"g2 E

-,-3 ,,,4. r--1 5

20 4.0 60 80 SULFUR 0Jg/sampte) Fig. 13. Gradient of elemental sulfur in a 7-ram-longcross-sectioned core. l-ram Slices were extracted with methanol and partitioned with n-hexane. Absorption of sulfur was read at 260 nm.

tioned core is shown in Fig. 13. The concentration of elemental sulfur was highest in the upper 1 ram. This is conceivably the result of the abiotic oxidation of sulfide by oxygen, or the activity of colorless sulfur bacteria (e.g., Beggiatoa spp.), since very few purple sulfur bacteria are seen at these depths.


5.1. Mat development The initial colonization of the Menum intertidal sediments was performed by Oscillatoria sp. It has been shown that this Oscillatoria sp. has the capability of nitrogen fixation, even under complete aerobic conditions [31]. Other reports refer to this Oscillatoria sp. as Lyngbya (Stam, personal commtmication). Hoffmann [16] never observed Oscillatoria sp. as a dominant organism, but frequently found Lyngbya in Microcoleus mats, sometimes even as the dominant species. Because of our results, we suppose that Hoffmann's Lyngbya is the same organism as our Oscillatoria. Bauld [4], and Javor and Castenholz [2,37] reported mats with mixed populations of Lyngbya aestuarii and M. chthonoplastes, and even almost monospecific mats of L. aestuarii. Earlier, Van Baalen [38] reported the growth of L aestuarii in a medium free of combined nitrogen. This pointed to a possible capability to fix nitrogen. The data on concentrations of nitrogen compounds confirmed the importance and necessity of nitrogen fixation in the Mellum mats [18]. The tidal flats of Mellum Island never showed heterocystous cyanobacteria upon microscopic examination. Nevertheless, we were able to isolate heterocystous cyanobacteria (e.g., Calothrix scopulorum) from the Mellum mats by using media free of combined nitrogen (unpublished results). Conceivably, heterocystous cyanobacteria occur in very low frequencies, and are not adapted to the prevailing environmental conditions in the mat, which would allow them to compete with other cyanobacteria. However, other reports have shown that heterocystous cyanobacteria are not always excluded from marine mat systems [2,16,37]. M. chthonoplastes is the most important mat-

122 builder in the microbial mats of Mellum. This organism is a cosmopolite and has been reported as the main component of mat systems of many other sites around the world [2-4,16,39,40]. A well-established microbial mat in the intertidal zone of Mellum Island is dominated by M. chthonoplastes, mostly mixed with Oscillatoria sp. The presence of Oscillatoria sp. in such mats is correlated with nitrogenase activity [18]. This indicates that nitrogen fixation remains an important ecological factor even after the mat system is established. Other species, e.g. Spirulina sp., Phormidium sp., Gloeocapsa sp., Synechocystis sp. and Synechococcus sp., do occur, sometimes even at quite high frequencies, but predominantly in younger microbial mat systems. Their ecological position in the mat is not yet understood. Salinity changes may play an important role, as some of the isolated strains appear to be typical fresh-water organisms. Our results showed fairly constant moderate salinities in the interstitial water. Most of the isolated strains grow excellently at these salinities. It cannot, however, be excluded that very low salinities occur during and after heavy rainfall [16]. This may result in a very fast succession of the typical freshwater species.

5.2. Light penetration A red layer of purple sulfur bacteria sometimes develops under the cyanobacterial mat. Orstedt [13] mentioned Thiopedia rosea ( Erythroconys littoralis) as the origin of the red color. In the Mellum mats, we found Thiocapsa sp., Chromatium sp., Thiopedia sp., and Ectothiorhodospira sp. It is obvious from the presence of sulfur globules that these organisms grow (in part) at the expense of sulfide oxidation. Our results show very dearly that light is available for the photosynthetic growth of purple sulfur bacteria. These results are in agreement with the data of Javor and Castenholz [37]. No specific pigments of green sulfur bacteria were detected in extracts of samples from the microbial mats on Mellum, and attempts to isolate green sulfur bacteria were invariably unsuccessful. In meromictic lakes, Chlorobiaceae are usually dominant under layers of Chromatiaceae [41]. Presumably the high redox potential and the possible oxygenation of deeper layers of the sedi-

ment (e.g., by upwelling of aerobic seawater), exclude the growth of Chlorobiaceae, which are strict anaerobes. Chromatiaceae, on the other hand, are able to withstand oxygen, and may even grow at low oxygen concentrations [42]. Microbial mats may be covered by fine sand. Therefore, the cyanobacteria need to glide in the direction of the light to find optimal growth conditions. Phototactic movement depends on noticeable light penetrating through the sand above. Light penetrated through up to 5 mm of sand. Wet sand allowed better light penetration than dry sand (Table 5). These results agree with the measurements of Hoffmann [32], who also found better penetration of light of long wavelength. This red light is very efficient in stimulating positive phototaxis in cyanobacteria [43].

5. 3. Oxygen and sulfide gradients The oxygen gradient measured in the Mellum mats showed the same pattern as that measured by Revsbech et al. [24] in mats at Solar Lake (Sinai). In contrast to the work of Revsbech et al. [24], we were unable to measure any S 2- in the Mellum mats. At the pH prevailing in the microbial mats of Mellum, virtually all free sulfide will exist in the form of HS-. Although we cannot exclude the presence of considerable amounts of free sulfide, we suppose that most of the sulfide produced by the sulfate-reducing community occurs as bound sulfide (FeS or FeS2). The presence of high concentrations of sulfide becomes evident from the measurements of acid-distilled samples. Sulfide is a toxic agent for most organisms. Cyanobacteria and purple sulfur bacteria show pH-dependent sulfide toxicity, H2S being the more toxic compound [44,45]. For cyanobacteria, Howsley and Pearson [44] suggested that sensitivity to sulfide might exclude heterocystous cyanobacteria from these benthic environments. However, because of the apparent low concentrations of free sulfide, and because virtually no H2S will be present at the prevailing pH, we suppose that other mechanisms exclude heterocystous cyanobacteria from these environments. One possibility is that heterocystous cyanobacteria cannot cope with anaerobic conditions in the dark [46,47]. Recently, Howarth and Merkel [48] and How-

123 arth amd Marino [49] reported the formation of pyrite and elemental sulfur as the major products of sulfate reduction in salt marsh sediments. Pyrite is not included in the measurements of acid-distilled sulfide. The measurement of pyrite in the microbial mats needs to be done and deserves close attention. Elemental/sulfur, on the other hand, may be formed not only during sulfate reduction but also by purple sulfur bacteria during photosynthesis [28], cyanobacteria in the presence of sulfide [36,50], or colourless sulfur bacteria e.g., Beggiatoa spp. [51], or by a chemical oxidation of sulfide with oxygen. The redox potential in the sediment was relatively high. This agrees very well with the measurements of Fenchel [27] in comparable sediments. The redox gradients measured by Jorgensen et all [52] in Solar Lake microbial mats were totally different. Here, a decrease in redox potential of > 400 mV was encountered at a depth between 3-5 mm, while we found only a decrease of 100 mV between 2-5 mm depth. At the mat surface, the difference in redox potential between day and night was about 300 mV in Solar Lake, but only 50 mV in the Meilum mats. In view of the high redox potential, it can be concluded that the concentration of free sulfide must be low. However, free sulfide will be released from the bound form due to the actively growing population of purple sulfur bacteria. The presumably low concentration of free sulfide and the high redox potential in the North Sea microbial mats are highly significant, and unique compared with hypersaline laminated microbial ecosystems with high sulfate reduction rates, or with thermal microbial mats with a high natural supply of hydrogen sulfide.

ACKNOWLEDGEMENTS We sincerely thank the Mellumrat for allowing us to carry out fieldwork on the nature reserve MeUum. We are grateful to Dr. F. Wunderlich, and to Captain K. Kummer and the crew of the MS 'Senckenberg' for the numerous crossings to Mellum. W. Tecklenburg and T. Prause surveyed the position and heights of our measuring stations. We also thank Mr. D. Beyer and Mrs. S. Gross-

berger for skilled technical assistance. C. Giele is acknowledged for her skilled operation of the SEM. This work was supported in part by grant Kr 333/16-2 of the Deutsche Forschungsgemeinschaft.

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