The cyanobacterial community of the Zerka Ma’in hot springs, Jordan: morphological and molecular diversity and nitrogen fixation

Danny Ionescu1,2, aharon oren1,*, orly levItan3, Muna hInDIyeh4, hanan MalkawI5 and Ilana BerMan-Frank3

1 The Institute of Life Sciences, The Hebrew University of Jerusalem, Jerusalem, Israel2 The School for Marine Sciences, The Ruppin Academic Center, Emek-Hefer, Israel3 Mina and Everard Goodman Faculty of Life Sciences, Bar-Ilan University, Ramat Gan, Israel4 Department of Water and Environmental Engineering, German Jordanian University, Amman, Jordan5 Yarmouk University, Irbid, Jordan

With 5 figures and 2 tables

Abstract: The area of the Dead Sea, bordering Jordan and Israel, contains many springs with different physical and chemical properties. Hot springs are found on the eastern shore of the lake: the springs of Zara (the ancient Callirrhoe) with tem-peratures up to 59 °C and those of Zerka Ma’in, 5 km inland. The latter springs, mentioned by Josephus (1st century C.E.), yield fresh water with a low sulfide content, near-neutral pH and temperatures up to 63 °C. Although the growth of green microorganisms was already noted in 1807 by the German explorer ulrich Jasper seetzen, the microflora of the springs has remained unexplored. In 2005 we began a series of surveys characterizing the microbial diversity of the springs. The sources of the Zerka Ma’in springs and their outflow channels are covered by green to orange mats, containing a diverse community of unicellular and filamen-tous cyanobacteria. Conspicuous types include Thermosynechococcus, unicellular Gloeocapsa types, and Spirulina-like filaments. Large colonies of Scytonema were also found. Several representative types of Thermosynechococcus, Gloeocapsa, and Mastigocladus/Fischerella were isolated and cultured. Amplification of cyano-bacterial 16S rRNA genes from DNA extracted from the mats showed a high di-versity of Thermosynechococcus in the springs. Low concentrations of ammonium and nitrate in the spring water and the presence of heterocystous cyanobacteria indicated that biological nitrogen fixation may be important in the spring ecosys-

* corresponding author

Algological Studies 130 109–124 Stuttgart, October 2009

DOI: 10.1127/1864-1318/2009/0130-0109 1864-1318/09/0130-109 $ 4.00

© 2009 E. Schweizerbart’sche Verlagsbuchhandlung, D-70176 Stuttgart

110 D. Ionescu et al.

tem. nifH genes related to those of Fischerella, Phormidium sp., and Lyngbya sp. were amplified from the community DNA. Laboratory experiments with hetero-cystous isolates showed the occurrence of nitrogen fixation (acetylene reduction) at 52 °C but not at 63 °C. However, the possibility that nitrogen is fixed in situ at 63 °C cannot be excluded as reverse transcription PCR with mRNA isolated from the site yielded an amplified gene with 100 % homology to the nifH gene found in a filamentous cyanobacterial culture obtained from the site, and low but significant rates of acetylene reduction were found during in situ incubations.

Key words: cyanobacteria, 16S rRNA gene sequences, hot springs, nitrogenase, thermophilic

Introduction

The Dead Sea Rift, within the Syrian-African Rift Valley, is rich in thermal springs. Some are freshwater springs, others are saline or hypersaline. Ex-amples are Hamat Gader (up to 52 °C) and the hot springs of Tiberias (up to 60 °C). The area of the Dead Sea, on the border between Jordan and Israel, contains many springs which differ in their physical and chemical properties. The eastern bank of the Dead Sea (Jordan) is especially rich in thermal springs. These include the springs of Zara on the Dead Sea shore (the ancient Callirrhoe; Donner 1963) with temperatures up to 59 °C, and the hot springs of Zerka Ma’in, 5 km inland, up to 63 °C (abu aJamieh

Fig. 1. Map showing the location of the Zerka Ma’in and the Zara hot springs east of the Dead Sea and sampling sites A1, A2, B1, and B2.

The cyanobacterial community of hot springs 111

1980, 1989, swarieh 2000) (Fig. 1, Table 1).Very little is known about the phototrophic communities inhabiting

the hot springs of the Dead Sea Rift. A morphological description of the cyanobacteria of the Tiberias hot springs has been given (Dor 1967), and taxonomic identifications of the species present in the Zara springs (limited to areas in the temperature range 35–40 °C) were published long ago (Frémy & rayss 1938). The German explorer ulrich Jasper seetzen (1767–1811) wrote after his visit to the hot springs of Zerka Ma’in in 1807: “In dem Wasser wuchs eine grüne schleimige Conferve” [In the water grew a green slimy microscopic alga] (seetzen 1854). blanckenhorn (1912) provided a somewhat more detailed macroscopic description of the green material in the springs, yet no further studies are available on their biota.

We have recently begun a microbiological survey of the Jordanian hot springs (ionescu et al. 2007). Here we present information on the morphological diversity of the cyanobacteria inhabiting the Zerka Ma’in springs and 16S rRNA gene sequence-based diversity, as found both in environmental samples and in cyanobacteria cultured from these samples. In addition, we evaluated the potential for nitrogen fixation by the cyanobacterial community, by examining the presence of nifH genes, their expression, and nitrogen fixation activity.

Table 1. Chemical and physical characteristics of water samples collected at the dif-ferent sampling sites of the Zerka Ma’in springs, as analysed at the Department of Earth and Environmental Sciences, Yarmouk University.

Sampling site

A1 A2 B1 B2

TemperaturepHH2SConductivityTotal dissolved saltsAlkalinityHardness[Cl–][NO3–][SO42–][HCO3–][Mg2+][Ca2+][K+][Na+][Sr2+]

(°C)

(mM)(µS cm–1)

(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)(ppm)

636.690.3

21401306120560

860.10

1972391

9018533

86.33.31

626.620.3

41801267110520

810.41.02196ND9118635

85.6ND

586.380.0324601428130540

900.90.72210ND9319337

78.5ND

586.81NA24701445130580

790.60

211ND9219538

78.2ND

1As determined by I. Gavrieli, The Geological Survey of Israel. ND = not determined.

112 D. Ionescu et al.

Materials and methods

Samples were collected from the sampling sites indicated in Fig. 1 between December 2005 and June 2007 into sterile plastic 50 mL tubes and stored in a refrigerated coolbox until processed within 40 h of sampling. Photomicro-graphs were made in a Zeiss Axiovert model 135 TV microscope equipped with phase-contrast optics. Cultures were prepared in 250 mL Erlenmeyer flasks containing 100 mL BG-11 growth medium (stanier et al. 1971), and incubated in a New Brunswick Innova 44 illuminated incubator at 45 °C and an incident light intensity of 175 µmol quanta m–2 s–1. For DNA ex-traction, cultures and environmental samples were incubated for 30 min at 100 ºC in lysis buffer (100 mM Tris-HCl, 50 mM EDTA, 10 mM NaCl, 1 % SDS, pH 8), followed by extraction with phenol-chloroform-isoamyl alco-hol (25:24:1). The extracts were washed with chloroform-isoamyl alcohol (24:1) until no interphase could be observed. DNA was then precipitated with cold ethanol, washed with ice-cold 70 % ethanol, and resuspended in water. Fragments of the 16S rRNA gene were then amplified by PCR, using cyanobacteria-specific primers as specified in Table 2. The primer sets 29F – 809R and 740F – 1494R were used for cultures, while 106f and 781R were used for environmental samples. Amplicons originating from environmen-tal sequences were cloned using the InsTAclone kit (K1214, Fermentas, Lithuania) and verified using colony PCR before being sequenced.

Samples for RNA analysis were collected into 1 mL of TriReagent (TR 118, Molecular Research Center Inc., OH, USA) and immediately placed in liquid nitrogen to be later processed according to the manufacturer’s in-structions. To evaluate the existence of the nifH mRNA in total RNA ex-tracts, cDNA was produced using a first strand cDNA synthesis kit (K1612, Fermentas, Lithuania) together with the nifHR primer (Table 2). The

Table 2. PCR primers used in this study.

Primer Primer target Sequence

27F General 16S rRNA1 AGA GTT TGA TTT ACG CGA CA

809R Cyanobacterial 16S rRNA1 GCT TCG GCA CGG CTC GGG TCG ATA

740F Cyanobacterial 16S rRNA2 GGC YRW AWC TGA CAC TSA GGG A

1494 General 16S rRNA GGY TAC CTT GTT ACG ACT

106F Cyanobacterial 16S rRNA3 CGG ACG GGT GAG TAA CGC GTG A

781R Cyanobacterial 16S rRNA3 GAC TAC WGG GGT ATC TAA TCC CWT T

nifHF Cyanobacterial nifH4 CGT AGG TTG CGA CCC TAA GGC TGA

nifHR Cyanobacterial nifH4 GCA TAC ATC GCC ATC ATT TCA CC

1 pomati et al. 2004. 2 F. Goh, University of New South Wales, personal communi-cation. 3 nübel et al. 1997. 4 mazarD et al. 2004.

The cyanobacterial community of hot springs 113

cDNA was further amplified using the nifHF and the nifHR primers. The same primer was used to screen our cultures for the nifH gene.

Nitrogen fixation activity was assessed using the acetylene reduction as-say (stewart et al. 1967). Subcultures of 32 mL or 11 mL in case of in situ experiments were placed in rubber-stopped glass bottles, 72 mL and 25 mL respectively. Acetylene (8 mL and 2 mL respectively) was injected into the head space. Control experiments included systems without acetylene and sterile medium with acetylene. To preserve the ethylene/acetylene ratio ob-tained immediately after the in situ incubation was terminated, 2.5 mL of gas from the head-space were transferred to 4 mL vacuum tubes (Vacuette, Greiner Bio-One) containing saturated NaCl solution. Acetylene reduction was assessed by injecting 1 mL of gas taken from the head space into a gas chromatograph equipped with a flame ionization detector (FID-GC SRI 310). To evaluate various factors which affect the N2 fixation abilities of our isolates, we tested activity in: 1) cultures which were initially grown in N-free BG-11 medium (BG-11 0), transferred into fresh BG-110 medium and incubated at 31, 45, 53 and 63 ºC under constant light for 24 h; 2) cultures initially grown in BG-11 medium, transferred to BG-110 two days prior to acetylene reduction assay, and incubated at 45 ºC under constant light and under a 12 h dark/light regime for 48 h; 3) cultures initially grown in BG-11 medium, washed with and transferred to BG-110 immediately prior to acetylene reduction assay initiation and incubated at 48 ºC under a 12 h dark/light regime for 100 h. Cyanobacterial mats were collected from site A1 and from two nearby streams running at a temperature of 51 ºC and 39 ºC. Duplicate samples were incubated in situ at their respective sampling sites under various conditions: 1) in pond/stream water under constant light; 2) dark incubation in bottles covered with aluminum foil; 3) in BG-110 medium under constant light. Incubation time ranged between 1:45–2:45 hours.

The Bunsen gas solubility coefficient for ethylene was calculated for each of the various incubation temperatures according to breithbart et al. (2004). N2 fixation rates were calculated according to capone (1993); a ratio of 4:1 was used to convert reduced acetylene to nitrogen fixed accord-ing to staal et al. (2001) and Jensen & cox (1983). Chlorophyll was ex-tracted either by 90 % methanol at 80 ºC (experiments 1 and 2) or by 90 % acetone (experiment 3).

Results

The microbial mats at sites A1 and A2 (Fig. 1) were dark-green in colour. At sites B1 and B2 and along the channel connecting them, the dominant colour was orange, with prominent patches of green material being present as well. Microscopic examination shows a highly diverse community of cy-

114 D. Ionescu et al.

A B

D C

E F

H G

Fig. 2. Cyanobacteria observed in the Zerka Ma’in springs (a–d) and strains cul-tured from the samples (e–h). A – unicellular cyanobacteria from site A1, B – thin filamentous organisms from site A2, C – spirally wound filaments from site B, D – unicellular, Gloeocapsa-like cells from site B, E – filamentous organisms cultured from site B1, F – Fischerella/Mastigocladus-like filaments, G – cultured Gloeo-capsa-like cells, H – heterocystous filaments. Scale bars = 10 µm.

The cyanobacterial community of hot springs 115

anobacteria, unicellular as well as filamentous (Fig. 2). Site A1 was char-acterized mainly by unicellular organisms (Fig. 2a), while thin filamentous cyanobacteria dominated at site A2 (Fig. 2b). The in situ diversity of cyano-bacteria at site B was higher than that of site A. Spiral filaments reminis-cent of the genus Spirulina were found abundantly at site B (Fig. 2c). The orange coloured mats from site B contained mainly small Gloeocapsa-like unicellular cyanobacteria (Fig. 2d).

After incubation of the environmental samples at 55 °C for periods up to one month, we obtained enrichment cultures with a variety of filamen-tous cyanobacteria (Fig. 2e, 2f). Some isolates resembled those seen in the original samples; others were of types not observed before. An example of the latter is seen in Fig. 2f: Fischerella/Mastigocladus-like filaments, which rapidly became dominant in part of the enrichment cultures. The orange-brown Gloeocapsa-like cells found in the springs (Fig. 2d) were also ob-tained in culture (Fig. 2g). Cultured cells showed a thick capsule, which was less prominent in field-collected material. The filamentous cyanobacteria depicted in Fig. 2h, not observed in the original samples, are of great inter-est because they developed heterocysts.

The 16S rRNA gene sequences obtained from the environmental samples and from the cultures derived from these were aligned and plotted as a tree, using the neighbour joining algorithm and the Mega 3.1 software (Fig. 3). The heterocystous strains designated tBTRCCn 101 and 403 (Fig. 2f, 2h) showed 98 % 16S rRNA gene similarity with Fischerella major, and cluster with filamentous cyanobacteria from thermal environments in Australia and in the Philippines. The unicellular strains tBTRCCn 23 and 28 (Fig. 2g) clus-ter together with Chroogloeocystis siderophila, with which it shows 98 % 16S rRNA gene similarity. The thin filamentous isolate tBTRCCn 302 (Fig. 2e) clusters with other filamentous organisms such as Oscillatoria J24 originating from thermal environments. The Synechococcus-like organisms which thrive in site A1, not yet obtained in culture, were shown by culture-independent techniques to be a diverse cluster of Thermosynechococcus -like organisms (Fig. 3). Other clones clustered together with filamentous cyanobacteria of the Limnothrix type.

We obtained three different nifH sequences (Fig. 4). The sequence from strain nBTRCC 101 and similar isolates showed 94 % similarity with the nifH of Fischerella major (with which it shares 98 % 16S rRNA gene se-quence similarity, see Fig. 3). The other two sequences recovered cluster with nifH of Phormidium sp., Lyngbya sp., and a number of environmental samples from the marine environment. Analysis of environmental mRNA extracted from site A1 revealed two different nifH sequences, one of which has over 99 % sequence identity with one of our filamentous cyanobacteria cultures (tBTRCC24; Fig. 4).

Acetylene reduction assay experiments showed that culture tBTRCCn 101 and similar strains are the only ones capable of N2 fixation. tBTRCCn

116 D. Ionescu et al.

Fig. 3. Phylogenetic tree showing cyanobacterial 16S rRNA gene sequences ampli-fied from the Zerka Ma’in springs and from enrichment cultures obtained from these springs. The sequences were aligned using the ClustalW algorithm and were plotted as a tree using the Neighbour Joining algorithm. Escherichia coli K12 was

The cyanobacterial community of hot springs 117

101 showed high fixation rates at 45 ºC (24.5 nmol N µg chlorophyll-a–1 day–1) and much lower values both at lower and higher temperatures (2.2 and 0.4 nmol N µg chlorophyll-a–1 day–1 at 31 ºC and 53 ºC, respectively). N2 fixation rates under constant light were 10 times lower than when ex-posed to equal length dark/light cycles (0.85 vs. 8.6 nmol N µg chlorophyll- a–1 day–1). Microscopic examination of the cultures at various stages after being transferred from a medium containing combined nitrogen to BG-110 showed that the filaments underwent a morphology change from Mas-

used as outgroup. The enrichment cultures are marked as temporary Bridging the Rift Culture Collection number (tBTRCCn). Selected GenBank accession num-bers are given after the strain designations. Environmental sequences are labeled as clones.

Fig. 4. Phylogenetic tree showing nifH gene sequences obtained from enrichment cultures from the Zerka Ma’in springs, as well as from cDNA collected from site A1 at 63 °C. The sequences were aligned using the ClustalW algorithm and were plotted as a tree using the Neighbour Joining algorithm. Methanothermobacter thermautotrophicus was used as outgroup. The enrichment cultures are marked as temporary Bridging the Rift Culture Collection number (tBTRCCn), and Gen-Bank accession numbers are given after the strain designations. Sequences derived from environmental mRNA amplification are labeled as clones.

118 D. Ionescu et al.

tigocladus-like branched filaments to straight Oscillatoria-like filaments (Fig. 5). The identity of both forms of the organism was confirmed by se-quencing 16S rRNA and nifH genes. When a culture grown in nitrogen-rich medium was transferred to BG-110 medium after extensive washing in the same medium, acetylene reduction started after 48 h, when part of the culture had already changed to the non-branched form.

In situ acetylene reduction assays yielded significant rates in all springs sampled, including at site 1A at 63 ºC. As in the culture experiments, the highest ethylene production was obtained at 51 ºC, with lower rates ob-tained at 63 ºC and 39 ºC. Rates at 51 °C were in the order of 0.0025–0.012 nmol N µg chlorophyll–1 h–1, but even at 63 °C we obtained rates of 0.001–0.0025 nmol N µg chlorophyll–1 h–1, rates significantly higher than back-ground values. Acetylene reduction rates in the light were 30–50 % lower than those measured in the dark (0.0023–0.006 vs. 0.034–0.012 nmol N µg chlorophyll–1 h–1, respectively).

60μm

Fig. 5. Morphological changes upon transfer of isolate tBTRCCn 101 from high-nitrate medium to nitrogen starvation and vice-versa. A – Thick, branched fila-ments in nitrate rich culture, B – Branching of thin filaments 24 h after transfer to BG-110 medium at 48 ºC, C – Branching of thin filaments 24 h after transfer to BG-110 medium at 53 ºC, D – Culture with mixed types of filaments 72 h after transfer to BG-110 medium at 48 ºC, E – Culture with mixed types of filaments 48 h after transfer back to nitrate rich medium. Scale bars = 10 µm.

The cyanobacterial community of hot springs 119

Discussion

Unicellular cyanobacteria generally dominate hot spring communities at temperatures approaching the upper temperature limit for photosynthesis of 74 ºC (castenholz 1969a, brock 1978). Site 1A, which has the high-est temperature of the Zerka Ma’in thermal springs area, shows a fairly diverse community of organisms belonging to the Thermosynechococ-cus clade, as observed microscopically (Fig. 2a) as well as by phylogenetic analyses of samples (Fig. 3). Hot springs at lower temperatures are gen-erally dominated by filamentous cyanobacteria (copelanD 1936, casten-holz 1969a, 1984, brock 1978), and the same is true for site B of the Zerka Ma’in springs (Fig. 2c, 2d). These include yet uncultured spiral cyanobacte-ria (Fig. 2d). Similar tightly coiled filaments were previously reported from Mammoth Hot Springs, Yellowstone (Spirulina cf. labyrinthiformis), where they occurred at an upper temperature limit of 51–52 ºC (castenholz 1977, weller et al. 1992). Phylogenetic analyses of environmental samples thus far failed to reveal any Spirulina -like 16S rRNA gene sequences; how-ever, some of our clones clustered with Limnothrix, a genus that includes a spiral organism (Limnothrix chlorospira), but does not have any known thermophilic species. The nature of the spiral organism in the Zerka Ma’in springs remains to be ascertained.

In previous descriptions of hyperthermal springs, including the Zara springs in Jordan (Deranieh 1990, weller et al. 1992), orange mats were often reported to contain green anoxygenic photosynthetic, Chloroflexus-like organisms. We detected the fatty acids 15:0 and 17:0 in the orange mate-rial, found also in Chloroflexus mats at Yellowstone (zenG et al. 1992). How-ever, a microscopic examination of orange mats in the Zerka Ma’in springs showed prominent presence of a unicellular Gloeocapsa-like cyanobacte-rium (Fig. 2d), with carotenoids probably contributing most to the colour of the mats. These unicellular organisms have been obtained in culture, and their 16S rRNA gene sequence shows up to 99 % similarity with the recently isolated Chroogloeocystis siderophila. This organism was originally found in iron-rich thermal environments in Yellowstone, and requires as much as 30 µM iron for optimal growth (brown et al. 2005). Chemical analyses of the Zerka Ma’in spring waters showed far lower iron concentrations, rang-ing from 0.4 µM (khoury et al. 1984) to 2.2–3.6 µM (rimawi & salameh 1988). At this stage there is no evidence as to the use of siderophores and no measurements of internal iron concentration have been done.

The occurrence of heterocysts in thermophilic filamentous cyanobacteria has been previously documented in Fischerella/Mastigocladus spp. (nierzwicki-bauer et al. 1984, stevens et al. 1985). The isolation of heterocystous Fischerella/Mastigocladus-like filamentous cyanobacteria from the Zerka Ma’in spring (Fig. 2f, 2h, 5) suggests that nitrogen fixation may occur in these springs. Although recent studies show nitrogen fixation

120 D. Ionescu et al.

to take place at temperatures of up to 92 ºC (metha & baross 2007), cyanobacterial nitrogen fixation appears to be limited to much lower temperatures. Mastigocladus-containing microbial mats were shown to fix nitrogen up to a temperature of 55 °C (FoGG 1952, stewart 1970, wickstrom 1980). To further evaluate the nitrogen fixation potential of the microbial community in the Zerka Ma’in thermal springs, we screened our enrichment cultures for the presence of the gene for dinitrogenase reductase, nifH, known as a good phylogenetic marker for nitrogen fixing prokaryotes (zehr et al. 1997). Additionally we sampled the sites themselves for nifH mRNA to check whether any of the amplified sequences obtained from our cultures may match an expressed nifH gene. Although we found mRNA with a nifH sequence over 99 % identical to of one of our cultures (Fig. 4), this culture did not show any nitrogen fixation ability as measured by the acetylene reduction assay under any of the conditions used thus far.

In spite of the short incubation time used (due to experimental constraints), the in situ acetylene reduction assay shows for the first time that nitrogen is being fixed in the themal springs of Zerka Ma’in. The results agree with the data obtained from culture studies, showing a peak in nitrogen fixation at intermediate temperatures (51 ºC). mRNA analyses of these samples are not yet available, and therefore it cannot be ascertained whether the activity was due to cyanobacteria. However, the fact that light inhibited the reaction suggests that the activity was due to an oxygenic phototroph whose nitrogenase was inhibited by the oxygen produced in the light, and may therefore possess a nitrogen fixation strategy which allows for higher rates in the dark. Also in culture experiments much higher acetylene reduction rates were obtained when the culture was exposed to 12 h light/dark cycles than under continuous light.

We observed the branched form of Mastigocladus only in media containing combined nitrogen, and as far as we have been able to ascertain, the change in morphology to unbranched filaments following nitrogen starvation has not been described before. After bound nitrogen was removed from the system, the culture started to fix nitrogen after 48 h. During this induction period the culture went through a morphology change to an unbranched filamentous form (Fig. 5). Microscopic analysis suggests that the process occurs in two stages: first, branching generates thin filaments instead of regular Mastigocladus-type, then these filaments become detached. The morphology change is completed in over a week. Cultures grown from the beginning in BG-110 completely lacked branched filaments. This morphology change can explain why the “classical” Mastigocladus form was not observed in the springs.

Up to now three types of Mastigocladus have been defined. Two strains were characterized as a high temperature type growing at 62–64 ºC (HTF), and a low temperature type grows at 57–58 ºC (MTF) (castenholz

The cyanobacterial community of hot springs 121

1969b, 1972). These two types could also be distinguished by the nature of their branching. A third type (EHN), which incorporates morphological and physiological traits found in the first two types, was described by melick et al. (1991). The latter undergoes morphological changes similar to the Mastigocladus-like cyanobacteria from Zerka Ma’in; however, the trigger for these changes is different. Young cultures of EHN strain are characterized by thin, non-branching filaments, while older cultures resemble the conventional Mastigocladus morphology. The EHN strain is incapable of forming heterocysts and fixing nitrogen, and nitrogen starvation does not induce a morphology change (melick et al. 1991). The Zerka Ma’in organism appears to represent a fourth type, and therefore a more detailed characterization is required.

In conclusion, the diverse cyanobacterial community of Zerka Ma’in consists of organisms similar to those previously described in such springs, as well as novel types. Our findings suggest a higher upper temperature boundary (63 ºC) for cyanobacterial nitrogen fixation than previously published (55 ºC). The physiology and ecology of nitrogen fixation in these springs, including phenomena of light inhibition and oxygen toxicity, remain to be further explored.

Acknowledgements

We thank the Bridging the Rift Foundation for financial support. O.L. was financially supported by an Israeli Ministry of Sciences Fellowship for Women in Science.

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Manuscript received July 11, 2007, accepted October 27, 2007.

The authors’ addresses:

Danny ionescuProf. aharon orenThe Hebrew University of JerusalemDepartment of Plant and Environmental SciencesInstitute of Life Sciences, andThe Moshe Shilo Minerva Center for Marine BiogeochemistryIL-91904 Jerusalem, IsraelE-mail: orena@cc.huji.ac.ilionescu@pob.huji.ac.il

orly levitanDr. ilana berman-FrankBar-Ilan UniversityThe Mina and Everard Goodman Faculty of Life SciencesIL-52900 Ramat Gan, IsraelE-mail: orlylevitan@biu.013.net.ilirfrank@mail.biu.ac.il

Prof. muna hinDiyehDepartment of Water and Environmental Engineering German Jordanian University, Amman, JordanE-Mail: muna.hindiyeh@gju.edu.jo

124 D. Ionescu et al.

Prof. hanan malkawiDepartment of Biological SciencesYarmouk UniversityIrbid, JordanE-mail: hanan@yu.edu.jo, hananmalkawi@gmail.com