Copyright 2010 The Society for Actinomycetes Japan VOL. 24, NO. 2
Actinomycetologica (2010) 24:31–38
Isolation of a Halotolerant Streptomyces sp. from a Constructed Wetland that Biodegrade Phenol and Various Biopolymers Eyal Kurzbaum, Yoram Zimmelsy, and Robert Armon Faculty of Civil & Environmental Engineering, Division of Environmental, Water & Agricultural Engineering, Technion-Israel Institute of Technology, Haifa 32000, Israel (Received Mar. 8, 2010 / Accepted Jun. 17, 2010 / Published Sep. 15, 2010)
In the present study, Streptomyces sp. CW1 was isolated from a constructed wetland system mesocosm and identified as such based on 16S rDNA analysis, and additional biochemical properties were evaluated. This isolate was found to be halotolerant (up to 11% NaCl) and able to grow and utilize biopolymers such as: agarose, agar, gellan gum (a bacterial polysaccharide), polypectate and chitin as the sole carbon and energy source. Since the primary isolation was performed based on its capability to degrade phenol, the high cell yield coefficient showed rapid growth on phenol (0.82–0.98 mg dry biomass/mg phenol). Its halotolerance and the capability to biodegrade biopolymers found mainly in the marine environments, suggest on its primordial oceanic origin. In the present study some of its characteristics are described and discussed along with its beneficial use for wastewater purification processes in constructed wetland systems. proteobacteria, Flavobacterium sp. and Flexibacter-Cytophaga-Bacteroides. The role of actinobacteria phylum in the filed of wastewater treatment was investigated extensively especially in activated sludge systems (Soddell & Seviour, 1990; Wagner et al., 1994; Bond et al., 1999). Nevertheless, there are only few studies that describe the role of actinobacteria in CW systems. Hatano et al. (1994) and El-Shatoury et al. (2004) investigated actinomycetes diversity and role in CW. Hatano et al. (1994) reported that the majority of actinobacteria isolates from pulp-mill wastewater treated by CW were identified as the genus: Streptomyces. Additionally, El-Shatoury et al. (2004) showed that biofilm-attached actinomycetes reflect successful competition with other microorganisms based on their biodiversity. In the present study, microorganisms capable to degrade phenol were isolated from a CW. Phenol as an aromatic contaminant was chosen due to its use in various biodegradation studies and our knowledge on its biodegradation chemistry (Gurujeyalakshmi & Oriel, 1989; Gassner & Neugebohrn, 1994; Van Schie & Young, 2000; Sa´1 & Boaventura, 2001). Along phenol biodegradation experiments (as the sole carbon and energy source) of CW different isolates, it was noticed that a particular bacterium is capable of growing on agar plates without any supplemented carbon source. This capability to grow on agar alone and utilize it as a carbon source is not a prevalent characteristic among terrestrial bacteria but mainly among several marine bacteria. Generally, agaro-
INTRODUCTION Constructed wetland (CW) is an artificial marsh or swamp, created for anthropogenic discharge such as wastewater and stormwater runoff treatment. Commonly it serves as a habitat for wildlife, or for land reclamation after mining and other interferences with nature due to development works. The main biochemical activity in CW occurs by plants and microorganisms presence. Due to this attribute, natural and constructed wetlands act as biofilters removing sediments, heavy metals and organic matter from the polluted incoming water (Kadlec & Knight, 1996). Microorganisms present in CW play a key role in organic carbon removal. These microorganisms exist in two states: free and attached. The attached form is either on rhizoplane or support bed (Kadlec & Knight, 1996; Morris & Monier, 2003; Danhorn & Fuqua, 2007). Their diversity and abundance can vary significantly according to environmental conditions, water quality and flow regimen. The diversity of microorganisms in wetland environment may be critical for proper functioning and maintenance of the system, however, little is known about bacterial populations inhabiting these ecosystems. Ibekwe et al. (2003) while studying bacterial community composition in a CW treating dairy effluents, showed that bacterial community was predominantly composed from phylogenetic clusters related to Bacillus, Clostridium, Mycoplasma, Eubacterium, Proteobacteria, Nitrosospira and Nitrosomonas. Jin & Kelley (2007) found the following microorganisms groups in a pilot scale CW: fungi, protozoa, diatoms, Beta
Corresponding author: Faculty of Civil & Environmental Engineering, Division of Environmental, Water & Agricultural Engineering, Technion-Israel Institute of Technology, Haifa 32000, Israel. E-mail:
[email protected]; Tel: +97248292377; Fax: +97248293309 y We would like to dedicate this article in the memory of Prof. Y. Zimmels that unfortunately passed away prematurely. 31
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lytic bacteria were isolated mainly from oceanic habitats, however several bacteria from terrestrial habitats were also reported (Hosoda et al., 2003). The present study describes the identification and characterization of this agarolytic/ phenol degrader bacterium isolated from CW rhizosphere and the aspects of its biochemical properties according to its habitat are further discussed.
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without phenol was used as control. Inoculated plates were incubated at 30 1 C for 48 hours. Following incubation, white colonies appeared on solid HMM-phenol medium as well on the control medium without phenol. A number of isolated colonies were further plated and transferred several times on solid HMM without phenol. Without phenol, the isolates grew on the solid HMM, revealing the capability of these bacteria to utilize agar as the sole carbon source.
MATERIALS AND METHODS Bacterial growth minimal medium The basic growth media were based on half-strength Hoagland’s minimal medium (HMM) (Hoagland & Arnon, 1950). HMM was prepared by mixing three stock solutions with the following formula: Solution 1- macroelements (g L1 ): KNO3 10.2, Ca (NO3 )2 4H2 O 7.08, NH4 H2 PO4 2.3, MgSO4 7H2 O 4.9 in double distilled water; Solution 2- (g 250 mL1 ): FeSO4 7H2 O 1.9, EDTA-Na2 1.25; and Solution 3-microelements (g L1 ): H3 BO3 2.86, MnCl2 4H2 O 1.81, CuSO4 5H2 O 0.08, H2 MoO4 H2 O 0.09, ZnSO4 7H2 O 0.22. Final Hoagland’s composition was made of solution 1- 100 mL, solution 2- 0.6 mL and solution 3- 1 mL and distilled water to 1 L. pH was adjusted to 6.5 with sterile NaOH (1M).
Identification of the isolated bacteria by 16S rRNA gene analysis Chromosomal DNA was extracted from the isolated bacteria using the alkali-lysis technique, as described by Hartas et al. (1998). Eubacteria-domain-targeted PCR primers were used to amplify a conserved region of eubacterial DNA coding for 16S rRNA (1392 bp), as described previously (Amann et al., 1995). The following primers, purchased from Sigma Genosys (Israel) were used: 11F (50 -GTTTGATCMTGGCTCAG-30 ) and 1392R (50 ACGGGCGGTGTGTAC-30 ). PCR was performed in a Biometra/Tgradient thermocycler. Reactions were carried out in a final volume of 40 ml, with 0.5 pmol of each primer and 20 ml of PCR reaction master mix (Fermentas, Ontario, Canada). PCR conditions were: 95 C for 4 min followed by 36 cycles of 94 C for 30 s, 58 C for 30 s, and 72 C for 1.5 min. The final cycle was followed by extension at 72 C for 10 min. PCR products were visualized on agarose gel (1%) following the purification step, and sent for sequence determination (Hy Laboratories, Rehovot, Israel). Sequence identity was determined with the BLAST program (Altschul et al., 1997).
Isolation of Streptomyces strain from CW Common Reed (Phragmites australis) plants were taken from a CW mesocosm system containing limestone gravel bed as already described (Zimmels et al., 2008). The system was supplemented with diluted domestic sewage (1:10 v/v) for 6 months prior to sampling experiments. To separate rhizospheric bacterial population, 5 g of excised root sections were vortexed in 200 mL sterile saline (0.85% NaCl) containing Tween 20 (0.1%) (Sigma, Israel) for 60 minutes at 200 rpm (temperature 22 2 C) on a rotatory shaker (MRC, TS-400). The suspension was pre-filtered through a sterile cheese-cloth to remove large particles, the filtrate was centrifuged at 6,000 g for 10 minutes and the pellet washed three times with saline. Finally, 50 mL of resuspended pellet was inoculated into sterile 300 mL flasks containing 100 mL sterile HMM supplemented with 20 mg L1 phenol (Merck, USA) as the sole carbon and energy source. The inoculated flasks were shaken (100 rpm) for one month at 30 C, supplemented daily with 20 mg L1 phenol. From the experimental results (data not shown) phenol at this concentration was biodegraded completely in 24 hours, therefore daily addition of phenol was requested to maintain a continuous selective environment. Weekly, the growing suspended bacterial cells were centrifuge and washed with saline three times. The pellet was used as inoculum in a fresh medium containing phenol as mentioned above. After one month, five samples from each subculture were spread on solid HMM (addition of 15 g L1 Bacto agar, Difco, USA) containing 50 mg L1 phenol as the sole carbon and energy source. Solid HMM medium
Morphology, oxygen requirement, and salt tolerance of bacterial isolates The isolated bacterium was grown on nutrient agar (Difco), starch-casein agar (APHA, 1995) and nutrient broth (Difco) for 24 hours at 30 C. Isolated colonies were Gram stained and examined for morphology with Zeiss Axiolab epifluorescent microscope ( 1,000 magnification). Oxygen requirement of the bacterial isolate was performed by anaerobic growth. Nutrient agar stroke with bacterial isolates were incubated in BBL jars containing GasPak anaerobic system (Becton-Dickinson, Cockeysville, MD) at 30 C up to 7 d. Halotolerance was investigated by incubation of the isolated bacteria into sterile 100 mL nutrient broth containing various concentrations of NaCl (Sigma) (0 to 15%) incubated on rotary shaker at 30 C (100 rpm). Bacterial growth was monitored daily by increase in optical density (OD600 nm) and Gram staining for identification. Antibiotic susceptibility test Antibiotic susceptibility tests were performed on the isolated strain using disk diffusion technique on nutrient 32
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cultures were incubated at 30 C under continuous shaking (150 rpm). Cell growth was determined by measuring OD at 600 nm using a spectrophotometer (Spectronic 20 Genesys, Spectronic Instruments, USA) at different time intervals. Phenol concentrations were measured periodically using the 4-aminoantipyrine colorimetric approach (APHA, 1995) on supernatant of samples centrifuged at 6,000 g for 10 min. Each experimental culture dry biomass was measured as already described (APHA, 1995). Phenol degradation rate (mg L1 h1 ) and cell yield coefficient (mg cell/mg phenol) of the biodegradation processes were determined by a method reported by Pirt (1975). Phenol degradation rate was determined from the logarithmic slope of phenol concentration (mg L1 ) versus time. Cell yield coefficient was calculated as the dry cell mass per mass of phenol utilized.
agar (Difco) (Bauer et al., 1966). Commercially available disks (Difco) were applied as recommended, and the plates were incubated at 37 C for 48 h. The antibiotic types (and concentration in mg) were as follows: doxycycline (30), spectinomycin (100), colistin (polymyxin E) (10), ceftriaxone (30), amikacin (30), sulfamethoxazole trimethoprim (23.75 and 1.25, respectively), nalidixic acid (30), amoxicillin/clavulanic acid (30), erythromycin (5), chloramphenicol (30), tetracycline (30), rifampin (5), ampicillin (10), gentamicin (10), penicillin (30), kanamicine (30), polymyxin B (300), and cycloheximide (100). Biopolymers degradation tests According to section ‘‘Isolation of Streptomyces strain from CW’’ the isolated bacterium able to grow on agar minimal medium was further tested for its capability to biodegrade additional biopolymers such as: agarose, polypectate, chitin, gellan gum, starch, and different agar brands as the sole carbon source. These biopolymers were prepared as follows: agarose (high resolution agarose, Quantum Appligene), Bacto agar (Difco) and Noble agar (Difco) (15 g L1 ) were added to 1 liter HMM, and Bacto agar (15 g L1 ) in double distilled water. Polypectate (Sigma, USA) in HMM was prepared as described in Pickaver (1977). Gellan gum (Gel-Gro, ICN Biochemicals, Cleveland, OH) was made by addition of 8 g Gel-Gro powder with 1 g of MgSO4 7H2 O into 1 L HMM, heating to boiling point while stirring and autoclave. Chitin medium was prepared as previously described (Hsu & Lockwood, 1975; Pleban et al., 1997). Briefly, colloidal chitin (Sigma, USA) was suspended in HMM (4 g L1 ) and 15 g L1 Bacto agar added. Following sterilization, the hot medium pH was adjusted to 8.0 with sterile NaOH (1 M) through continuous stirring and finally dispersed into Petri dishes at 50 C. Starch agar was prepared by addition of 2 g starch (Merck, USA) and 15 g Bacto agar to 1 L HMM. Starch hydrolysis was demonstrated by Gran’s test as described elsewhere (e.g. Stanier, 1942). All media were sterilized by autoclave at 121 C (15 psi) for 17 minutes, cooled to 50 C and equally divided into Petri dishes (90 mm). The capability of newly isolated experimental bacteria to grow on the various biopolymer sources was examined by direct inoculation and additional incubation at 30 C for 4 d.
Colonization of Zea mays root by Streptomyces sp. CW1 observed with Confocal Laser Scanning Microscopy (CLSM) Corn (Zea mays) (Gedera Seeds Co., Gedera, Israel) seeds were surface sterilized for 10 min in 70% ethanol and 1 min in 2% sodium hypochlorite and grown in a sterile hydroponic chamber as previously described (Burdman et al., 1996). 7 days old germinated sterile seedlings (approximately 4 cm long roots) were soaked for 5 minutes in HMM containing Streptomyces sp. CW1 suspension (107 CFU/mL) previously grown in HMM supplemented with 50 mg L1 phenol. Inoculated seedlings grown in the hydroponic chamber for 3 additional days were examined for surface adsorption of bacterial cells with confocal laser scanning microscope (CLSM). Adsorbed bacteria were stained by soaking the seedlings in 0.01% acridine orange for ten minutes and rinsed three times in sterile HMM. Confocal microscopy of the adsorbed bacteria onto rhizosphere were performed by CLSM (model: CTR 5500CS microscope, Leica, Germany) equipped with detectors and filter sets that simultaneously monitored red and green fluorescence. The green autofluorescence exhibited by the root material was used to visualize the root surface (for further details see Assmus et al., 1995). Image combination and process were performed with the standard software package provided by Leica (Leica application suite 3.0). RESULTS AND DISCUSSION Characteristics of the isolated bacteria and colony morphology The CW bacterial rhizospheric isolate CW1 is a filamentous, Gram (+) bacterium that resembles morphological characteristics of Streptomyces group. Single cell has a rod morphology with rounded ends (1:5 0:6 mm), while the hyphae are long and branched (between few to tens mm long) (Fig. 1). In some cases spores connected to the edge of the hyphae were observed. Colonies appearance on nutrient agar and other media is round and floury white
Phenol degradation kinetic The isolate was grown in 100 mL HMM supplemented with 100 mg L1 phenol as sole carbon source for 12 hours on a rotary shaker (150 rpm) at 30 C. The incubated culture was centrifuged and washed three times with sterile saline (0.85% NaCl). For kinetic experiments, inoculums from the phenol acclimatized culture were divided into 400 mL flasks. Each flask contained the following phenol concentrations of 5, 10, 30 and 50 mg L1 in 100 mL HMM solution (experiments were done in triplicates). The 33
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Fig. 2. (Color online) Colony morphology of the isolated Streptomyces sp. CW1 on nutrient agar plate (inset: enlargement of few colonies showing floury appearance of aerial mycelia and spores).
Fig. 1. (Color online) Gram staining of Streptomyces sp. CW1 from liquid medium (A) and agar plate (B) ( 1000, Zeiss, Axiolab). Singular cells, filaments and aerial mycelia are visible, which is characteristic of Streptomyces. Fig. 3. (Color online) In situ visualization of Streptomyces sp. CW1 cells on the root surfaces of Zea mays seedling. The red fluorescence emitted by the Streptomyces sp. CW1 cells and the green autofluorescence emitted by the root material were visualized by CLSM.
with strong hold into the solid agar (Fig. 2). In liquid culture the bacterium usually revealed hypheal growth, forming granules of 0.5 to 3 mm in diameter accompanied by solution clearing. The bacterium did not grow when incubated under anaerobic conditions. Therefore the CW1 was identified as a member of the genus Streptomyces based on the morphological and physiological characteristics. As some CW systems receive a variety of effluent qualities, salinity may increase as a result of prolonged retention time as well continuous evapotranspiration. In the present experimental set-up, the new isolated Streptomyces sp. CW1 exhibited halotolerance. This characteristic is not unusual for Streptomyces (up to 7%) (Buchanan & Gibbons, 1974), however the upper halotolerance of the new isolate was 11% (w/v) (data not shown). At 11% NaCl, growth was delayed but still measurable. Increasing the NaCl concentration in the growth media caused the granules to have a more compressed appearance which was associated with faster sinking characteristics. Streptomyces sp. CW1 revealed good attachment properties onto various materials such as glass, activated carbon, lime-stone gravel, textile fibers (data not shown), and plant roots (Fig. 3).
Spores originating from growing culture on solid media, were highly hydrophobic (floating on water surface) while those grown in liquid medium were equally suspended in the whole water volume. Spores collected from dried agar layer remained viable for more then one month at 36 C and grew well when retransferred to fresh liquid/solid media. Identification of the isolated bacteria by 16S ribosomal RNA analysis The 16S rRNA sequence used for similarity search using the BLAST program indicated a relatively close relationship to the DNA sequence of many species from the genus Streptomyces (94% identity). Extra specific identification of the new isolate to species and strain levels was outside the scope of the present study. The sequence data have been submitted to the GenBank database under accession number GU132307. 34
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Table 1.
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Utilization of different biopolymers by Streptomyces sp. CW1
Biopolymer name (experimental medium) Agarose (HMM) Bacto agar (HMM) Noble agar (HMM) Bacto agar (DDW)b Polypectate (HMM) Starch (HMM) Chitin (HMM) Gellan gum (HMM)
Characteristic Structure linear polymer of alternating D-galactose and 3,6-anhydrogalactose units linear polymer of alternating D-galactose and 3,6-anhydrogalactose units and -1,3-glycosidically linked D-galactose units linear polymer of alternating D-galactose and 3,6-anhydrogalactose units and -1,3-glycosidically linked D-galactose units linear polymer of alternating D-galactose and 3,6-anhydrogalactose units and -1,3-glycosidically linked D-galactose units linear chain of D-galacturonic acid glucose monosaccharide units joined together by glycosidic bonds N-acetyl-D-glucos-2-amine L-rhamnopyranosyl-D-glucopyranosyl-D-glucuronopyranosyl-D-glucopyranosyl
Growth efficiencya ++ +++ ++ + ++ +++ +++ +++
a)
- Growth efficiency was measured qualitatively by relative coverage with Streptomyces sp. CW1 colonies of the petri dish: +, low growth; ++, medium growth; +++, intensive growth. b) - DDW, double distilled water
aquatic bacteria involves pectin degrading enzymes, and Gummadi & Kumar (2005) showed that Streptomyces spp. owe this properties. Streptomyces sp. CW1 was grown on a bacterial biopolymer (gellan gum). This biopolymer is a low-acyl microbial polysaccharide produced by the bacterium Sphingomonas paucimobilis that is generally resistant to bacterial enzymatic degradation (Lin & Casida, 1984; Bajaj et al., 2007). In its highly-purified form natural anionic heteropolysaccharide is composed of glucose, glucuronic acid and rhamnose moieties. Experimental results showed that Streptomyces grew efficiently and even better compared to agar (Table 1). Thus far, only three bacterial species: Bacillus sp., Sphingomonas sp., and Geobacillus sp., were reported to biodegrade gellan gum polysaccharide (Sutherland & Kennedy, 1996; Hashimoto et al., 1997; Derekova et al., 2006). In the present study, Streptomyces sp. isolate showed capability to utilize gellan gum as a carbon source. Gellan gum and other microbial biopolymers have many attractive pharmaceutical, chemical, and food-industrial applications (Giavasis et al., 2000), therefore lyases probably secreted by the newly isolated bacterium, may have potential industrial applications.
Antibiotics susceptibility tests Streptomyces sp. CW1 was subjected to a variety of antibiotic disks at various concentrations revealed that among the eighteen antibiotics tested, sulfamethoxazole trimethoprim, nalidixic acid, polymixin B sulfate, and cycloheximide did not inhibit its growth. Growth of the Streptomyces sp. on diverse biopolymers Streptomyces sp. CW1 grew on agar plates without any supplementary carbon sources (Table 1). The agarolytic property is mostly uncommon among terrestrial bacteria but mainly common among marine bacteria (Belas et al., 1988; Hosoda et al., 2003). The rationale of this biochemical characteristic can be explained by agar production of some marine algae. Agar is a mixture of a neutral, dominating polysaccharide called ‘‘agarose’’ and a charged polymer called ‘‘agaropectin’’. Agar breakdown occurs through an extracellular enzyme called -agarase (Belas et al., 1988). Among the genus Streptomyces, Streptomyces coelicolor was found to have agarolytic capability (Parro & Harwood, 1998). Agarolytic and halotolerance properties of the Streptomyces sp. CW1 isolate allude on its possible marine origin in spite of its terrestrial habitat. Additional biodegradation experiments with Streptomyces sp. CW1 on colloidal chitin agar, revealed clear zones surrounding the colonies (Table 1). Growth in HMM liquid medium containing chitin was also observed (data not shown). Chitinase activity was reported for phylum Actinobacteria to which Streptomyces belong (Reynolds, 1954; Hsu & Lockwood, 1975; Watanabe et al., 1999). Streptomyces sp. CW1 was found also to biodegrade polypectate (Table 1). Pectin degrading enzymes include several types and are commonly found in soil and plant pathogenic bacteria (Grac¸a et al., 2005). Suberkropp & Klug (1980) found that decomposition of leaf litter by
Biodegradation of phenol Phenol degradation kinetics are shown in Fig. 4. Phenol initial concentration ranged from 5 to 50 mg L1 and the bacterial inocula were 0.013 OD (35 mg dry biomass/L). An interesting kinetic behavior of the biomass was observed at different initial phenol concentrations. When initial phenol concentrations of 5 and 10 mg L1 were applied, bacterial biomass continued to increase even when phenol concentration was 0 mg L1 . At 30 and 50 mg L1 initial phenol concentrations, bacterial biomass stopped to grow when phenol concentration reached 0 mg L1 . In the present 35
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Fig. 4. Phenol degradation at various initial concentrations and biomass formation by Streptomyces sp. CW1 (A) 5 mg L1 , (B) 10 mg L1 , (C) 30 mg L1 , and (D) 50 mg L1 . Note: Figures B and D include a control (phenol without bacteria).
study, phenol concentrations of 5 to 50 mg L1 are relatively low compared to previous studies where bacteria were exposed to high concentrations mainly for field bioreactors and possible industrial utilization (Mo¨rsen & Rehm, 1987; Peyton et al., 2002; Kumar et al., 1992). The experimental phenol concentrations applied in the present study are typical to effluents feed of CW worldwide (Kadlec & Knight, 1996; Polprasert et al., 1996; Abira et al., 2005). According to published literature on phenol biodegradation, it can be assumed that in the case of the newly isolated Streptomyces sp. at phenol concentrations of 5 and 10 mg L1 the intermediate compounds such as catechol are inhibitory. The intermediary compound originating from the oxygenation of the phenol aromatic ring through monohydroxylation by a mono oxygenase phenol hydroxylase at ortho position to the pre-existing hydroxyl group to form catechol (Agarry et al., 2008), does not inhibit bacterial growth (biomass increase), while at 30 and 50 mg L1 the inhibitory effect is much more pronounced. At 50 mg L1 phenol, a significant lag in biomass growth was observed. Similar phenomenon of inhibition due to high phenol concentration for an actinobacteria strain was reported by Straube et al. (1990). Generally, phenol
Table 2. Cell yield coefficient and degradation rate of phenol by Streptomyces sp. CW1 Initial Phenol concentration (mg L 1 )
Cell yield coefficient (mg dry biomass/ mg phenol)
Phenol degradation rate (mg L 1 h 1 )
5 10 30 50
0.82 0.90 0.90 0.98
0.94 2.96 7.85 8.41
biodegradation by pure cultures of bacteria has been adequately described by substrate inhibition models (Kumaran & Paruchuri, 1997). The phenol degradation rate increased proportionally with the initial phenol concentration (Table 2). Cell yield coefficient (mg dry biomass/mg phenol) versus phenol concentration ranges showed also an increase from 0.82 for 5 mg L1 to 0.98 for 50 mg L1 . The cell yield coefficient values calculated in this study are higher than other phenol degrading bacteria as already reported by Feitkenhauer et al. (2001). 36
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vated sludge systems. Appl. Environ. Microbiol. 65, 4077– 4084. Buchanan, R.E. & Gibbons, N.E. (1974). Bergey’s manual of determinative bacteriology, 8th ed. Williams Wilkins, Baltimore, MD. Burdman, S., Volpin, H., Kigel, J., Kapulnik, Y. & Okon, Y. (1996). Promotion of nod gene inducers and nodulation in common bean (Phaseolus vulgaris) roots inoculated with Azospirillum brasilense Cd. Appl. Environ. Microbiol. 62, 3030–3033. Danhorn, T. & Fuqua, C. (2007). Biofilm formation by plantassociated bacteria. Annu. Rev. Microbiol. 61, 401–422. Derekova, A., Sjoholm, C., Mandeva, R., Michailova, L. & Kambourova, M. (2006). Biosynthesis of a thermostable gellan lyase by newly isolated and characterized strain of Geobacillus stearothermophilus 98. Extremophiles 10, 321– 326. El-Shatoury, S., Mitchell, J., Bahgat, M. & Dewedar, A. (2004). Biodiversity of actinomycetes in a constructed wetland for industrial effluent treatment. Actinomycetologica 18, 1–7. Feitkenhauer, H., Schnicke, S., Mu¨ller, R. & Ma¨rkl, H. (2001). Determination of the kinetic parameters of the phenol-degrading thermophile Bacillus themoleovorans sp. A2. Appl. Microbiol. Biotechnol. 57, 744–750. Gassner, W. & Neugebohrn, L. (1994). The significance of higher plants for degradation of phenols in aquatic systems. Arch. Hydrobiol. 129, 473–495. Giavasis, I., Harvey, L.M. & McNeil, B. (2000). Gellan gum. Crit. Rev. Biotechnol. 20, 177–211. Grac¸a, M.A.S., Ba¨rlocher, F. & Gessner, M. (2005). Methods to study litter decomposition: A practical guide. Springer, The Netherlands. Gummadi, S.N. & Kumar, D.S. (2005). Microbial pectic transeliminases. Biotechnol. Lett. 27, 451–458. Gurujeyalakshmi, G. & Oriel, P. (1989). Isolation of phenoldegrading Bacillus stearothermophilus and partial characterization of the phenol hydroxylase. Appl. Environ. Microbiol. 55, 500–502. Hartas, J., Hibble, M. & Sriprakash, K.S. (1998). Simplification of a locus-specific DNA typing method (Vir Typing) for Streptococcus pyogenes. J. Clin. Microbiol. 36, 1428–1429. Hashimoto, W., Maesaka, K., Sato, N., Kimura, S., Yamamoto, K., Kumagai, H. & Murata, K. (1997). Microbial system for polysaccharide depolymerization: enzymatic route for gellan depolymerization by Bacillus sp. GL1. Arch. Biochem. Biophys. 339, 17–23. Hatano, K., Frederick, D. & Moore, J. (1994). Microbial ecology of constructed wetlands used for treating pulp-mill Wastewater. Water Sci. Technol. 29, 233–239. Hoagland, D.R. & Arnon, D.I. (1950). The water-culture method for growing plants without soil. Circular 347. Agricultural Experiment Station, University of California, Berkeley. Hosoda, A., Sakai, M. & Kanazawa, S. (2003). Isolation and characterization of agar-degrading Paenibacillus spp. associated with the rhizosphere of spinach. Biosci. Biotechnol. Biochem. 67, 1048–1055. Hsu, S.C. & Lockwood, J.L. (1975). Powdered chitin agar as a selective medium for enumeration of Actinomycetes in water and soil. Appl. Microbiol. 29, 422–426. Ibekwe, M., Grieve, C.M. & Lyon, S.R. (2003). Characterization of microbial communities and composition in constructed dairy
CONCLUSIONS The newly isolated Streptomyces sp. CW1 revealed environmentally interesting features such as: phenol degradation, halotolerance, and utilization of biopolymers as the sole carbon source (agar, agarose, chitin, pectin, gellan gum, and starch). These characteristics may point on its previous marine origin with later on adaptation to terrestrial environments. Moreover, its presence in CW rhizosphere, harboring biodegradability potential towards a large variety of biopolymers, may suggest that this euryhaline bacterium can be useful in CW with variable salinity and still reduce anthropogenic contaminants present in these systems. ACKNOWLEDGMENTS This research was partially supported by a fund from Keren Kayemeth LeIsrael, Faculty of Civil & Environmental Engineering, and Grand Water Research Institute at Technion. REFERENCES Abira, A., Van Bruggen, J.J. & Denny, P. (2005). Potential of a tropical subsurface constructed wetland to remove phenol from pre-treated pulp and papermill wastewater. Water Sci. Technol. 51, 173–176. Agarry, S.E., Durojaiye, A.O. & Solomon, B.O. (2008). Microbial degradation of phenols: a review. Int. J. Environ. Pollut. 32, 12–28. Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W. & Lipman, D.J. (1997). Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402. Amann, R.I., Ludwig, W. & Schleifer, K.H. (1995). Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143–169. APHA 1995. Standard methods for the examination of water and waste water, American Public Health Association, 19th ed., Washington. Assmus, B., Hutzler, P., Kirchhof, G., Amann, R., Lawrence, J.R. & Hartmann, A. (1995). In situ localization of Azospirillum brasilense in the rhizosphere of wheat with fluorescently labeled, rRNA-targeted oligonucleotide probes and scanning confocal laser microscopy. Appl. Environ. Microbiol. 61, 1013– 1019. Bajaj, I.B., Survase, S.A., Saudagar, P.S. & Singhal, R.S. (2007). Gellan gum: fermentative production, downstream processing and applications. Food Technol. Biotechnol. 45, 341–354. Bauer, A.W., Kirby, W.M.M., Sherris, J.C. & Turck, M. (1966). Antibiotic susceptibility testing by a standardized single disk method. Am. J. Clin. Pathol. 45, 493–496. Belas, R., Bartlett, D. & Silverman, M. (1988). Cloning and gene replacement mutagenesis of a Pseudomonas atlantica agarase gene. Appl. Environ. Microbiol. 54, 30–37. Bond, P.L., Erhart, R., Wagner, M., Keller, J. & Blackall, L.L. (1999). Identification of some of the major groups of bacteria in efficient and nonefficient biological phosphorus removal acti37
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