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Isolation, characterization and localization ofextracellular polymeric substances from thecyanobacterium Arthrospira platensis strain MMG-9Mehboob Ahmedab, Tanja C.W. Moerdijk-Poortvlietb, Anita Wijnholdsb, Lucas J. Stalbc &Shahida Hasnainad

a Department of Microbiology and Molecular Genetics, Quaid-e-Azam Campus, Universityof the Punjab, Lahore-54590, Pakistanb Department of Marine Microbiology, Royal Netherlands Institute of Sea Research (NIOZ),NL-4400 AC Yerseke, The Netherlandsc Department of Aquatic Microbiology, IBED, University of Amsterdam, 1090 GEAmsterdam, The Netherlandsd The Women’s University Multan, Multan, PakistanPublished online: 25 Apr 2014.

To cite this article: Mehboob Ahmed, Tanja C.W. Moerdijk-Poortvliet, Anita Wijnholds, Lucas J. Stal & Shahida Hasnain(2014) Isolation, characterization and localization of extracellular polymeric substances from the cyanobacteriumArthrospira platensis strain MMG-9, European Journal of Phycology, 49:2, 143-150, DOI: 10.1080/09670262.2014.895048

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Isolation, characterization and localization of extracellularpolymeric substances from the cyanobacterium Arthrospiraplatensis strain MMG-9

MEHBOOB AHMED1,2, TANJA C.W. MOERDIJK-POORTVLIET2, ANITAWIJNHOLDS2,LUCAS J. STAL2,3 AND SHAHIDA HASNAIN1,4

1Department of Microbiology and Molecular Genetics, Quaid-e-Azam Campus, University of the Punjab, Lahore-54590,Pakistan2Department of Marine Microbiology, Royal Netherlands Institute of Sea Research (NIOZ), NL-4400 AC Yerseke, TheNetherlands3Department of Aquatic Microbiology, IBED, University of Amsterdam, 1090 GE Amsterdam, The Netherlands4The Women’s University Multan, Multan, Pakistan

(Received 28 July 2013; revised 16 December 2013; accepted 13 January 2014)

Arthrospira platensis is a cyanobacterium known for its nutritional value and secondary metabolites. Extracellular polymericsubstances (EPS) are an important trait of most cyanobacteria, including A. platensis. Here, we extracted and analysed differentfractions of EPS from a locally isolated strain of A. platensis. Three different fractions of EPS were distinguished. These were EPSreleased into the medium (REPS), EPS loosely bound to the organism (LEPS) and EPS tightly bound to the organism (TEPS),which were extracted by different procedures. The LEPS fraction was smaller than the other two fractions. The EPS of A. platensisexhibited high diversity. Total protein and carbohydrate content was determined in each of these fractions. The largest amount oftotal carbohydrates and total proteins was in the TEPS fraction. Eight sugar moieties were detected and analysed in all EPSfractions using HPAE-PAD. Fructose, mannose and ribose were rare sugar residues in all fractions of EPS. With the exception offructose, all sugars tested for were detected in TEPS. The amount of sugars detected was significantly higher in TEPS comparedwith the two other fractions, especially for galactose, xylose and glucose. The EPS were localized by confocal laser scanningmicroscopy (CLSM) after staining with different fluorescent dyes and it was found that A. platensis possessed a thick and smoothlayer of EPS around the spiral trichomes.

Key words: Arthrospira platensis, confocal laser scanning microscopy, extracellular polymeric substances, lectins, monosac-charide analysis

Introduction

Extracellular polymeric substances (EPS) are impor-tant compounds produced by many cyanobacteria;they surround the organisms as sheaths, capsules ormucilage. EPS provide protection from various bioticand abiotic stresses and facilitate the attachment of theorganism to a solid substrate (Palmer et al., 2007;Pereira et al., 2009; Jittawuttipoka et al., 2013).Generally, EPS are categorized into three types:sheaths, capsules and mucilage, based on their loca-tion, structure and consistency. Capsules are consid-ered to be highly structured, strictly delimited andtightly attached to the organism, whereas sheaths areless structured, not delimited, of varying thickness andnot so intimately attached to the organism.Mucilage isunstructured and loosely associated with the organism

or freely dispersed into the environment (Richertet al., 2005). In recent years, there has been greatinterest in the exploitation of EPS by various indus-trial sectors because of the chemical and physicalproperties of these polymers. For instance, EPS andEPS-producing microorganisms have been used forsoil conditioning (Abed et al., 2009).

Arthrospira platensis (previously known asSpirulina platensis) belongs to the familyOscillatoriaceae and is a filamentous cyanobacteriumwith spiral trichomes. It is one of the best-studied andmost widely used cyanobacteria in biotechnologybecause of its fast growth in open ponds under alka-line conditions that largely exclude contaminatingorganisms (Ahmed et al., 2010b). Arthrospira platen-sis has enormous potential for applications in a varietyof fields such as food, cosmetics, environmentalimprovement, aquaculture and in the future perhapstherapeutics (Rossi et al., 2005; Trabelsi et al., 2009).Correspondence to: Mehboob Ahmed. E-mail: mehboob.

mmg@pu.edu.pk

Eur. J. Phycol. (2014), 49(2): 143–150

ISSN 0967-0262 (print)/ISSN 1469-4433 (online)/14/020143-150 © 2014 British Phycological Societyhttp://dx.doi.org/10.1080/09670262.2014.895048

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Arthrospira platensis produces many metabolites thatare either unique or found in higher concentrations inthis organism than others and thus may be producedeconomically (Cohen, 1997). As is the case for manyother cyanobacteria, Arthrospira secretes a variety ofdifferent EPS that can be extracted and used in differ-ent fields (Trabelsi et al., 2009), for instance, arthros-piral EPS has been reported to enhance human innateimmunity (Lobner et al., 2008; Parages et al., 2012).The objective of the present work was (1) to extractdifferent EPS fractions and characterize them bio-chemically and (2) to visualize the EPS of A. platensiswithout disturbing its structural integrity by usingvarious lectins and confocal laser scanningmicroscopy.

Materials and methods

Strain and growth conditions

Arthrospira platensis MMG-9 (NCBI GenBank Accessionnumber FJ839360 for the 16S rRNA gene sequence) wasisolated from a rice field in Pakistan (Ahmed et al., 2010b).This Arthrospira strain was cultured and maintained inSpirulina medium (George, 1976) at 25°C at a 16 : 8 hlight-dark cycle (200 µmol photons m−2 s−1). Cultures wereharvested after 3 weeks of incubation for extraction of EPS.

Growth measurement

For estimation of growth, chlorophyll a was determined fol-lowing the method of Tandeau de Marsac & Houmard (1988).Cells were harvested by centrifugation at 10 000 × g for 10minat 4°C. The pellets were extracted with 80% methanol for 2 hin the dark at 4°C. The extract was centrifuged at 10 000 × gfor 10 min at 4°C and the absorbance of the supernatant wasmeasured spectrophotometrically at 665 nm against 80%methanol. Chlorophyll content (µg ml−1) was calculatedfrom the absorption at 665 nm (OD665nm) × 13.9.

Extraction of EPS

EPS were extracted according to Staats et al. (2000) withmodifications. The EPS can be tightly bound and remainattached to the cells, or released into the medium as REPSwhich can be recovered from the supernatant. Tightly boundEPS were extracted from the cells in two steps. Firstly, theloosely bound EPS (LEPS) were obtained by shaking inlukewarm water and secondly, the remaining tightly boundEPS (TEPS) was extracted with 0.1 M EDTA.

Arthrospira platensis was grown for 3 weeks in 1 litreculture flasks. Cultures were harvested by centrifugation at5000 × g for 20 min at 20°C. For REPS, the supernatant wasfiltered through membrane filters (Millipore, 0.45 µm), freeze-dried, weighed and stored at −20°C until further analysis. ForLEPS, 4.5 ml of MilliQ water was added to the cell pellet,vortexed for 1 min and subsequently incubated in a shakingwater bath (120 rpm) at 30°C for 1 h. The cells were checkedmicroscopically to ensure that they were not damaged.Subsequently, the extracts were centrifuged at 5000 × g for20 min at 20°C. The supernatant containing the LEPS was

filtered through membrane filters (Millipore, 0.45 µm), freeze-dried, weighed and stored at −20°C until further analysis. Forthe extraction of TEPS, the pellets resulting from the LEPSextraction were re-suspended in 4.5 ml of 0.1 M EDTA andincubated for 4 h in a shaking (120 rpm) water bath at 25°C.The cells were checked microscopically to ensure that no celllysis had occurred, in order to minimize contamination of theextracts with intracellular carbohydrate. The extracts werecentrifuged for 20 min at 5000 × g at 20°C and the supernatantcontaining the TEPS was filtered through a membrane filter(Millipore, 0.45 µm), freeze-dried, weighed and stored at−20°C until further analysis.

Measurement of total carbohydrates, total proteins,uronic acids and glycosaminoglycans (GAGs)

Known amounts of the three fractions of EPS were dissolvedin MilliQ water. Total sugar content was determined accord-ing to Dubois et al. (1956); total protein content was mea-sured with the Bradford Protein Assay (Bio RadLaboratories, Veenendaal, the Netherlands); uronic acidswere measured with carbazole in 80% sulphuric acid withborate ions added (0.8% sodium borate – tetrahydrate andconcentrated sulphuric acid mixed in ratio of 17 : 83) (Taylor& Buchanan-Smith, 1992), and glycosaminoglycan wasmeasured with the dimethyl methylene blue method(Chandrasekhar et al., 1987).

Monosaccharide analysis

Monosaccharide composition of the three EPS fractions wasanalysed qualitatively and quantitatively using HighPerformance Anion Exchange Chromatography with PulsedAmperometric Detection (HPAE-PAD). Known quantities ofEPSwere suspended in 4.5 mlMilliQ water and subsequentlyhydrolysed by adding 0.5 ml of 11 M H2SO4 to 4.5 ml EPSsolution in a glass vial and incubating for 1 h at 120°C.Immediately after hydrolysis the glass vials were put on ice,BaCO3 was added to neutralize the solution and the vialswere vigorously shaken. The pH of the solution was mea-sured and BaCO3 was added until the pH reached 5–6. Thehydrolysed solution was centrifuged for 20 min at 10 000 × gand 4°C, then the supernatants were filtered through a0.22 µm filter (Millex-GV4; Millipore, Bedford, MA, USA)and stored in 1 ml glass vials at −20°C until further analysis(Boschker et al., 2008).

HPAE-PAD was carried out on a Dionex System (Dionex,Breda, the Netherlands) consisting of an HPLC pump(Dionex, P580), an autoinjector (Dionex, ASI-100) and anelectrochemical detector (Dionex, ED40) fitted with aCarboPac PA20 guard and narrow-bore analytical column(3×150 mm; Dionex, Breda, the Netherlands) and eluted iso-cratically with 300 µl min−1 of 1 mMNaOH. The column wasregenerated regularly with 200 mM NaOH. All eluents weredegassed with helium before NaOH was added and furtherdegassed with helium during analysis. All pump heads wererinsed at least once a day to prevent crystallization. Peaks wereanalysed with ChromeleonC Dionex Version 6.80 SR5 Build2413. Peak identification was based on retention times incomparison with external standards. Concentration measure-ments were based on peak areas of the separated compoundsand calibrated against external standards.

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Localization of EPS by CLSM

EPS were localized using a confocal laser scanning micro-scope (Leica TCS-NT, Leica Lasertechnik Heidelberg,Germany) equipped with an Argon-Krypton laser and usingthe 564 and 488 nm lines. Leica 25 (NA 0.75) oil immersionand Leica 63 (NA 0.9) water immersion objectives were used(Ahmed et al., 2010a).

Autofluorescence of cyanobacterial chlorophyll was usedfor observation of the cells and the different components ofEPS were stained using a variety of lectins and fluorescentdyes (Table 1). Multi-fluorochrome staining was performedusing fluorescent dyes in series. Eighteen-day-old cultureswere harvested by centrifugation for 10 min at 10 000 × g inEppendorf tubes and the pellets were treated with thefluorescent dye according to the manufacturer’s instructions.Excitation wavelengths and emission filters of the CLSMwereadjusted according to the specificity of fluorescent dye.

Results

Extraction of EPS

Extracted EPS appeared as a white material. A total of561 mg EPS (g Chl a)−1 was extracted. Extraction andquantification of the three EPS fractions of A. platen-sis MMG-9 revealed that TEPS was the most abun-dant (49%) whereas the LEPS fraction was the leastabundant: 15% of total EPS (Table 2).

Measurement of total carbohydrates, total proteins,uronic acids and glycosaminoglycan

The quantity of total carbohydrate was highest (605mg g−1) in TEPS and lowest (502 mg g−1) in REPS.

Total protein content was significantly higher(ANOVA-DMRT at P = 0.05) in TEPS and REPS(Table 2). TEPS contain the highest amount of uronicacids and glycosaminoglycan (GAG), 21 mg g−1 and16 mg g−1 of EPS dry weight, respectively (Table 2).

Monosaccharide analysis

Monosaccharide composition analysis revealed the het-erogeneous nature of the EPS of A. platensis MMG-9.Of the eight sugars analysed by ourmethod, sevenweredetected in the EPS (only fructose was not detected inany of the EPS fractions). Five sugars, i.e. fucose,galactose, glucose, rhamnose and xylose, were presentin all three EPS fractions; ribose and mannose wereonly detected in TEPS. Neither ribose nor fructose havebeen reported to be common in cyanobacterial EPS.

Quantitative analysis of the three EPS fractionsrevealed that the different monosaccharides were pre-sent in varying amounts (Table 3). Among the sevenmonosaccharide residues measured in the three EPSfractions, glucose (198 µg (Chl a)−1) and galactose(237 µg (Chl a)−1) were present in significantly higheramounts in TEPS. Rhamnose was the most prominentfraction in REPS and LEPS, comprising 41% and38%, respectively. Mannose and ribose were unde-tectable in REPS and LEPS. Xylose was present inabundance in TEPS (23%).

Confocal laser scanning microscopy

EPS of A. platensis MMG-9 stained well with DTAFand the attached EPS were visible at high resolution

Table 1. Fluorescent dyes and conditions.

Name of dye Buffer used Incubation time Excitation λ Emission λ

DTAF (1 mM) Carbonate buffer, pH 9.0 Overnight 493 nm 517 nmCon A-FITC (1 mM) Phosphate buffer saline, pH 8 20 min 494 nm 518 nmLotus-FITC (1 mM) Phosphate buffer, pH 8 20 min 494 nm 518 nmCSA-TRITC (1 mM) Phosphate buffer, pH 8 20 min 540 nm 605 nmAAA-Alexa 488 Sodium bicarbonate buffer, pH 8.5 20 min 488 nm 518 nm

Table 2. Biochemical characteristics of EPS fractions.

Total EPS(mg g chl a−1) *

Carbohydrates (mg g−1) ofdry EPS*

Protein (mg g−1) of dryEPS*

Uronic acid (mg g−1) of dryEPS*

Glycosaminoglycan (mg g−1) ofdry EPS*

REPS 201.6 ± 0.15(b) 502.1 ± 0.51(a) 16.2 ± 1.26(b) 15.2 ± 1.5(b) 8.2 ± 1.75(b)LEPS 84 ± 0. 13(a) 574 ± 0.22(b) 11.1 ± 2.23(a) 6.3 ± 1.31(a) 5.5 ± 0.89(a)TEPS 275.4 ± 0.14(c) 605.4 ± 0.54(c) 23.3 ± 1.12(c) 21.8 ± 2.81(c) 16.3 ± 1.78(c)

Relative percentages of each component

Total EPS Carbohydrates Protein Uronic acid Glycosaminoglycan

REPS 36 30 32 35 27LEPS 15 34 22 15 18TEPS 49 36 46 50 55

* Mean ± SE (N = 3). Different letters indicate significant differences, Duncan’s multiple range test (P = 0.05).

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along with details of the surface of the trichomes (Figs1–4). CSA-TRITC stained EPS up to 1 µm distantfrom the cells (Figs 5–7). Con-A-FITC stained thinlayers of EPS that seemed to be tightly associated withthe cell and did not extend to the outer layers of theEPS (Fig. 8). Staining with Alexa Fluor 488 showedthe presence of proteins attached to the trichomes butalso in REPS, which is detached from the trichomes(Fig. 9). Lotus-FITC stains fucose in EPS and itseemed to stain the REPS in particular (Figs 10–11).A dense region consisting of a thick layer of EPS (upto 20 µm) was observed around the Arthrospira tri-chomes showing the presence of fucose-bindingdomains in this EPS (Fig. 12).

Staining with individual lectins resulted in thevisualization of EPS, but in order to obtain a detailedinsight into the EPS matrix around the cyanobacterialtrichomes, three lectins with different dyes were usedsimultaneously (Lotus-FITC, CSA-TRITC and AlexaFluor 488). Visualization after combined stainingresulted in the detection of a cloud of EPS aroundthe spiral trichome (Fig. 13). Analysis of 3D stacksshowed the trichome spiral width to be 15–20 µmwhereas the length of the trichome was 100 µm. EPScovering the trichomes was 10–20 µm in thicknessshowing EPS granules of various sizes that are moredensely stained (Fig. 14); 1–2 µm thickness of TEPSwas visible around the trichome.

Discussion

Cyanobacteria produce diverse exopolysaccharides interms of their physical and chemical properties.Because of the large amounts of EPS produced bycyanobacteria and their potential industrial applica-tions, cyanobacterial EPS have been studied exten-sively (Li et al., 2001). In the present study, EPSfrom A. platensis strain MMG-9 were the subject ofinvestigation. They were extracted as three operation-ally defined fractions, of which REPS was the mostabundant. Arthrospira platensis strain MMG-9 pro-duced copious amounts of EPS composed of carbohy-drates and protein in equal proportions in TEPS and

LEPS. The latter fraction is composed of mucilage andits presence indicated that A. platensis strain MMG-9continuously releases EPS. The proportions ofreleased and bound EPS in various studies show ahigh variability (Gloaguen et al., 1995; Nicolauset al., 1999); such variability may be strain dependentor the result of the cultivation conditions. Virtuallyany environmental condition could influence the pro-duction of the different fractions of EPS, which makesit difficult, if not impossible, to compare our resultswith other published data.

Protein is an integral component of cyanobacterialEPS that determines in part the physicochemical prop-erties of these polymers (Kawaguchi & Decho, 2000).Uronic acids enhance the stickiness of cyanobacterialEPS which aids, for instance, biofilm or microbial matformation (Pereira et al., 2009; Stal, 2012).Glucosaminoglycan (GAG), in combination with uro-nic acids, contributes to the anionic nature of EPS;GAG and uronic acids bind calcium and magnesiumions forming ionic bridges between EPS moleculesand charged minerals in sediments. This property ofEPS is particularly important in biofilms and micro-bial mats as it forms the matrix in which the organismsare embedded, helping them to attach to a surface andstabilize their environment (Kawaguchi & Decho,2000; Gao & Zou, 2001). It is therefore not surprisingthat most (90%) of the cyanobacterial EPS studiedthus far contain uronic acid (Pereira et al., 2009).

Cyanobacterial EPS are complex molecules(Pereira et al., 2009); while many bacteria produceEPS containing less than four sugar moieties, cyano-bacterial EPS may be composed of up to 12 mono-saccharides (Pereira et al., 2009). One speciality ofcyanobacterial EPS is the presence of one to threepentose sugars, which differentiate cyanobacterialEPS from other microbial EPS (Sutherland, 1994).The composition of EPS of A. platensis is complex(Trabelsi et al., 2009). Quantitative analysis of EPSfractions considered in this study, i.e. REPS, LEPSand TEPS, revealed the ratio in which the componentmonosaccharides were present in the three operationalfractions to be different, with seven out of eight tested

Table 3. Comparison of constituent monosaccharidic residues in different EPS fractions of A. platensis MMG-9.

Monomer

REPS LEPS TEPS

* µg mg (Chl a)−1 Relative % * µg mg (Chl a)−1 Relative % * µg mg (Chl a)−1 Relative %

Fructose 0 ± 0.0 (a) 0 0 ± 0.0 (a) 0 0 ± 0.0 (a) 0Fucose 37.9 ± 0.5 (c) 15 22.4 ± 0.6 (b) 9 99.2 ± 1.3 (i) 11Galactose 47.1 ± 0.4 (d) 19 75.3 ± 3.02 (d) 32 237.8 ± 3.8 (m) 26Glucose 42.2 ± 1.4 (c) 17 39.1 ± 2.2 (c) 17 198.4 ± 6.5 (k) 22Mannose 0 ± 0.0 (a) 0 0 ± 0.0 (a) 0 30.7 ± 1.2 (d) 3Rhamnose 102.3 ± 1.0 (e) 41 79.8 ± 1.7 (d) 34 114.7 ± 1.8 (j) 13Ribose 0 ± 0.0 (a) 0 0 ± 0.0 (a) 0 15.2 ± 0.8 (b) 2Xylose 18.3 ± 0.8 (b) 8 19.9 ± 1.3 (b) 8 205.0 ± 8.9 (l) 23

* Mean ± SE (N = 3). Different letters indicate significant differences, Duncan’s multiple range test (P = 0.05).

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monosaccharide residues detected in the TEPS frac-tion. Although a wide range of analytical techniqueshave been used to study the monosaccharide compo-sition of EPS, including TLC, HPLC, GC and GC-MS, these techniques can only detect four–eight resi-dues in a single run (Saravanan & Jayachandran,2008; Jindal et al., 2013; Ohki et al., 2014). Glucoseis the predominant monosaccharide in the majority ofcyanobacterial released polysaccharides (REPS); anexception is the REPS of Anabaena sphaerica whichcontains galactose as the main sugar (Li et al., 2001).The EPS fractions of A. platensis strain MMG-9 wererich in glucose, galactose and rhamnose, particularlyin REPS, confirming the findings of Majdoub et al.

(2009) who reported that rhamnose made up to 49.7%of the EPS of Arthrospira platensis. Rhamnose andfucose are deoxy sugars that make EPS hydrophobicand contribute significantly to the emulsifying proper-ties of these polysaccharides (Pereira et al., 2009). Acombination of the hydrophobic and hydrophilic fea-tures of cyanobacterial EPS, along with adhesion tosubstrates, contributes to water storage, thus confer-ring desiccation resistance (Rossi et al., 2012). Fucoseis not an unusual component in EPS. It occurs in theglycoconjugates of many microorganisms and is animportant component of the cell wall and capsulestructures of Gram-negative and Gram-positive bac-teria (Maki & Renkonen, 2004). The relative

Figs 1–9. Localization of EPS from A. platensis MMG-9 using confocal laser scanning microscopy. Figs 1, 3. Overlaid images ofautofluorescence and stained EPS with DTAF. Figs 2, 4. Spiral trichomes embedded in EPS stained with DTAF. Figs 5–6. Overlaidimages of autofluorescence and stained EPS with CSA-TRITC. Fig. 7. Image of EPS stained with CSA-TRITC. Fig. 8. Staining withCon-A-FITC. Fig. 9. Staining with Alexa Fluor 488. Scale bars = 10 µm (Figs 1, 2), 15 µm (Figs 3, 4), 5 µm (Fig. 5), 10 µm(Figs 6–9). Red = cyanobacterial autofluorescence, green = EPS.

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proportion of fucose in EPS of Nostoc has been asso-ciated with the salt resistance of this cyanobacterium(Yoshimura et al., 2012). Xylose has also beenreported as an important component of EPS (FilaliMouhim et al., 1993; Trabelsi et al., 2009), supportingour finding of the large proportion of this sugar in theTEPS of A. platensis (23%), although previous studiesof TEPS have not reported the rare sugar ribose as wasfound in this study.

Confocal laser scanning microscopy providedinformation with regard to the localization of EPSand supported the view that the different operationalEPS fractions could indeed be related to recognizablestructures (Ruas-Madiedo & de los Reyes-Gavilan,2005). The use of various fluorescently labelled lec-tins and fluorescent stains gave insight into the loca-lization of the EPS (Strathmann et al., 2002) and auto-fluorescence of the cyanobacteria visualized the cells(Ahmed et al., 2011). DTAF stains the EPS of cyano-bacteria and plant cells (Ahmed et al., 2011) andshowed nicely the cross-walls between adjacent

cells, which is a typical feature of Arthrospira tri-chomes (Waterbury, 2006). CSA-TRITC, a lectin spe-cific for N-acetylgalactoamine, depicted the galactoserichness in Arthrospira EPS. Both DTAF and CSA-TRITC stained the TEPS but to variable degrees;CSA-TRITC stained only the outer EPS covering ofthe trichomes whereas DTAF stained particularly theregions between two cells with a strong fluorescence(Figs 1, 4 & 7). Besides carbohydrates, DTAF showsalso affinity for proteins and this may be the reason forthe more intense staining of the EPS (Schumann &Rentsch, 1998). Alexa is a sensitive dye for protein(Huang et al., 2004) and allowed us to detect andlocate protein in the EPS. Con-A has affinity formannose, which is present in TEPS in small amountsbut also binds to the glucose in EPS (Strathmann et al.,2002). Con-A staining confirmed the presence ofmannose in the TEPS as there was no glucose foundin this fraction. Some fucose-specific lectins, includ-ing Lotus tetragonolobus (Lotus), which shows higheraffinity for (α-1,2)-linked fucose residues, have been

Figs 10–14. Localization of EPS from A. platensisMMG-9 using confocal laser scanning microscopy. Figs 10–12. Spiral filamentsembedded in EPS stained with Lotus-FITC. Fig. 13. EPS clouds from A. platensisMMG-9 using confocal laser scanning microscopyand three dyes (Lotus-FITC, CSA-TRITC and Alexa Fluor 488). Fig. 14. z-stacks showing the depth of the image (numbers showdepth in µm). Scale bars = 15 µm (Fig. 10), 10 µm (Fig. 11), 50 µm (Fig. 12), 10 µm (Fig. 13). Red = cyanobacterial autofluorescence,green = EPS.

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reported to be less suitable for detection of the sheathor TEPS (Zippel & Neu, 2011). Lotus-FITC stainsfucose in EPS and it seemed to stain particularly theREPS.

Regardless of the complications involved with theuse of multiple fluorochromes, multifluorochromestaining is a valuable technique that allows the detailedinvestigation of microbial samples and is becomingincreasing popular for the study of EPS-producingmicrobes (Chen et al., 2007). Visualization after com-bined staining resulted in the detection of a thick EPSmatrix surrounding the spiral trichome (Fig. 13). Theinner core of the spiral trichome seemed to be staineddensely suggesting a higher amount of EPS in thatregion.

EPS that surround microbes are involved in theattachment of cells to a substrate and also in the for-mation of a matrix in which microbes are embedded(de los Rios et al., 2004). In terrestrial cyanobacteria, athick EPS envelope also serves as a reservoir to holdwater in order to avoid desiccation (Yoshimura et al.,2012). This matrix of mucilage could enhance thebiotechnological importance of Arthrospira as it effi-ciently traps metal ions and other toxic compounds(Garcia-Meza et al., 2005; De Philippis et al., 2011).The ability of cyanobacterial EPS to chelate metalions enables cells to accumulate these trace nutrientsfor growth and/or to prevent them from direct contactwith toxic metals (Pereira et al., 2009).

The use of lectins is an elegant method to visualizedifferent types of microbial EPS (Zippel & Neu, 2011).Furthermore, the use of optical sectioning with confo-cal laser scanning microscopy enables 3D imaging ofundamaged cyanobacterial trichomes along with theirsurrounding EPS (Barranguet et al., 2004). The thickheterogeneous layer of EPS covering the cyanobacter-ial trichomes illustrates how these polymeric sub-stances could act as a physical barrier preventingdirect contact between the cell and its environment.

Acknowledgements

This research was part of the PhD thesis of Dr MehboobAhmed. The Higher Education Commission of Pakistanis acknowledged for providing funding to MehboobAhmed (IRSIP No.1-8 ⁄HEC⁄HRD⁄ 2007⁄923) to visitthe Netherlands Institute of Ecology (NIOO-KNAW)and use Confocal Laser Scanning Microscopy and ana-lyses of EPS.

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