III. ISOLATION, IDENTIFICATION, AXENIC CULTURE AND BIOMASS …shodhganga.inflibnet.ac.in/bitstream/10603/106059/12/12_chapter3.… · Isolation, Identification, Axenic culture and

III. ISOLATION, IDENTIFICATION, AXENIC

CULTURE AND BIOMASS

PRODUCTION

Chapter III………………Isolation, Identification, Axenic culture and Biomass production

31

3.1. INTRODUCTION

In recent past, a number of cyanobacteria have been screened and recognized as a

rich but not yet extensively examined source of food, feed and pharmacological as well as

structurally interesting secondary metabolites (Ugwu et al. 2008). Cyanobacteria

characterize by their blue green color and can be distinguish from other organism. Further

classification of them is based on microscopic and molecular characterization. These

organisms are susceptible and their growth is affected by sudden physical and chemical

fluctuation of environmental conditions such as light, salinity, temperature and nutrient

limitations (Tomaselli and Giovannetti 1993; Oliveira et al. 1999). In literature it is

mentioned, cyanobacteria can grow where moisture is available but in actual experience

the cyanobacterial behavior in nature and in laboratory culture is differ (Chlipala et al.

2011). There is a need to observe the behavior in nature and growth requirements for

individual cyanobacterial species and to apply it in laboratory to scale up the biomass

production.

In the early 1950’s, the increase in the world’s population and predictions of an

insufficient protein supply led to a search for new alternative and unconventional protein

sources. Algal biomass appeared to be a good candidate for this purpose (Becker 2004;

Cornet 1998; Spolaore 2006). Commercial large-scale culture of algae started in the early

1960’s especially using the algal genera Chlorella, Scenedesmus and Dunaniella (Iwamoto

2004; Borowitzka 1999). The Spirulina has been traditionally consume by tribal people in

Central Africa and Mexico therefore in 1970’s, for cyanobacteria Spirulina, the culturing

and harvesting facility established at Lake Texcoco by Sosa Texcoco S.A. (Borowitzka

1999; Muller-Feuga 1996). Among the green and blue green algal species, Spirulina have

received greater attention as a source of human food and poultry feed. Now most species

of Spirulina are mass cultivated globally. The growth of Spirulina is faster and become

possible to obtain the large scale biomass in open and closed culture systems.

However, till date little industrial and economic success is achieved in utilization

of other cyanobacteria. Except Spirulina meager information is available on biomass

production of cyanobacteria. There are evidence that almost a billion year earlier

cyanobacterial forms present in stromalites (Charpy et al. 2012) and they are the pioneer

oxygen producing phtotosynthetic organisms. Additionally, new methods need to be

developed to allow the cultivation of previously ‘uncultivable’ strains.

Axenic cultures are important in genetic, biochemical, physiological and

taxonomic studies. Although numerous methods such as single cell isolation,


32

centrifugation cleaning (Vaara et al. 1979; Bolch and Blackburm 1996), UV irradiation or

gamma irradiation (Kraus MP 1966), differential filtration (Meffert and Chang 1978) and

bactericidal chemical treatment, lysozyme treatment, antibiotic (Pinter and Provasoli,

1958; Rippka R 1988; Vaara et al. 1979; Carmichael et al. 1974; Anderson 2005) have

been suggested to produce the axenic culture of cyanobacteria. Since cyanobacteria are

grows in moist and watery conditions, exhibiting enormous variation in growth,

morphology, possess complex multilayer envelop, filaments are tightly aggregated and

metabolic capabilities, application of any particular approach cannot guarantee success of

their purification. Cyanobacteria and bacteria have similar prokaryotic cell organization

showed the more or less similar responses to selective procedure such as antibiotic

treatment.

Therefore in this chapter, the attempts were made for isolation, identification and

establishment of culture and purification of culture. Further the emphasis was given for

large scale biomass production of some cyanobacteria.

3.2. MATERIALS AND METHODS

3.2.1 Source of cyanobacteria

A total of 840 soil and water samples were collected randomly from paddy field, water

bodies and moist rock from different locations belonging to Pune and adjoining area of

Ahmednagar, and Satara district of Maharashtra state, India during 2009 and 2010 (June –

October) (Table 3.2; Fig. 3.1). The Ahmednagar, Pune and Satara districts are situated at

19° 8′ N 74° 48′ E, 18° 31′ N, 73° 51′ E, 17° 42′ N 74° 02′ E, respectively on a plateau

which is about 43187.89 km2 in area.

Fig. III. 1 District location of study area under Maharashtra (India).


33

The collection of sample and isolation of cyanobacterial strains was carried out by

following the method described by Rippka et al. (1979); Anand et al (1990) and protocol

described by National Facility for Blue Green algae, IARI, New Delhi (Singh et al.

2001).The visualize blue-green filamentous forms were collected from water bodies and

moist soil and rock surface.

The samples were collected in the sterilized culture bottle (0.3 L capacity) and

brought to the laboratory. Some portions of algal samples were preserved in 4% formalin

for microscopic study.

3.2.2. Isolation and establishment of cultures

The soil sample (1g), scrape from moist rock (1g), few filaments from water samples and

Nostoc ball were suspended separately in 100 ml sterile BG-11 medium. The suspension

was shake thoroughly and 10ml suspension was diluted serially using sterile BG-11

medium as: 100, 10

-1, 10

-2, 10

-3 up to 10

-9. Each dilution was separately streaked on BG-11

agar plate. The filamentous forms exhibited radial spreading growth on the agar surface.

The filaments at the peripheral portions were picked up with the nichrome loop and

washed in the sterilized distilled water. Then the filaments were inoculated in the liquid

medium. This procedure was repeated till the unialgal cultures were established. At

weekly interval the cultures were observed under the research microscope for

contamination if any. Then the unialgal cultures were transferred to the culture bottles

with 50 mL of the liquid culture media. The cultures were incubated in culture room at

25±2oC temperature and 50-60% humidity. The cultures were exposed to 8 h light and 16

h dark cycle (30-40µmolm-2

S-1

light intensity).

3.2.3. Identification and enumeration of the cyanobacterial isolates:

The algal biomass was analyzed for chlorophyll and phycobilliprotein pigments using

paper chromatography (Jeffery 1961). After confirmation of cyanobacterial nature on the

basis of pigments the forms were observed for microscopic character. The dimension of

cell and filaments were measured using ocular and stage micrometer under the

microscope. The morphological characters of the cyanobacterial isolates were compared

with the characters enlisted in the monographs, research paper and keys published by

Prescott (1951), Desikachary (1959), Anand (1990, 1993), Kumar (1999), Graham and

Wilcox (2000), Trivedi (2001), Hoek et al. (2002), Anderson (2005), Komarek (2006),

Adhikary (2006), Lee (2008), Amsler (2008), Markou and Georgakakis (2011), Komarek

and Mares (2012). For some of the forms the recorded observations were verified and


34

confirmed with the help of experts from Krishnamurthy Institute of Algology, Chennai.

Identification of some of the cyanobacterial isolates was confirmed with 16s rRNA (Nubel

1997; Shih et al. 2013). The identified isolates were classified and arranged as per the

system of classification described by Desikachary (1959), Komarek and Mares (2012).

The relative abundance and percent of cyanobacterial species were determined

following the method of Devi et al. (1999). The relative abundance was calculated using

the formula: Relative abundance = (Y/X) × 100, where X is the total number of samples

collected and Y is number of samples from which a cyanobacterial strain was isolated.

The percentage of cyanobacterial strains in the soil samples was calculated as: The

percentage of cyanobacterial species (z) = (p/q), where q is the total number of

cyanobacterial species and p is number of particular species of cyanobacteria.

3.2.4. Nutrient media

The recommended nutrient media namely BG-11 (Stanier et al. 1971), Fogg’s medium

(Fogg’s 1949; Jacobson 1951), Allen and Arnon’s medium (Allen and Arnon 1955),

Zarrouk’s medium (Zarrouk 1966), CFTRI medium (Venkataraman and Becker 1984)

were used for establishment and growth studies of cyanobacterial isolates (Table 3.1).

3.2.4.1. Glasswares and plastic wares

The experiments were carried out using borosil make glasswares. The cultures were

grown in autoclavable glass culture bottles with polypropylene caps (0.3L, 5L, 10L, 20L).

For cleaning, the glasswares were soaked overnight in liquid soap and washed in running

tap water thoroughly and placed in an hot air oven at 80oC for 2 hours for drying. The

plastic semi-transparent culture trays (22.5 x 15 x 5.5 cm) were used for growing the

cyanobacterial isolates. After cleaning, the trays were sterilized by soaking in 4 %

formalin overnight. After drying the trays were rinsed with sterilized distilled water.

3.2.4.2. Preparation of stock solutions and nutrient media

The nutrient salts and other chemicals used for preparation of different media were

obtained preferably from HiMedia, Qualigens, SRL, E-Merck and Sigma Company. Glass

distilled water was used for the preparation of stock solutions of nutrient constituents.

Milli Q water (Millipore) was used for the preparation of the nutrient media. The nutrient

constituents and the concentrations recommended for different media are presented in

Table 3.1. The desired quantity of nutrient constituents for 10X stocks were weighed


35

accurately on Contech make single pan digital balance and dissolved in 75ml of distilled

water individually. The final volume (100 mL) was made by adding distilled water in

volumetric flask. The stock solution of the micronutrient was made by adding 100 X

concentration of each nutrient separately in 10 ml of water and finally all the dissolved

nutrients were mixed to make final volume of 100 mL with distilled water. From this stock

solution the desired quantities were picked up for preparation of particular volume of

medium. All stock solutions were stored at 0-4oC in refrigerator and used within a period

of month.

3.2.4.3. BG-11 (Stanier et al. 1971)

The required stock solutions of nutrient components for BG-11 medium were

added so as to maintain the recommended concentration of nutrient constituents as

described in Table 3.1. The stock solutions of were added in distilled water (volume less

by 10 mL than that of final volume). The pH of the medium was adjusted to 7.1 with 0.1 N

NaOH or 0.1 N HCl and final volume made with distilled water.

3.2.4.4. Fogg’s Medium (Fogg 1949; Jacobson 1951)

Required quantity of macro nutrients and micro nutrient stock solutions were

added in distilled water (volume less by 10 mL than that of final volume) as given in Table

3.1. The stock of Fe-EDTA was prepared separately; 26.1 gm of EDTA in 268 ml of 1 N

KOH was taken and 24.9 gm FeSO4. 7H2O was added to it and contents were dissolved by

stirring. The solution was aerated for 16-18 hours and Fe-EDTA was stored in amber

colored reagent bottle (Jacobson 1951). The pH of the medium was adjusted to 7.5 by drop

wise addition of 0.1 N NaOH or 0.1 N HCl.

3.3.4.5. Allen and Arnon medium (Allen and Arnon 1955)

Allen and Arnon medium was prepared as described by Allen and Arnon (1955).

The required stock solutions of nutrient components of Allen and Arnon medium were

added as described in Table 3.1. The stock solutions of different minerals were added in

distilled water (volume less by 10 mL than that of final volume). The pH of the medium

was adjusted to 8.2 with 0.1 N NaOH or 0.1 N HCl.

3.2.4.6. Zarrouk’s medium (Zarrouk 1966)

The required stock solutions of nutrient components of Zarrouk’s medium were

added as described in Table 3.1. The stock solutions of different minerals were added in


36

Table 3.1: Composition of nutrient media.

Constituent BG-11

medium

(g L-1

)

Fogg’s

Medium

(g L-1

)

Allen and

Arnon

medium

(g L-1

)

Zarrouk's

Medium

(g L-1

)

CFTRI

(g L-1

)

NaNO3 1.5 - - 2.5 1.5

K2HPO4.3H2O 0.04 0.20 0.456 1.0 0.5

MgSO4.7H2O 0.075 0.20 0.246 0.02 2.0

CaCl2.2H2O 0.036 0.10 0.074 0.04 0.04

Citric acid 0.006 - - - -

Ferric ammonium

citrate

0.006 - - 0.01 -

EDTA 0.001 - 0.004 0.01 -

Na2CO3 0.002 - - - -

KNO3 - - 2.02 - -

NaCl - - 0.232 1.0 1.0

K2SO4 - - - 1.0 1.0

NaHCO3 - - - - 4.5

FeSO4 - - - - 0.01

Micro nutrients mg/L mg/L mg/L mg/L mg/L

H3BO3 2.86 2.86 2.86 2.86 2.86

MnCl2.4H2O 1.81 1.81 1.81 1.81 1.81

ZnSO4.7H2O 0.222 0.222 0.222 0.222 0.222

Na2MoO4.2H2O 0.390 - - - 0.390

CuSO4.5H2O 0.080 0.079 0.079 0.079 0.080

CO(NO3)2.6H2O 0.040 - - - 0.040

MoO3 - 0.0177 0.0177 0.0177 -

pH 7.1 7.5 7.8 9.2 10.0


37

distilled water (volume less by 10 mL than that of final volume). The pH of the medium

was adjusted to 9.2 with 0.1 N NaOH or 0.1 N HCl.

3.2.4.7. CFTRI medium (Venkataraman and Becker 1984)

The required stock solutions of nutrient components of CFTRI medium were added

as described in Table 3.1. The stock solutions of different minerals were added in distilled

water (volume less by 10 mL than that of final volume). The pH of the medium was

adjusted to 10.0 with 0.1 N NaOH or 0.1 N HCl.

3.2.5. Preparation of solid medium

The desired quantities of liquid media were prepared and after adjusting recommended pH

2% w/v of agar-agar (Himedia, India) powder was added. The flask was kept in

microwave oven to dissolve agar. After well mixing, pour the medium in culture vessels

and sterilized by autoclaving or otherwise for petri-plate preparation the media was

sterilized in conical flask and poured in sterilized petri-plate in an aseptic condition under

laminar air flow.

3.2.6. Sterilization of media, glasswares and equipments

The clump of 7 or 10 plugged culture tubes containing medium was wrapped by

paper partially (above half portion) to prevent wetting of plugs during autoclaving. The

medium was sterilized at 105 kPa pressure and 121oC for 12 minutes. The required

glasswares and equipments such as scalpels, forceps, scissors, petri dishes, and surgical

blade holders were wrapped in the paper and sterilized in an autoclave. Distilled water was

sterilized in the conical flask. As per the recommendations for larger volumes of media or

water, the sterilization time was increased 12 min for 20ml; 20 min upto 100ml; 25 min up

to 2 lit and 35 min up to 5lit and more (Barsanti and Gualtieri 2006).

3.2.7. Inoculation

All the operations were carried out under laminar airflow fitted with ultra violet

(UV) tube and HEPA filter. The sterilized culture tubes, flasks, petri dishes, forceps and

surgical blades were kept in the laminar air flow and exposed to UV for 20 minutes to

sterilize surfaces of all glassware and instruments prior to use for transfer. 1g inoculum of

fresh biomass of cyanobacterial isolates was used for each culture vessel.


38

3.2.8. Culture conditions

All the cultures were maintained in the culture room at temperature 25±2°C under

8-h light/16-h dark photoperiod with a photosynthetic photon flux density of 40 µmoles-2

S-

1 provided by cool white fluorescent tube lights (Philips, India).

The growth of cyanobacterial isolates was observed in different culture media

namely BG-11 (Stanier et al. 1971); Fogg’s medium (Fogg 1949; Jacobson 1951); Allen

and Arnon’s medium (Allen and Arnon 1955), Zarrouk’s medium (Zarrouk 1966) and

CFTRI medium (Venkataraman and Becker 1984).

3.2.9. Purification of cyanobacterial isolates with antibiotics

By using centrifugation method (1000 x for 2 min), the filaments of unialgal isolates

(Table 3.3), were washed 7 times with sterile BG-11 medium. The filaments were

homogenized in BG-11 media using sterile homogenizer. The density of suspension of

was adjusted to 0.5 OD at 540 nm using sterile BG-11 medium. Inoculum size maintained

as 15-20% (v/v) of medium. For antibiotic treatment, the cyanobacterial suspensions were

initially starved for 24 h in dark at 300C. After starvation, sterile Luria Broth media

(Sambrook and Russell 2001) was added to the suspension and incubated for 24 h in dark

at 300C on rotary shaker at 180rpm. After incubation, the culture was centrifuge at

5000rpm for 1 min and washed 3 times with sterile distilled water. The pellet was

suspended in BG-11 medium containing gentamycin, penicillin, streptomycin, pipericillin,

carbenicillin, ampicillin, kanamycin, cefotaxim, imipenem and cephalosporin (100 µg/ml)

individually and incubated at 300C in dark for 24 h. Then the suspensions were washed

with sterile distilled water for 3 times and suspended in sterile BG-11 medium. These

antibiotics were used along with the cyclohexamide (100µg/ml) to avoid the fungal

contamination.

The above treatments of antibiotics were not effective for establishment of axenic

cultures of Leptolyngbya foveolarum. Therefore by following above method the

suspension of Leptolyngbya foveolarum was treated with lysozyme (0.5 mg mL-1

for 10

min), ppm solutions (Plant preservative mixture) and antibiotics namely Meropenem,

Augmentin and Amikacin. The lysozyme treatment was given by following the method of

Sarchizian and Ardelean (2010).


39

3.2.10. Confirmation of axenic culture

After antibiotic treatment and washing with sterile distilled water, the algal suspensions

were inoculated in culture tube containing 10 ml BG-11 medium and incubated at 22±2 °C

for 7 days with 8h light: 16 h dark cycle. To detect the presence of bacteria in the cultures,

1ml of algal suspensions were inoculated with 4ml of broth media enriched with 0.05%

peptone and 0.01% yeast extract and incubated for 24 h. The contaminants were detected

by observing the turbidity in the medium. The cultures were also streaked on the nutrient

agar plate to observed growth of bacteria. To check the purity of the cyanobacterial

strains, the aliquots were examined critically under the phase contrast microscope and oil

immersion microscopy.

3.2.11. Molecular identification

Among 18 cyanobacterial isolates, the morphological characters of Leptolyngbya sp. did

not match with species recorded in literature therefore the attempts were made to identify

this genus with help of 16 s rRNA. For this identification the 16 s rRNA analysis was

carried out with the help of Genome bio Pvt. Ltd Pune.

3.2.12. Extraction of genomic DNA

The total genomic DNA was isolated from the fresh material by the following methods:

600µl of TNES buffer (10mM Tris, pH 7.5, 400 mM NaCl, 100mM EDTA, 0.6% SDS)

and 35µl of Proteinase-K (20mg/ml) was added to the sample and mixed well by inverting

the tube several times. The sample was incubated overnight at 500C after that 166.7 µl of

6M NaCl was added to it. The sample shakes vigorously for 20 seconds and microfuge at

14,000 rpm for 5 min at room temperature. Equal volume of ethanol was added and gently

mixed by inverting the tube a couple of times. The sample was then centrifuged at 14,000

rpm for 10 min at 40C. DNA pellet was washed with 200-700 µl of 100% ethanol followed

by 70% ethanol. The DNA sample was air dried and the pellet was resuspended in Tris-

EDTA.

3.2.13. Amplification of 16s rDNA primers

The PCR amplification of the bacterial 16S rDNA was performed using two bacterial

universal primers, CYA 106F (5’-CGGACGGGTGAGTAACGCGTGA-3’) and

CYA781R (GACTACTGGGGTATCTAATCCCATT-3’), with temperature cycler. DNA

was amplified in a 100 µL mixture composed of fifty picomole of each primer, 25nmol of


40

each dNTP, 200 µg of bovin serum albumin, 10 µl of 10X PCR buffer (100 mM Tris –

HCl [pH9.0], 15mM MgCl, 500mM KCl, 1% [v/v] Triton X-100, 0.1% [w/v] gelatin), and

10ng of template DNA and sterilized ddH2O. To minimize non specific annealing of the

primers to nontarget DNA, 0.5 U of super Taq DNA polymerase (make) was added to the

reaction mixture after the initial denaturation step (5min 1t 940C), at 80

0C.

After PCR is completed, the PCR products were checked on 2% Agarose by

Agarose Gel Electrophoresis and amplicons size was compared using reference Ladder.

2% agarose gel spiked with Ethidium bromide at a final concentration of 0.5 µg/ml was

prepared using Agarose (LE, Analytical Grade, Promega Corp., Madison, WI 53711 USA)

in 0.5X TBE buffer. 5.0 µl of PCR product was mixed with 1 µl of 6X Gel tracking dye.

5µl of gScale 100bp size standard (geneOmbio technologies, Pune; India) was loaded in

one lane for confirmation of size of the amplicon using reference ladder. The DNA

molecules were resolved at 5V/cm until the tracking dye is 2/3 distance away from the

lane within the gel. Bands were detected under a UV Trans illuminator. Gel images were

recorded using BIO-RAD GelDocXR gel documentation system. The PCR product of size

675 bp was generated through reaction using primers CYA106F/ CYA781R1 and 422 bp

product was generated using primer pair CYA359F/ CYA781R

The thermal cycler profile was as follows: initial denaturation at 94°C for 3 min,

followed by 35 cycles of denaturing at 94°C for 1 min, annealing at 60°C for 1 min, and

extension at 72°C for 1 min, and a final extension at 72°C for 10 min.

3.2.14. Sequence analysis and data processing

After this amplification, products were purified by using a geneO-spin PCR product

Purification kit (geneOmbio technologies, Pune; India) and were directly sequenced using

an ABI PRISM BigDye Terminator V3.1 kit (Applied Biosystems, USA) on an automated

DNA sequencing machine (3130 Genetic Analyzer). DNA sequencing was performed

using primer CYA106F and CYA781R1.

The sequences were analyzed using Sequencing Analysis 5.2 software. BLAST (Basic

local alignment search tool) analysis was performed at BlastN site at NCBI (National

center for biotechnology information) server (http://www.ncbi.nlm.nih.gov/BLAST).

Guiding phylogenetic tree was drawn using cluster algorithm with first five hits in NCBI

nucleotide sequence database. Tree generated using TreeTop – Phylogenetic analysis tool


41

(http://www.genebee.msu.su/services/phtree_reduced.html). The distance matrix was also

generated using TreeTop phylogenetic analysis tool.

3.2.15. Large scale biomass production:

For large scale biomass production, the unialgal cultures were grown in 0.3 L, 5lit,

10 lit and 20 lit capacity bottles containing 150 ml, 2.5, 5.0 and 10 lit medium

respectively. For establishment of tray culture system the culture rack unit was design (2.6

m H X 1.35 m L X 0.9m W) which can accommodate 160 trays (22.5 x 15 x 5.5 cm) in 11

compartments. The rack system was equipped with drip system to add the media as per

requirements. The unit was covered with highly transparent polythene. The racks were

placed in a shade net (50%). The experiments for large scale biomass production were

carried out at ambient conditions (light 80-120 µmol m-2

s-1

, temp 23±90 C. humidity 80-

90%). The inoculum of cyanobacterial isolates was prepared in 300 ml bottles. In each

tray or bottle 1g (approx.) of fresh inoculum was added.

3.2.16. Harvesting and measurement of growth:

The biomass from culture bottles and trays was harvested at the end of fifth week

of culture by filtration through double layer of fine nylon net. The harvested biomass was

washed 3 times with sterile distilled water and then air dried in desiccator for 5 min to

remove the surface water. The resultant biomass was weighed and fresh weight was

recorded. The biomass was dried on application of air blower in shade at room

temperature till constant weight achieved. The growth recorded on the basis of fresh

weight and dry weight.

3.2.17. Experimental design and statistical analysis

The experimetns were carried out in completely randomized design and repeated at

least thrice. Results were represented as analysis of variance (ANOVA) followed by

Duncan’s Multiple Range Test (DMRT) at 5% probability level. Otherwise variability in

data was shown as mean ± standard error.

3.3. RESULTS

3.3.1. Cyanobacterial diversity in study area

In the present investigation, cyanobacterial samples were collected from Western ghat

region especially from the locations of Pune, Ahmednagar and Satara district of


42

Maharashtra state, India (Table 3.2). The Western Ghats better known as Sahyadri range is

a hilly range running parallel to the coast, at an average elevation of 1,200 m, is one of the

hot spots of biodiversity in India. The area chosen for this study is represented by

prominent peaks like Kalsubai (1646 m), old Mahabaleshwar (1326 m), Bhor ghat (1285

m), Purandar fort (1313 m), Rajgad (1103 m). Kalsubai is the highest peak of the Sahaydri

ranges. The samples were randomly collected from 35 different localities, collection site,

latitude; longitude and altitude are shown in Table 3.2.

Total 840 samples were collected from 35 localities, out of which 378 were

identified as cyanobacteria and were described based on their morphology. Eighteen

cyanobacteria taxa belonging to 12 genera were recorded; among them twelve were

heterocystous. These identified cyanobacteria belong to 2 order and 5 families. The

number of species found per family was in order Oscillatoriaceae > Nostocaceae >

Scytonmeataceae > Rivulariaceae > Stigonematacae. Among the identified species, 7

species belong to the family Oscillatoriaceae and 6 to Nostocaceae. However only single

species was recorded from the Rivulariaceae and Stigonemataceae. The most densely

populated genus was Nostoc punctiforme followed by Nostoc ellipsosporum. However, the

genus Leptolyngbya fovelolarum found only in 4 habitats. Total 18 species (Table 3.3)

were identified with the help of morphological characters described in the monographs

and keys as per the references mentioned in material and method section.

Recently, the identification and classification of cyanobacteria is based on the

polyphasic approach (Johansen and Casamatta 2005; Komark and Mares 2012) which is a

combination of molecular and traditional morphological characters. In the present

investigation, the morphological characters of the cyanobacterial isolates were compared

with the characters enlisted in the monographs, research paper and keys published by

Prescott (1951), Desikachary (1959), Anand (1990, 1993), Kumar (1999), Graham and

Wilcox (2000), Trivedi (2001), Hoek et al (2002), Anderson (2005), Komarek (2006),

Adhikary (2006), Lee (2008), Amsler (2008), Markou and Georgakakis (2011); Komarek

and Mares (2012). For some of the forms the recorded observations were verified and

confirmed with the help of experts from Krishnamurthy Institute of Algology, Chennai.

Identification of Leptolyngbya foveolarum and Leptolyngbya sp. of the cyanobacterial

isolates was confirmed with 16s rRNA (Nubel 1997; Shih et al. 2013) The identified

isolates were classified and arranged as per the system of classification described by

Desikachary (1959), Anand (1989), and recommendation of Graham and Wilcox (2000),

Bergys (2002), Komarek 2006, Komarek and Mares (2012).


43

Table 3.2 Collection sites of the cyanobacterial samples from the state of Maharashtra

(India)

District Location Latitude N

Longitude E

Altitude

(meters)

Pune Pune University campus 18o 32' 59.21” 73

o 49' 31.75” 583

Pashan Lake 18o 31' 56.64” 73

o 74' 23.96” 592

NCL ground 18o 32' 22.13” 73

o 84' 28.14” 589

Bhimashankar 19o 04' 15.16” 73

o 32' 09.15” 933

Pirangut Ghat 18o 30' 03.25” 73

o 42' 33.08” 699

Paud 18o 31' 34.92” 73

o 36' 58.29” 584

Paud-Sus Road 18o 33' 42.19” 73

o 40' 22.38” 606

Mulshi 18o 31' 47.98” 73

o 31' 28.71” 577

Hinjewadi 18o 34' 53.97” 73

o 43' 28.57” 572

Katraj Ghat 18o 23' 42.62” 73

o 51' 13.60” 915

Nasarapur 18o 15' 21.29” 73

o 53' 23.79” 663

Shirwal 18o 08' 03.47” 73

o 58' 53.23” 595

Bhor Ghat, Bhor 18o 01' 44.50” 73

o 51' 06.75” 1285

Dive Ghat 18o 24' 55.63” 74

o 00' 22.07” 846

Purandar-Saswad Road 18o 18' 28.50” 73

o 58' 48.70” 861

Purandar Fort, Purandar 18o 16' 36.40” 73

o 58' 18.56” 1312

Paud Mulshi Road 18o 31' 41.46” 73

o 36' 40.47” 568

Mulshi Road 18o 34' 26.61” 73

o 32' 36.75” 590

Mulshi area 18o 31' 52.97” 73

o 33' 03.89” 574

Mulshi Dam 18o 28' 53.70” 73

o 29' 53.00” 633

Shivneri 19o 11' 39.66” 73

o 51' 29.76” 918

Ahmednagar

Arangao 19o 01' 15.96” 74

o 43' 19.23” 655

Bhandardara 19o 32' 45.25” 73

o 47' 05.14” 702

Akole 19o 32' 12.82” 73

o 59' 01.38” 601

Sangamner 19o 35' 10.52” 74

o 19' 06.61” 576

Rahata 19o 42' 03.91” 74

o 29' 06.05” 523

Shrirampur 19o 33' 37.25” 74

o 38' 33.02” 503

Rahuri 19o 22' 55.70” 74

o 38' 54.82” 509

Newase 19o 26' 29.84” 75

o 04' 16.40” 497

Shevgaon 19o 20' 39.20” 75

o 12' 54.59” 491

Pathardi 19o 07' 42.36” 74

o 58' 30.60” 647

Satara

Khandala 18o 02' 57.72” 74

o 00' 54.01” 665

Khambataki Ghat 18o 00' 48.87” 74

o 49' 46.95” 920

Pachgani 17o 55' 55.56” 73

o 51' 04.33” 952

Old Mahabaleshwar 17o 57' 43.42” 73

o 39' 45.19” 1326

Lingmala 17o 55' 21.00” 73

o 42' 17.36” 1272


44

Table 3.3: Cyanobacterial diversity from Pune, Ahmednagar and Satara district

Order, Family, Genus and

Species

No. of samples

in which

particular

cyanobacteria

observed

Relative

abundance

(%)

% distribution

of

Cyanobacterial

species

Nostocales

Oscillatoriaceae

Spirullina platensis (Nordst.)

Geitler 07

0.83 1.85

Oscillatoria chalybea 39 4.64 10.3

Phormidium fragile(Meneghini)

Gomont 19

2.26 5.02

Leptolyngbya sp. 01 0.11 0.26

Leptolyngbya fovelarum 04 0.47 1.05

Lyngbya bipunctata Lemm. 32 3.80 8.46

Microcoleus lacustris (Rabenh.)

Farlow 28

3.33 7.40

Nostocaceae

Nostoc punctiforme Born. Et Flah. 68 8.09 17.9

Nostoc entophytum Born. Et Flah. 32 3.80 8.46

Nostoc ellipsosporum (Desm.)

Rabenh. Ex Born. et Flah. 41

4.88 10.8

Nostoc calcicola Brebsson ex Born.

et Flah. 17

2.02 4.49

Nostoc muscorum Ag. ex. Born. at

Flah. 13

1.54 3.43

Anabaena subcylindrica. 18 2.14 4.76

Scytonemataceae

Scytonema tolypothrichoides

Kützing ex Born. et Flah 14

1.66 3.70

Scytonema mirabile (Dillw.) Born. 08 0.95 2.11

Tolypothrix fragilis (Gardner)

Geitler 11

1.30 2.91

Rivulariaceae

Calothrix javanica de Wilde 23 2.73 6.08

Stigonematales

Stigonemataceae

Westiellopsis prolifica Janet 03 0.35 0.79

Total no of cyanobacterial form 378

Total no. of sample collected 840


45

3.3.1.1. Taxonomic enumeration of the species isolated (Plate-III-1&2)

1. Spirulina platensis (Nordst.) Geitler (Desikachary 1959: 190) (Plate-III-1E) Trichome

spirally coiled, filamentous, Multicellular, unbranched, 6-8 µm broad not attenuated at

the ends, spirals 20-30 µm broad, 3-8 µm long; end cells broadly rounded, Cross wall

of cells not distinct, composed of cylindrical cells.

2. Oscillatoria chalybea (Mertens) Gomont (Desikachary 1959: 218) (Plate-III-1J)

Thallus dark blue green, trichome straight, attenuated at the apex and somewhat bent,

8-13 µ broad, cells broad, quadrangular, without calyptra, trichome single forming flat

or thallus sheath, motile end cells showing typical oscillatory movements unbranched,

hormogones present.

3. Phormidium fragile (Meneghini) Gomont (Desikachary 1959: 253) (Plate-III-1K)

Filaments parallel many forming a sheet like thallus, attached to soil, sheath diffluent,

thallus yellowish blue-green, mucilaginous, lamellated; trichomes more or less

flexuous, entangled, constricted at the cross walls, 1-3 µm broad; cells nearly quadrate,

attenuated at the ends; end cell conical, calyptras absent.

4. Lyngbya bipunctata Lemm. (Desikachary 1959: 290) (Plate-III-1E). Filament free or

entangled forming expanded thallus, trichome solitary, free floating; sheath narrow,

colorless cells, 1-2 µm broad, 4-5 µm long, not constricted at the cross walls, end cell

rounded not attenuated.

5. Microcoleus lacustris (Rabenh.) Farlow (Desikachary 1959: 345) (Plate-III-2D).

Filamentous, many trichomes in bundles inside the firm or gelatinizing sheath,

contorted like rope, thallus blackish blue-green; sheath colorless, slimy, trichome

distinctly constricted at the cross-walls, 4-5 µm broad, 6-12 µm long; end cell

rounded, conical not capitate.

6. Leptolyngbya kmn-1(KC589411) (Plate-III-1D): New species recorded in the

present investigation

Filament in cluster and mats, enveloping in thin, firm and colourless sheaths opened at

the apical end. It is immotile with rounded apical cells, usually constricted at the cross

walls, and very rarely false branching. Cells is pale blue-green or olive-green

cylindrical-shaped, 0.5-3 µm wide, length is longer than wide up to several times,

rarely with prominent granules, without heterocytes and akinetes.

7. Leptolyngbya foveolarum (Montagne ex Gomont) Anagnostidis & Komarek 1988

(Plate-III-1C): filamentous, long, thin, bright blue green trichome, cell content

homogenous or sparsely granulated, cells cylindrical, shorter, rarely longer than wide,

0.9 µm -2.3 µm wide, 1-4 µm long, constricted at ungranulated cross wall, sheath often


46

extend beyond end of thrichome, firm, thin, colorless, diffluent, variously curved,

sometimes straight filaments and in parallel arranged, often tangled together, pseudo-

branching rare.

8. Nostoc punctiforme Born. Et Flah. (Desikachary 1959: 374) (Plate-III-1G). Thallus

without firm outer layer, soft formless, trichome densely coiled hardly seen, trichome

1-3 µm broad, filament densely entangled, sub-globose, blue green, 2 mm in diameter,

scattered, attached, sheath delicate, hyaline, mucilaginous; cells barrel-shaped;

heterocyst 2-5 µm broad.

9. Nostoc entophytum Born. et Flah. (Desikachary 1959: 375) (Plate-III-1H). Thallus

macroscopic, blue-green, small, trichome densely entangled with distinct hyaline

sheath; trichome 2-3 µm broad; cells short, 3.8 µ broad, barrel shaped; heterocyst

broader than vegetative cells.

10. Nostoc ellipsosporum (Desm.) Rabenh. Ex Born. et Flah. (Desikachary 1959: 383)

(Plate-III-1F). Thallus irregularly expanded, gelationous; filaments flexuous, loosely

entangled; cells 3-4 µm wide, 5-8 µm long; heterocysts subspherical, 4-7 µm wide, 4-

10 µm long; akinetes ellipsoidal, wall smooth, colorless. Spore ellipsoidal to

cylindrical, 5.8-7.8 µ X 7.8-11.7 µ.

11. Nostoc calcicola Brebsson ex Born. et Flah. (Desikachary 1959: 384) (Plate-III-1B).

Thallus olive grey, up to 2-3 cm in diameter, mucilaginous, slightly diffluent; filament

loosely entangled; sheath indistinct, only at the periphery of the thallus, colorless;

trichome blue-green, 2 µm broad; cells subspherical, longer than broad; heterocysts

subspherical, 2-4 µm broad. Spores spherical, 4.5-5.8 µ diameter.

12. Nostoc muscorum Ag. ex. Born. et Flah. (Desikachary 1959: 385) (Plate-III-1I).

Colony expanded, 3 cm in diameter, olive green; filaments thickly entangled; sheath

distinct only at the periphery of the colony, yellow brown; trichome 4 µm broad; cells

short barrel shaped; heterocysts subspherical, 6 µm wide; akinete oblong, 3-6 µm

broad, 7-10 µm long, wall smooth, yellow. Trichome 5 µ broad, cells barrel shaped,

shorter or longer than broad.

13. Anabaena subcylindrica Borge 1921 (Plate III-1A): Trichome straight, often

aggregated to form blue green colony, Mucilage thin and usually colorless, cells

subspherical or ellipsoidal, 3-4 µ wide, 5-7 µm long, heterocyst 4.5-5.5 µm wide, 5.5 -

8 µm long. Akinetes cylindrical 5-8 µm wide, 21-30 µm long, wall smooth colorless.

14. Scytonema tolypothrichoides Kutzing ex Born. et Flah (Desikachary 1959: 479)

(Plate-III-2B). Thallus dense, brownish green; filaments 8-10 µm broad, 2-3 mm long,


47

repeatedly false branched, false branches very similar to main filaments; sheath

lamellated; trichome 6-8 µm broad; cells longer, densely granulated; heterocyst varied.

15. Scytonema mirabile (Dillw.) Born. (Desikachary 1959: 483) (Plate-III-2A) Colony

bushy, blackish brown; filaments entangled, 2-12 mm long, mostly false branched;

cells 8-12 µm wide, cylindrical, at the end of trichome disc-shaped; sheath blue green;

heterocyst rounded.

16. Tolypothrix fragilis (Gardner) Geitler (Desikachary 1959: 500) (Plate-III-2F) Colonies

dense, blue-green; filament heteropolar, united, free apical ends, falsely branched,

solitary lateral branches; sheath distinct, colorless; trichome with basal cylindrical

heterocyst; cells cylindrical.

17. Calothrix javanica de Wilde (Desikachary 1959: 525) (Plate-III-2C) Filaments

heteropolar, simple, lateral false branches, 30-40 µm broad; trichome with widened

basal, constricted at the cross walls; sheath present, thick, yellow-brownish colored;

heterocyst basal; cells cylindrical. Spores spherical upto 6.5 µ diameter.

18. Westiellopsis prolifica Janet (Desikachary 1959: 596) (Plate-III-1L) Thallus

filamentous, filament loosely entangled, lateral branches clearly seen, cells of main

filament short, 5.0 -9.1 µ broad, 3.5 -7.5 µ long, barrel shaped, cylindrical, lateral

filament 5.5 – 9.8 µ broad, 4.5 -11.5 µ long, heterocyst intercalary, solitary,

cylindrical, true-branched; filament of two types, primary thicker, creeping, torulous,

two seriate, intensely constricted at cross walls, secondary branches thinner, composed

of rounded cells, two seriate, pseudohormocyst.

3.3.2. Purification of Cyanobacteria

The serial dilution and streak plate method were effective to minimize the bacterial and

fungal contaminants associated with cyanobacterial isolates (Plate III-3). Precautiously the

filaments of isolates were picked up and subculture by maintaining all possible aseptic

conditions but the bacterial and fungal contaminants was not removed completely.

Therefore different antibiotics were used to obtain the axenic culture of 18 cyanobacterial

isolates. The data presented in Table 3.4 showed that streptomycin, penicillin, ampicillin

and pipericillin were ineffective to control the bacteria present in the culture of 18

cyanobacterial isolates. Among the different antibiotics, the treatment of 100 µg/ml of

cefotaxim was found to be effective to inhibit the growth of bacteria in the culture of

Oscillatoria chalybea, Scytonema mirabile, Nostoc calcicola, Nostoc entophytum,

Scytonema tolypothrichoides and Calothrix javanica (Plate III-4).


48

Table 3.4: Effect of antibiotic on growth of bacteria associated with the

cyanobacterial filaments.

Antibiotics 100µg/ml

Cyanobacterial isolates 1 2 3 4 5 6 7 8 9 10

Lyngbya bipunctata +++ +++ +++ ++ +++ - - - - -

Phormidium fragile +++ +++ +++ ++ +++ - - - - -

Spirulina platensis +++ +++ +++ +++ +++ + ++ + - -

Oscillatoria chalybea +++ +++ +++ ++ - + - - - -

Leptolyngbya sp. +++ +++ +++ ++ ++ + ++ + - -

Scytonema mirabile +++ +++ +++ + - + - - ++ - Nostoc muscorum +++ +++ +++ ++ ++ + - + - -

Anabaena

subcylindrica

+++ +++ +++ +++ +++ - - + + -

Nostoc calcicola +++ +++ +++ + - - - ++ + -

Nostoc ellipsosporum +++ +++ +++ + + + - + - -

Nostoc punctiforme +++ +++ +++ ++ + - + + - -

Nostoc entophytum +++ +++ +++ +++ - - - + + - Scytonema

tolypothrichoides

+++ +++ +++ ++ - - - - - -

Tolypothrix fragilis +++ +++ +++ +++ + - - + - -

Westellopsis prolifica +++ +++ +++ + + - - + + -

Leptolyngbya

foveolarum

+++ +++ +++ +++ +++ +++ +++ +++ +++ +++

Calothrix javanica +++ +++ +++ ++ - + - + - -

Microcoleus lacustris +++ +++ +++ +++ + - - + - -

1- Streptomycin , 2- Penicillin 3 – Ampicillin, 4- Pipericillin, 5- Cefotaxim, 6-

Cephalosporin, 7- Gentamicin, 8- Carbenicillin, 9-Kanamycin, 10-Imipenem

The antibiotic cephalosporin, gentamycin, carbenicillin, kanamycin and imipenem

were effective to control the growth of bacteria present in cultures of Lyngbya bipunctata

and Phormidium fragile. The bacteria present in the culture of Spirulina platensis were

sensitive to 100 µg/ml of kanamycin or 100 µg/ml of imipenem.

Gentamicin was effective for inhibiting the growth of bacteria present in culture of

Oscillatoria chalybea, Scytonema mirabile, Nostoc muscorum, Anabaena subcylindrica,

Nostoc calcicola, Nostoc ellipsosporum, Nostoc entophytum, Scytonema tolypothrichoides,

Tolypothrix fragilis, Westellopsis prolifica, Calothrix javanica and Microcoleus lacustris.

But at the same time it resulted in decolorization of the filament.

Among the antibiotics used (Table 3.4) imipenem was found to be effective to

check the growth of bacteria present in the culture of all the cyanobacterial isolates except

Leptolyngbya foveolarum. The treatment of this antibiotic did not affect the growth of

cyanobacterial isolates, the filaments were healthy and dark blue green in color.

The treatments of ampicillin, penicillin, streptomycin, pipericillin, kanamycin,

carbenicillin, cefotaxim, gentamicin, cephalosporin, individually and in combinations

were not effective in controlling the bacterial growth completely in cultures of


49

Leptolyngbya foveolarum. The aliquots staining of Leptolyngbya foveolarum showed the

bacteria firmly attached to the filaments (fig 3.2) Therefore the lysozyme, commercial

plant preservative mixture (ppm) solution, meropenem, amikacin and augmentin were

used to control the bacteria. The lysozyme treatment (500µg/ml) and ppm solution (10-

100µl) were observed to be ineffective for preparation of axenic culture. The bacterial

growth observed immediately after 24h on streaking the treated algal suspension on

nutrient agar. The higher concentration of imipenem and meropenem (7 mg/ml) required

to control the growth of bacteria present in the culture of Leptolyngbya foveolarum. 25

mg/ml of amikacin was effective for controlling the growth of bacteria. At the same time,

very low concentration of augmentin (500 µg/ml) (Table 3.5) was highly effective to

control the bacterial growth and finally to eliminate the bacteria from the cultures of L.

foveolarum.

Fig 3.2: Bacterial association with the filaments of Leptolyngbya foveolarum

Table 3.5 Effect of antibiotic on purification of L. foveolarum

Concentration Imipenem Augmentin Meropenem Amikacin

200µg + + + +

250µg + + + + 500µg + - + +

750 µg + - + +

1mg + - + +

1.5mg + - + +

2mg + - + +

3mg + - + +

4mg + - + +

5mg + - + +

6mg + - + +

6.5mg - - - +

7mg - - - +

10mg - - - +

15mg - - - +

20mg - - - +

25mg - -

+ = bacteria present, - = bacteria absent after 72 h incubation at 300 C


50

3.3.3. Identification of Leptolyngbya foveolarum and Leptolyngbya sp by 16 s rRNA

Phylogenetic relationships of the 16S rDNA sequences from the isolated strains were

compared with other cyanobacterial sequences retrieved from GenBank. Molecular

characterization showed that the isolated cyanobacterium was Leptolyngbya foveolarum

and all result inferred from 16s rRNA sequence analysis were in agreement (Table 3.6).

The isolates morphological feature also suggested that the isolate was L. foveolarum

(Plate-III-1C)

Table 3.6. Result from the blast searches using 16s rRNA of Leptolyngbya foveolarum

and Leptolyngbya sp.

Marker gene Length

(bp)

Closest math

(Gene bank Accession

number)

Overlap % Sequence

similarity %

Taxanomic

affinity

16SrRNA 611 Leptolyngbya

foveolarum VP1-08

(FR798945)ab

100% 100% Leptolyngbya

foveolarum

16SrRNA 562 Leptolyngbya kmn-1.

(KC589411)

95% 95% Leptolyngbya

sp.

a=the closest culture match b=isolated from concrete-made fountain with stagnant water,

gray dry crust, Italy

Fig 3.3: Analysis of PCR products from the cyanobacterial 16s rRNA

gene related to genus-specific sequence signatures

Lane 1: 200-1000bp DNA Molecular Marker; Lane 1: Leptolyngbya sp. 1 PCR with CYA106F/

CYA781R1; Lane 2: Leptolyngbya foveolarum PCR with CYA106F/ CYA781R1; Lane 3: Sample

1 PCR with CYA359F/ CYA781R1; Lane 4: Negative control reaction CYA106F/ CYA781R1;

Lane 5: Negative control reaction CYA359F/ CYA781R1.


51

Fig 3.4 Phylogenetic tree based on 16srRNA Phylogenetic Tree (Guiding tree) Guiding phylogenetic tree drawn using cluster algorithm with first five

hits in NCBI nucleotide sequence database. Tree generated using TreeTop – Phylogenetic analysis tool. (B-

128 Leptolyngbya sp, DQ431002 - Leptolyngbya sp,, EU068733 - Leptolyngbya sp, GU186906 – Calothrix

sp., FJ839352 – Leptolyngbya sp., AY493575-Leptolyngbya frigida.

3.3.3.1. 16S rRNA gene sequence of the Leptolyngbya kmn-1(KC589411)

1 accgctaaga ccccatatgc cggaaggtga aatagttttc tgcctgagga tgagctcgcg

61 tccgattagc tagttggtgg ggtaagagcc taccaaggcg acgatcggta gctggtctg

121 gaggatgatc agccacactg ggactgagac acggcccaga ctcctacggg aggcagcagt

181 ggggaatttt ccgcaatggg cgaaagcctg acggagcaag accgcgtgag ggaagaaggt

241 ctgtggattg taaacctctt ttgattggga agaagcactg acggtaccaa tcgaatcagc

301 ctcggctaac tccgtgccag cagccgcggt aatacggagg aggcaagcgt tatccggaat

361 tattgggcgt aaagcgtccg taggtggttt gtcaagtctt ctgtcaaagc gcggagctta

421 actccgtaaa ggcagaggaa actgacaggc tagagtgcga taggggcaag gggaattccc

481 agtgtagcgg tgaaatgcgt agatattggg aagaacaccg gtggcgaaag cgccttgctg

541 ggtctgcact gacactgagg ga

3.3.4. Culture media and growth of isolates

Total 18 species were identified in the laboratory and were studied for the growth and

biomass production. These 18 species were grown in BG-11, Fogg’s, Allen and Arnon,

Zarrouk's Medium and CFTRI media and the growth was recorded on the basis of dry

weight (Table 3.4). The growth of Leptolyngbya foveolarum, Phormidium fragile,

Tolypothrix fragile, Microcoleus lacustris, Scytonema mirabile, Scytonema

tolypothrichoides, Calothrix javanica and Westellopsis prolifica was higher in BG-11

medium as compared to other media (Table 3.4). The obtained dry weight in the BG-11

medium for these different isolates ranges between 12 to 95 mg per bottle. Among these

the growth of Phormidium fragile was higher (95 mg). The dry weight obtained of these

isolates was comparatively very low in Fogg’s and Allen and Arnon medium. The isolates

Nostoc entophytum, N. ellipsosporum, N. muscorum, N. punctiforme, N. calcicola and


52

Anabaena subcylindrica showed maximum growth in Fogg’s media. The dry weight of

these forms remains in the range of 13 to 34 mg per culture. The maximum 34mg dry

weight/ bottle was observed in Nostoc calcicola. The Spirulina platensis showed

maximum 18 mg dry weight/ bottle in CFTRI medium.

Table 3.7. Growth of cyanobacterial isolates in different nutrient medium.

Sr.

No.

Cyanobacteria Growth in media in terms of dry wt (mg/culture)

BG-11 Allen &

Arnon

Fogg's

Medium

Zarrouk's

Medium CFTRI

1 Phormidium fragile 95±2.4a 35±0.9a 22±1.9b 21±2.3a 18±0.5a

2 Lyngbya bipunctata 84±1.5b 34±1.7a 12±1.7cd 15±1.8bc 11±2.1b

3 Microcoleus lacustris 25±1.9d 13±1.2c 12±0.5cd 09±0.4c 10±2.3b

4 Nostoc punctiforme 12±1.6f 13±0.8c 16±2.3c 04±0.2d 05±0.6d

5 Nostoc entophytum 12±0.9f 07±0.5de 13±2.5cd 10±0.4c 08±0.3c

6 Tolypothrix fragile 12±1.4f 08±0.2d 07±0.4d 10±1.2c 05±0.2d 7 Anabaena subcylindrica 12±0.5f 11±0.4c 14±2.9cd 05±0.5d 10±0.3b

8 Nostoc ellipsosporum 14±0.2f 10±1.2cd 17±1.7c 12±1.4bc 10±0.2b

9 Nostoc calcicola 21±1.8d 13±1.7b 34±1.9a 18±1.7b 12±0.9b

10 Nostoc muscorum 12±2.4f 11±1.7c 16±1.4c 09±0.7c 07±0.4c

11 Leptolyngbya sp. 22±1.2d 06±1.2e 09±0.2d 05±0.2d 02±0.5e

12 Leptolyngbya foveolarum 45±2.8c 12±1.3c 15±1.9c 19±1.7b 02±0.5e

13 Scytonema

tolypothrichoides

29±3.6d 18±1.4b 12±1.6cd 13±1.3bc 09±0.2bc

14 Westellopsis prolifica. 11±4.5f 13±1.9c 12±1.3cd 11±1.8c 10±0.2b

15 Calothrix javanica. 22±1.6d 15±1.8bc 13±1.2cd 19±0.9b 10±0.4b

16 Scytonema mirabile 18±2.9e 12±0.9c 09±0.5d 04±0.05 07±1.8c

17 Spirulina platensis 12±2.3f 09±0.6d 12±0.2cd 13±0.1bc 18±1.2a

18 Oscillatoria chalybea 17±1.5e 09±0.3d 03±0.01e 06±0.7d 02±0.6e

Values are Mean± SE of three independent experiments. Means followed by the same

letters are not significantly different at p=0.05 by Duncan’s Multiple Range Test

3.3.5. Biomass production

In pilot experiments the BG-11 medium supported the maximum growth of Leptolyngbya

foveolarum, Phormidium fragile, Tolypothrix fragile, Microcoleus lacustris, Scytonema

mirabile, Scytonema tolypothrichoides, Calothrix javanica and Westellopsis prolifica.

While Fogg’s media supported the growth of Nostoc entophytum, N. ellipsosporum, N.

muscorum, N. punctiforme, N. calcicola and Anabaena subcylindrica. CFTRI medium

was best for the growth of Spirulina platensis. Therefore for the large scale biomass

production of these isolates in 0.3 L (Plate III-5), 5 L, 10 L and 20 L bottle and

polypropylene trays (22.5 x 15 x 5.5 cm) (Plate III-6) the respective media were used and

data on biomass production is depicted in Table 3.5. The biomass of Spirulina platensis in

0.3 L, 5L, 10L, 20L bottle and tray was 18, 28, 59, 116 and 213 mg respectively (Table

3.8). Similar trend was observed for biomass production in other isolates of cyanobacteria.

For all isolates of cyanobacteria, the maximum growth in terms of dry weight was


53

Table 3.8. Biomass production of cyanobacterial isolates in BG-11 medium in different culture vessel.

Values are Mean± SE of three independent experiments. Means followed by the same letters are not significantly different at p=0.05 by

Duncan’s Multiple Range Test.

Sr.

No.

Cyanobacterial

isolates

Type of culture vessel

Biomass production in terms of dry weight (mg/vessel)

0.3 L 5 Lit 10 lit 20 Lit Tray

1 Phormidium fragile 95±2.4a 190±2.8a 294±4.5a 528±1.6a 743±0.8d

2 Lyngbya bipunctata 84±1.5a 183±1.4b 273±3.7a 481±2.7b 635±0.4e

3 Microcoleus lacustris 25±1.9cd 94±2.4f 141±3.2c 234±1.2g 327±0.5l

4 Nostoc punctiforme 16±2.3ef 75±1.9g 118±2.6e 286±1.6f 394±0.3j

5 Nostoc entophytum 13±2.5fg 36±1.5j 139±2.2de 342±1.4e 649±0.2e

6 Tolypothrix fragile 12±1.4fg 42±2.4i 98±3.8f 181±1.8h 405±0.9h

7 Anabaena subcylindrica 14±2.9ef 53±1.2h 87±4.5fg 194±2.7h 389±0.7i

8 Nostoc ellipsosporum 17±1.7de 93±1.7f 129±1.7e 197±1.2h 412±0.2h

9 Nostoc calcicola 34±1.9c 127±1.4e 219±5.8b 376±1.9d 839±0.1c

10 Nostoc muscorum 16±1.4ef 91±1.2f 143±9.5c 329±2.2e 525±0.2f

11 Leptolyngbya sp. 22±1.2d 40±2.3i 148±3.7c 312±2.3e 729±0.2d

12 Leptolyngbya foveolarum 45±2.8b 136±4.2d 217±4.8b 475±3.2b 982±0.1a

13 Scytonema tolypothrichoides 29±3.6cd 143±1.8c 223±4.3b 435±2.2c 862±0.4b

14 Westellopsis prolifica. 11±4.5fg 35±3.2j 74±3.2g 199±2.5h 373±0.5k

15 Calothrix javanica. 22±1.6d 73±1.9g 155±2.7c 236±3.1g 531±0.8f

16 Scytonema mirabile 18±2.9de 51±2.3h 137±3.5d 219±2.4g 453±0.2g

17 Spirulina platensis 18±2.3de 28±2.7k 59±3.6h 116±3.1j 213±0.1n

18 Oscillatoria chalybea 17±1.5de 27±1.7k 43±3.8i 138±2.2i 298±0.4m


54

Table 3.9. Biomass production of cyanobacterial isolates in BG-11 medium in culture rack unit

Sr.

no.

Cyanobacterial isolates Biomass production in culture rack unit

Fresh

weight (g)

Dry weight (g) Moisture content

%

1 Spirulina platensis 803.5±3.6h 70.8±2.7g 91.2

2 Scytonema tolypothrichoides 1587.6±2.4c 138.9±1.9c 91.3

3 Leptolyngbya foveolarum 1675.2±2.7b 162.7±1.4a 90.3

4 Nostoc calcicola 2032.3±3.4a 149.0±1.5b 92.6

5 Tolypothrix fragilis 896.4±4.5g 65.7±1.3h 92.6

6 Westellopsis prolifica. 678.7±5.6j 59.9±1.9h 91.9

7 Calothrix javanica. 1009.4±2.3e 86.4±2.4f 91.4

8 Phormidium fragile 1446.3±2.5d 112.1±3.2d 92.2

9 Lyngbya bipunctata 995.4±5.2f 98.3±2.9d 90.1

10 Nostoc entophytum 985.2±3.5f 102.5±3.5e 89.5

11 Scytonema mirabile 824.8±4.8h 72.92±2.4g 91.1

12 Microcoleus lacustris 557.8±0.8k 45.8±1.2i 91.7

13 Nostoc punctiforme 732.2±1.4i 59.6±2.4h 93.7

14 Anabaena subcylindrica 787.6±1.7h 57.4±2.9h 92.7

15 Nostoc ellipsosporum 895.7±1.8g 53.6±2.4h 94.0

16 Oscillatoria chalybea 865.3±2.4h 51.2±1.9h 94.0

17 Nostoc muscorum 978.5±2.6f 49.8±1.7hi 94.9

18 Leptolyngbya sp. 1088.6±2.8e 42.2±1.5i 96.1 Values are Mean± SE of three independent experiments. Means followed by the same letters are not

significantly different at p=0.05 by Duncan’s Multiple Range Test.


55

observed in trays with one liter medium compare to bottle cultures. Therefore the biomass

production was studied in trays.

Considering these factors the culture rack unit was designed (size) which can

accommodate 160 trays in 11 compartments. The result of the present investigation

suggested that cyanobacterial biomass production is possible by using the culture rack

covered with polythene sheet and application of 50% shed net at ambient conditions where

diffuse sunlight is available (light 80-120 µmol m-2

s-1

, temp 23±90 C humidity 80-90%).

The use of transparent plastic trays increase the area for the growth and therefore the

developed method was found to be suitable as compare to other containers for maximum

biomass production in static conditions. The arrangement of drip system was convenient

for timely addition of nutrient solution and water in each culture tray (Plate III-7). This

helps to reduce the time and labor cost for filling the tray with nutrient solutions. In tray-

rack system, the visible growth was observed within 5 to 7 days after initiation of culture

which was evident by the formation of thin film of biomass at the surface of the medium.

Maximum growth in terms of dry weight was observed at the end of 5th

week of culture.

The growth was not increased in the sixth week of culture and the filaments start turning

to yellowish-whitish in color. The maximum average dry weight of Leptolyngbya

foveolarum in tray-rack unit was 162.7±1.4 g less to it was of Nostoc calcicola >

Scytonema tolypothrichoides> Phormidium fragile> Nostoc entophytum> Lyngbya

bipunctata > Calothrix javanica> Spirulina platensis>Scytonema mirabile> Tolypothrix

fragilis> Westellopsis prolifica (Table 3.9, Plate III-8,9).

3.4. Discussion

3.4.1. Cyanobacterial diversity in study area

Cyanobacteria are prokaryotes and lacks sexual fusion of gametes. Therefore taxonomist

historically used morphological features to define cyanobacterial taxa including variation

in cyanobacterial thallus structure i.e. occurrence as unicells, colonies, unbranched

filaments or branched filaments, false or true branches, presence or absence of exospores,

endospores or akinets, heterocyst, mucilaginous sheath, trichome, harmogonia, separation

disc (Graham and Wilcox 2000). However, for the modern taxonomic classification of

cyanobacteria, molecular sequence comparisons together with morphological features,

ecophysiological characters and other biochemical and molecular marker is essential

(Komarek and Mares 2012).


56

In the present investigation, total 840 soils and water samples were analyzed and

existence of cyanobacteria was observed in 378 samples. On the basis of morphological

and molecular characters 18 cyanobacterial species were identified belonging to 12

genera, 5 families and 2 orders (Table 3.2, 3.3). Order Nostocales was dominant and at the

family level, Nostocaceae and Oscillatoriaceae showed a higher percent frequency of

distribution. The results on relative abundance of genera depicted in Table 3.3 shows that

Nostoc punctiforme Born. Et Flah (8.09 %) was dominant. Other prominent and common

cyanobacteria were Nostoc ellipsosporum (Desm.) Rabenh. Ex Born. et Flah. (4.88 %),

Nostoc entophytum Born. Et Flah and Lyngbya bipunctata Lemm (3.8 %). Leptolyngbya

foveolarum was observed in seven soil samples. However, Leptolyngbya sp. showed least

relative abundance (Table 3.3). Previous studies on the cyanobacterial diversity from other

Indian states revealed that Nostoc sp. was dominant in Asam, Hariyana, Kerala,

Tamilnadu, West Bengal (Venkatramn 1975), Orisa (Sahu et al. 1996) and Manipur (Devi

et al. 1999). A limited study from the arable domain of the Pravara area of Ahmednagar

district Maharashtra revealed that, in addition to Nostoc, the most common other genera

were Oscillatoria, Westellopsis, Scytonema, Microchaete, Anabaena and Tolypothrix were

the most common genera (Auti and Pingle 2010). The genus Westellopsis showed

restricted distribution in the rice field soils of Manipur (Devi et al. 1999). The genera

Scytonema and Oscillatoria were represented by maximum number of species in paddy

fields of Western Maharashtra (Patil and Chaugule 2009). Scytonema was most abundant

in salt-affected soils of Kolhapur district (Madane and Shinde 1993). In the area of the

present investigation the relative abundance of Scytonema was only 1-2%, which is very

low compared to Madane and Shinde’s (1993) study which reported that Scytonema was

most abundant in saline soils of Kolhapur district, Maharashtra state. This difference could

be because our study was not specific to saline soils. Cyanobacteria are the pioneer

oxygenic phototrophs on the earth whose distribution around the world is surpassed only

by bacteria (Adams 2000). Until past few decades of research cyanobacterial were of

academic interest and were mostly ignored as nuisance but now are proved as potential

organism for food and utilizable molecules (Thajuddin and Subramanian 2005). Literature

suggest that cyanobacteria comprising more than 150 genera and 2000 species. On

verification and exploitation of rest of the area the number may increase. (Gupta et al

2013).

In this context, identifying and cataloguing the diversity of cyanobacterial species

from different regions including the Sahyadri ranges, a biodiversity hot spot of India in


57

Western Ghats, will be very beneficial in exploiting them for source for producing

metabolites with novel biological activity.

3.4.2. Purification of cyanobacteria

Purification of cyanobacteria from bacteria is tiresome and time demanding process (Singh

et al. 2001). In the present investigation also difficulty encountered in the isolation of

axenic cultures of cyanobacterial isolates. In the present investigation series of attempts

were made including serial dilution, sonication, repeated streaking, centrifugation cleaning

and various antibiotic treatments. However, the methods of centrifugation cleaning and

treatment of some antibiotics were effective in purification of cyanobacterial isolates.

Except Leptolyngbya foveolarum, the treatment of 100 µg/ml of imipenem was best for

elimination of bacteria and purification of 17 isolates of cyanobacteria (Lyngbya

bipunctata, Phormidium fragile, Spirulina platensis, Oscillatoria chalybea, Leptolyngbya

sp., Scytonema mirabile, Nostoc muscorum Anabaena subcylindrica, Nostoc calcicola,

Nostoc ellipsosporum, Nostoc punctiforme, Nostoc entophytum, Scytonema

tolypothrichoides, Tolypothrix fragilis, Westellopsis prolifica, Calothrix javanica and

Microcoleus lacustris). Antibiotics are widely used to remove bacteria associated with

cyanobacteria (Han et al. 2010). In this study, thirteen broad-spectrum antibiotics, namely,

imipenem, meropenem, augmentin, penicillin, streptomycin, pipericillin, carbenicillin,

ampicillin, gentamycin, kanamycin, amikacin, cephalosporin, cefotaxim were used. The

variable results were obtained to control the growth of bacteria and purification of the

cyanobacterial isolates. The results indicate that Imipenem was good for 17 cyanobacterial

isolates. Among the used antibiotics some antibiotics were not effective and some others

were effective for elimination of bacteria from the cultures (Table 3.4). One of the reason

is the cyanobacterial isolates were filamentous and most of them possess mucilaginous

sheath which helps to adhere the heterotrophic bacteria. Earlier reports in Phormidium

animalis, even selective antibiotic treatment have proven to be ineffective, as

mucilaginous coats surrounding the algal filaments harbor and protect bacteria from

antibiotics and long filaments make it practically impossible to eliminate all the bacteria

associated along the whole extension of the filament (Varquez-Martinez et al. 2004).

Axenic cultures of Arthospira platensis SAG 21.99 were obtained with the

combination of a washing step and a consecutive treatment with antibiotics imipenem,

neomycin, and cyclohexamide (Choi et al. 2008). The treatment of 1 g/L of lysozyme for

establishment of axenic culture of Anabena flos-aquae and Aphanothece nidulans (Kim et

al. 1999). Bolch and Blackburn (1996) were succeeded in purifying 12 of the 17 strains of


58

Microcystis aeruginosa on application of centrifugation cleaning, sulfide gradient

selection and antibiotic imipenem treatment. Axenic cultures of Phormidium anamalis

was obtained by combination of washing in a series of centrifugation and resuspension in

liquid medium followed by vacuum driven filtration through 8 µm pore size membrane

and treatment with carbenicillin followed by streptomycin then chloramphenicol, and

finally with kanamycin (Varquez-Martinez et al. 2004). Hong et al. (2010) succeeded in

purifying the filament of Nodularia spumigena KNUA005 after three purification step;

centrifugation, antibiotic treatment and streaking. He used antibiotic imipenem, the

bacteria still survived after treatment of imipenem which was further eliminated by

kanamycin.

Ppm is commercially available formulation which is used to avoid contamination

in plant tissue culture. In the present study the ppm was not effective to control the growth

of bacteria. Earlier report suggests that imipenem is broad spectrum beta lactum antibiotics

which kill the most of the bacteria (Pinter and Provasoli 1958; Rippka 1988; Bolch and

Blacburn 1996; Ferris and Hirsch 1991). However, in the present investigation for L.

foveolarum the antibiotic imipenem was ineffective up to 7mg/ml to eliminate the bacteria

from the culture. Similar results were also observed in Microcystis aeruginosa (Han et al.

2010).

Augmentin was the most effective antibiotic of this pool to eliminate the bacteria

from the cultures of L. foveolarum. This might be possible being augmentin a broad-

spectrum beta-lactam antibiotic which inhibits peptidoglycan biosynthesis and making it

superior to the other antibiotics used i.e. imipenem, meropenem, amikacin, gentamicin for

reducing the number of heterotrophic bacterial contaminants associated with L.

foveolarum culture.

3.4.3. Identification of the cyanobacterium

Literature suggest that cyanobacteria comprising more than 150 genera and 2000

species (Gupta et al. 2013). Members of the cyanobacteria are generally classified into five

orders: Chroococcales, Nostocales, Oscillatoriales, Pleurocapsales, and Stigonematales

(Anagnostidis and Komarek, 1988). Members of the order Oscillatoriales are filamentous

and lack heterocysts and akinetes (Albertano and Kovacik 1994; Turner 1997). The order

Oscillatoriales includes the families Borziacaea, Homoeotrichaceae, Oscillatoriaceae,

Phormidiaceae, Pseudanabaenaceae, and Schizothrichaceae as discussed in Anagnostidis

and Komarek (1988).


59

Many of the species within the genera Oscillatoria, Lyngbya, Phormidium,

Schizothrix, and Plectonema were originally classified by Gomont (1892) based on sheath

characteristics and the presence or absence of false branching. However, the sheath

characteristics frequently used in identifying and classifying members of this group are

influenced by culturing and environmental conditions (Albertano and Kovacik 1994). This

led to the eventual transfer of a number of strains within the Oscillatoriales to a new group

designated by Rippka et al. (1979) as LPP-group B (Lyngbya, Phormidium, and

Plectonema).

The identification of microorganisms including cyanobacteria in the natural

ecosystem is difficult. Traditional taxonomic criteria developed over a century ago were

based on erratic characters such as false branching and sheath characteristics. The

morphology of cyanobacteria is severely influenced by environmental factors which

induced phenotypic plasticity was not observed or well understood by early taxonomists.

The problem in using botanical criteria to classify cyanobacteria is that culturing

conditions and environmental plasticity often induce morphological changes (Burja et al.

2001). The most plastic characters observed in cyanobacteria include sheath color, sheath

thickness, granulation, false branching, and cell length (Anagnostidis and Komarek 1985).

Less variable characters include pigmentation, cell length to width ratios, tapering,

trichome width, and the ability to form calyptra (Anagnostidis and Komarek 1985).

Furthermore, morphological characters such as type of cell division and thylakoid

structure are constant, and are not influenced by variations in environmental or culturing

conditions (Anagnostidis and Komarek 1985). Culture and environmentally induced

morphological changes among cyanobacteria often lead to inaccurate identification and

classification taxa (Nelissen et al. 1992; Nubel et al. 1997). Morphological changes can be

problematic in establishing species definitions in that they are usually defined based on

cell dimensions and ecology, such that distinct separations are often not evident (Nubel et

al. 1997).

Anagnostidis and Komarek (1988) reassigned many different taxa from the LPP-B

group into a new genus Leptolyngbya, thus creating over 75 new combinations (Albertano

and Kovacik 1994; Turner 1997). This newly established group is typified by the presence

of a number of distinguishing morphological characteristics such as thin sheaths,

immobility of filaments, thin uniseriate trichomes, arrangement of thylakoids, and cell

wall constrictions (Albertano and Kovacik 1994; Turner 1997).


60

In recent years, molecular techniques have provided excellent complementary data

to address the limitations of taxonomic, physiological and biochemical approaches in

biodiversity assessment (Graham and Wilcox 2000; Bursanti and Gualteri 2006;

Srivastava et al. 2007; Lin et al. 2012).

Sequencing of DNA play an important role in the reconstruction of evolutionary

relationships among organism’s leads to new genetic classifications which confirm or

conflict with traditional taxonomy. Molecular techniques used to amplify some portions

of the genome in order to characterize and deduce phylogenetic relationships of

cyanobacteria has increased considerably in the recent years (Neilan et al. 1995; Garcia-

Pichel et al. 1996; Orcutt et al. 2002; Taton et al. 2003; Premanandh et al. 2006). The

rRNA genes are the most widely used markers for the identification of bacteria and

cyanobacteria due to their conserved function and universal presence. Several researchers

have exploited the conserved regions of the 16S rRNA gene for phylogenetic analysis of

cyanobacteria (Nubel et al. 1997; Crosbie et al. 2003; Salomon et al. 2003).

The 16S rRNA is an effective tool in inferring phylogenetic relationships between

different genera within orders proposed by Komarek and Anagnostidis (Turner 1997).

Additionally, it has been useful in identifying and classifying strains that belong to a single

clade (Palinska et al. 1996; Otsuka et al. 1998). Nelissen et al. (1992) found that the 16S

rRNA sequences of five strains of Pseudanabaena were nearly identical, and therefore

concluded that Pseudanabaena was a single monophyletic taxon. In a more recent study,

Honda et al. (1998) found that trees constructed using 16S rRNA sequence data resulted in

the clustering of members of the genus Synechococcus, indicating that these strains were

closely related.

Leptolyngbya belongs to the most taxonomically problematic and poorly defined

genus (McGregor and Rasmussen 2008). To clarify this genus, further study of samples

from various environments is necessary (Lin et al. 2012). The heterogeneous Leptolyngbya

is one such cyano prokaryotic genus, which needs further study (Komarek 2007).

Leptolyngbya were created as new genera to include large number of oscillatorean species

with trichome up to 3µm wide (Anagnostidis and Komarek 1988).

In the present investigation, morphological characters of two cyanobacterial

isolates matches with the description provided by Anagnostidis and Komarek (1988) and

were classified in the genus Leptolyngbya. One of the isolate further classified with the

similarities in characters as L. foveolarum while the other isolate in the genus


61

Leptolyngbya did not matches with any of the species characters listed in the literature

(Anagnostidis and Komarek 1988; Singh et al. 2001; Komarek and Mares 2012). On the

basis of Bioguided assay L. foveolarum noted to be source of potent cytotoxic metabolites.

While another Leptolyngbya isolate also showed biological activity (antibacterial,

antifungal and cytotoxic). Therefore in the present investigation the two isolates of the

genus identified as Leptolyngbya foveolarum and Leptolyngbya sp. were confirmed by 16s

rRNA sequence.

The highest similarity for 16s rRNA was to Leptolyngbya foveolarum FM7988945

(100% identity), Leptolyngbya sp. HM 776036 (100%), Leptolyngbya sp. EF08833.1,

Leptolyngbya sp. AF132785 and Leptolyngbya boryana (100%). Using BLAST we

searched the NCBI database for the sequence that was most closely related and its closest

hits were Leptolyngbya foveolarum. Thus on the basis of morphological and 16 s rRNA

characterization the isolate is identified as Leptolyngbya foveolarum.

The sequence of Leptolyngbya sp. was not matches with the available sequence in

the NCBI library therefore it was identified as novel species. The sequence (562 bp) was

submitted to NCBI for accession number, Leptolyngbya sp. kmn-1(KC589411). The basic

local alignment search tool (BLAST) of the national center for biotechnology information

(NCBI) was utilized for locating the isolate sequences. Sequences of this Leptolyngbya sp.

have been deposited in GenBank under accession numbers KC589411.

Leptolyngbya foveolarum falls within the clade of the genus Leptolyngbya in all

analysis conducted using 16 s rRNA. The PCR amplified product of 16 s rRNA of another

isolate of Leptolyngbya on sequencing and pairwise alignment showed 95 % homology

with the 16 s rRNA of Leptolyngbya sp. (DQ134002), 93 % homology with Leptolyngbya

sp. (EUO68733), 92 % with Leptolyngbya sp. (EUO68733), Leptolyngbya frigidia

(AY493575), Calothrix sp. (GU186906) and Leptolyngbya sp. (FJ839352) available in

the NCBI database.

3.4.4. Growth of cyanobacteria in different culture media

The distribution of cyanobacterial species depends on chemo-physical environment and

organism ability to grow in a particular environment. This has led to development of

various culture media and was used for isolation and cultivation. The methods of

microalgal culture and basic culture media formulation were developed in the late 1800s

and early 1900s. Many of these methods and media formulations are in used today


62

(Anderson 2005). In the present investigation, the growth of cyanobacterial isolates

obtained in recommended nutrient media namely BG-11 (Stanier et al. 1971); Fogg’s

medium 1949; Jacobson, 1951); Allen and Arnon’s medium (Allen and Arnon 1955),

Zarrouk’s medium (Zarrouk 1966), CFTRI medium (Venkataraman and Becker 1984) is

depicted in Table 3.4. It was observed that the growth increase up to 4th

and 5th

week of

culture. In the 6th

week of culture the cyanobacterial filaments become yellowish whitish

and not showing any sign of growth. Decrease in the growth after certain time is a natural

phenomenon for any algal culture (Barsanti and Gualtieri 2006). Colla et al. (2007)

reported similar results in Spirulina platensis. Once the stationary growth phase is started,

the biochemical composition is changed (Vonshak 1997), which results in decrease in the

growth. The cultures of cyanobacteria pass through different growth phases. These growth

phases are lag phase, log phase, stationary phase and death phase (Becker 1994). Due to

different phases of growth of cyanobacteria, it is impossible to attain uniform growth

throughout the time of experiments. Similar observations have been recorded in our study

on cyanobacterial isolates. Initially the growth was very slow which increased

progressively up to the time of harvesting. Initially the culture mat was very thin hence

shading effect was not observed. Thickness of culture mat increased with time and it

resulted in self-shading effect because of which the inside cells did not receive sufficient

light and these cells might have undergone respiration (Vonshak 1983). Similar types of

results were obtained by Ramirez et al. (2000) in Calothrix sp. and Costa et al. (2001) in

Spirulina platensis.

The growth of Oscillatoria chalybea, Leptolyngbya foveolarum, Leptolyngbya sp.,

Microcoleus lacustris, Calothrix javanica, Phormidium fragile, Lyngbya bipunctata,

Westellopsis prolifica, Tolypothrix fragile, Scytonema mirabile and Scytonema

tolypothrichoides was more in BG-11 medium than in other media. However, Nostoc

entophytum, N. ellipsosporum, N. punctiforme, N. muscorum, N. calcicola and Anabaena

subcylindrica showed maximum growth in Fogg’s medium. CFTRI was the best medium

for the growth of Spirulina platensis. The variability in growth of cyanobacterial isolates

was attributed to the differences in recommended mineral salts and their concentrations in

the nutrient medium. The cyanobacterial growth also depends on the adaptation to the

chemo-physical environment (Anderson 2005). Spirulina sp. occurs in environments with

higher nutrient levels and varying temperature and salinity with pH 9.0– 10.0 and grows

well at 11.5 pH but not at 7.0 pH (Marquez et al. 1993, Vonshak and Tomaseli 2000,

Quasim et al. 2012). This might be the reason for less growth of Spirulina platensis in BG-

11, Fogg’s and Allen and Arnon media. While Anabaena sp. displays optimal growth at


63

pH 7.4–8.4 and its productivity decreases significantly at pH values higher than 9

(Marquez et al. 1993).

For optimum growth of cyanobacteria, appropriate Ka+: Na

+ ratio is required in

the cytoplasm (Barsanti and Gualtieri 2006). BG-11 medium consists of moderate

concentration of Na+ and in Allen and Arnon medium, Zarrouk’s medium and CFTRI

medium there is high concentration of Na+ while in Fogg’s medium; there is no Na

+

source. This might be the one of the reason for the variability in growth in cyanobacterial

isolates.

3.4.5. Large scale biomass production

Cultivation of green algae, Dunaliella and Chlorella and cyanobacteria Spirulina

in open and closed system has been extensively studied in the past few years and very

good results have been achieved for their biomass production (Vonshak 1997; Borowitzka

1999). However monoculture of these microalgae is possible only due to their specific

growth conditions such as Dunaliella grows in high salinity, Chlorella in high nutrition

and Spirulina in high alkalinity where with these condition growth and survival of other

microorganism is difficult (Lee 2001). For maximizing productivity in mass algal culture

system various growth conditions, physiological and technological approaches have been

proposed and investigated (Chaumont 1993; Richmond 1996; 2000; Lee 2001; Grobbelaar

2000; Anderson 2005; Wang et al. 2013).

In recent past, a number of cyanobacteria have been screened and recognized as a

rich but not yet extensively examined source of food, feed and pharmacological as well as

structurally interesting secondary metabolites (Becker 2004; Spolaore 2006). The major

constraint is of their biomass production in closed and open system is difficult. Therefore

till date little industrial and economic success is achieved (Ugwu et al. 2008; Wang et al.

2013). Closed culture system allow control over illumination, temperature, nutrient level,

contamination with predators and competing algae, whereas open culture system though

cheaper make it very difficult to grow specific cyanobacterial cultures for extended

periods and are more readily contaminated (Wang et al. 2013). It is also possible axenic

cultivation but it is too expensive and difficult because it requires a strict sterilization of all

glass wares, culture media and vessels to avoid contamination. These constrains make it

impractical and very expensive for commercial operation. On the other hand non axenic

cultivation cheaper and less laborious but associated with less predictable and inconsistent

quality (Lee 2001).


64

In the present investigation on use of tray - rack culture system it become possible

to achieve the biomass production of Spirulina platensis, Scytonema tolypothrichoides,

Leptolyngbya foveolarum, Nostoc calcicola, Tolypothrix fragilis, Westellopsis prolifica,

Calothrix javanica, Phormidium fragile, Lyngbya bipunctata, Nostoc entophytum and

Scytonema mirabile. Among these studied form maximum biomass production (162 g dry

weight/ rack) was observed for Leptolyngbya foveolarum. While minimum observed for

Westellopsis prolifica (59.9 g dry weight /rack) the other forms produced the biomass in

range of the 69-149 g/rack (Table 3.6). Under similar conditions the different forms

showed the variation for biomass production. This indicates that the requirement for

growth might vary with the cyanobacterial forms. Variation in biomass production was

also observed in the cyanobacterial forms grown in 0.3 L, 5 L, 10 L and 20 L bottles.

However, the produced biomass was low comparative to tray-rack culture system.

Although the height of nutrient medium level is more in bottle but in static condition the

growth of cyanobacterial forms was observed at the surface of the medium and some

forms also showed the growth at the wall of the bottle. This indicates that the buoyancy

characteristic and phototactic properties possess in cyanobacteria. This might limits the

growth and biomass production in rest of the volume of the medium in bottle culture of

system in static condition.

In tray-rack culture system on inoculation of less biomass (0.5 g FW /tray) not

resulted in proper growth and biomass formation. In this situation the growth of other

microorganisms (contaminant) was observed together with the inoculated cyanobacteria.

The proper growth and biomass formation and prevention of growth of other contaminant

organism was observed on addition of higher inoculum biomass (1 g FW /tray). According

to Lee (1986), Anderson (2005), Barsanti and Gualtieri (2006), high cell concentration is

necessary to achieve higher biomass productivity, and the need to maintain monoculture

for microalgae that grow in mild culture conditions.

As per the convenience and response of algal organism different types of culture

system have been developed the most routinely adapted include batch, continuous, semi-

continuous, ponds and photo-bioreactors. In the present investigation the tray-rack culture

system was developed (2.6 m H X 1.35 m L X 0.9m W) which can accommodate 160

trays (22.5 x 15 x 5.5 cm) in 11 compartments. This is a partially closed system. It is

equipped with drip system for addition of nutrient solution as per the requirement. On

addition of inoculum in trays with 1 lit medium within 1-3 days the inoculum floats and at

the surface forms a thin mat. This is the first sign of healthier growth of desirable


65

organism. In subsequent period, the cyanobacterial filament density increases constantly

and resulted in dark green cyanobacterial filament mat formation.

The nutrient medium components and water level decrease over time. Therefore at

two week period the fresh half strength nutrient medium was added to trays through drip

system. The evaporation of water is very high in summer season, showed the requirement

of addition of fresh medium and water at weekly interval. Maintenance of 2.5 to 3 cm

level of nutrient medium was suitable for proper growth and dark green filamentous mat

formation. Addition of more nutrient medium and presence of 4 to 5 cm level of nutrient

medium hamper the growth and filamentous mat formation in tray-rack culture system.

Application of drip system prevents the labor involved in physical addition of

nutrient medium. Provision of transparent plastic cover to the tray-rack culture system

helps to maintain the requirement of high humidity (80-90%) and being transparent allows

the passage of sufficient light for the growth (80-120 µmol m-2

s-1

). The tray culture unit

was placed under 50% category shade-net. Application of 50% of shade net was best to

filter the excess light intensity and also helps to prevent the increase of temperature.

Otherwise in open condition the culture becomes yellowish and finally whitish and did not

show any sign of growth.

The growth of cyanobacteria in tray –rack culture system showed a typical pattern

of growth which follows the sigmoid pattern, consisting of a succession of six phases,

characterized by variation in the growth rate. After the inoculation, 24 to 72 h no growth

or very slow growth was observed (i.e. the lag phase of growth curve). This is possible as

the cells may not be in condition to divide immediately or required time period to adapt to

the newer situation and start growing. In this situation the cell has to prepare with increase

in the level of enzymes and metabolites involved in cell division and carbon fixation

(Barsanti and Gualtieri 2006). After 72 h the mat formation started becoming dark-blue

green showed the good sign of growth. This continues for about four to five weeks after

inoculation. During the fifth and sixth week of culture the cells become yellowish in color

at the bottom. This might be due to shading of cells and not availability of sufficient light

and limiting of other factors such as nutrient, pH, carbon dioxide and other physical and

chemical factors (Anderson 2005). Similar trend in growth of Calothrix sp. was observed

in 12 days batch mass culture experiment with 8 g inoculum in 8 Lit disposal polythene

containers (Ramirez-Olvera et al. 2000). Similar results for different microalgal mass

culture system are difficult because of different geographic locations, culture strategies,

algae sp. etc. (Chaumont 1993). Biomass production in open pond system is


66

technologically simple but not necessary cheap due to the high down stream processing

cost and the products from such system could be marketed as value added health food

supplement, specially feed and alternative product for research (Lee 2001). In open pond

system the growth of desirable microalgae is affected by zooplankton, bacteria, virus

(Wang et al. 2013).

Despite that a great deal of work has been done to develop large scale microalgal

cultivation system, more efforts are still required to improve the technologies and know-

how of algal culture (Ugwu et al. 2008; Wang et al. 2013). In this context the attempts

made in present investigation in design, development and utilization of tray-rack culture

system is one of the good steps towards the improvement in large scale biomass

production of cyanobacteria.

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III. ISOLATION, IDENTIFICATION, AXENIC CULTURE AND BIOMASS …shodhganga.inflibnet.ac.in/bitstream/10603/106059/12/12_chapter3.… · Isolation, Identification, Axenic culture and