Asian Jr. of Microbiol. Biotech. Env. Sc. Vol. 17, No. (4) : 2015 : 1065-1070 © Global Science Publications ISSN-0972-3005

ISOLATION AND CHARACTERISATION OF MICROALGAE FROM UNIVERSITY OF MAURITIUS FARM FOR BIOETHANOL PRODUCTION RITESH BHAGEA1 AND DANESHWAR PUCHOOA1* 1

Faculty of Agriculture, University of Mauritius, Réduit, Mauritius (Received 14 August, 2015; accepted 17 October, 2015)

Key words : Bioethanol, Chrysophyceae, Microalgae, Molecular analysis. Abstract– Fossil fuels are depleting rapidly and are also the major sources of greenhouse gases, such as CO2 and CH4, which are causing global warming. A more promising alternative is the generation of biofuels from microalgae. This study was carried out to research on the potential of locally available microalgae for bioethanol production. Water samples were taken from the farm of the University of Mauritius and were analysed and spread on Bold’s Basal Medium agar. After 10 days of incubation, pure cultures were established and the growth of the isolated microalgae was optimised and scaled up in 5L plastic containers containing the Bold’s Basal Medium broth. The oven-dried harvested microalgae yielded a mean biomass of 0.2963g DW/L and total carbohydrate of 15.33 %( ±6.11) DW/L after acid hydrolysis. A mean biomass of 0.1363g DM/L and the total carbohydrate 21.36 %( ±12.28) DM/L was obtained when the isolated microalgae were cultured in a nitrogen deficient medium. A significant difference was observed only in the biomass yield. The presence of bioethanol was confirmed with qualitative tests after fermentation. Molecular analysis confirmed that the isolated pure cultured microalgae belonged to the Chrysophyceae class.

INTRODUCTION Human energy needs have been and are still being satisfied by fossil fuels on a very large scale. The world population is increasing at a fast rate and furthermore, the demand in energy is increasing proportionally. However, fossil fuels/oils are classified as non-renewable sources of energy and will be depleted by 2040 (Razzak et al., 2013). Plants are the largest source of biofuel, which is one of the alternatives to fossil fuels apart from solar, wind and nuclear energy. Corn and sugarcane bagasse have already proven to be good sources of biofuel in the form of bioethanol. However, the fact that they are food crops makes it difficult to use them for biofuel production as there is competition for food production and fertile land. Non food crops such as Arundo donax are also used as biomass for biofuel (Scordia et al., 2012). The fossil fuels have for long supplied electricity and fuel for transportation. In addition, the burning of fossil fuels produces huge volumes of CO2, NOX and SO2 which lead to global warming as they are greenhouse gases. Therefore, other environmental friendly sources of energy must be found and some *Corresponding author’s email: [email protected]

are wind energy, solar energy and geothermal energy. Biofuels such as bioethanol are best suited for transportation and the production can be done through microalgal culture. Biofuels also produced from microalgae, termed as 3rd generation biofuel, have many advantages such as no competition with food crops or derivatives no need for deforestation and their production is carbon neutral (Alam et al., 2012). Microalgae are tiny organisms that are called thallophytes as they are photosynthetic organisms but lack specialised organs (Alam et al., 2012). They can be found almost everywhere and are classified according to diverse characteristics (Gualtieri and Barsanti, 2006) such as colour, physical appearance or organisation. Nearly all microalgae contain proteins, carbohydrates and lipids but certain strains tend to have or produce more of one of them. This characteristic makes microalgae suitable for biofuel production in case lipids and carbohydrates are obtained at high levels. Dunaliela salina and Chlorella sp. are known to have high quantities of carbohydrates and lipids respectively (Efremenko et al., 2012). The aim of this work was to isolate and identify

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via molecular analysis the potential of local freshwater microalgae for the production of bioethanol. MATERIALS AND METHODS Sample collection identification

and

morphological

Water samples were taken from the pond of the University of Mauritius Farm. One sample consisted of 100 mL of pond water and was collected in sterile corning tubes. The samples were subjected to light microscopy for morphological identification of the microalgae present. Culture of microalgae with growth conditions For inoculation, 100μL of sample was aseptically spread on Bold’s Basal medium (BBM) agar. The plates were sealed with parafilm and kept in a culture room at 24 oC with a photoperiod of 14h Light: 10h Dark for 10 days with a light intensity of ~50μmol m-2 s-1. The most prominent microalgae was then cultured in 200 mL of BBM broth by direct inoculation with a loop-full of microalgae. Sterile air was supplied at a rate of 10L/min via a 0.2μm filter and an air stone. The culture conditions were maintained as mentioned previously. Two plastic containers of 5L were used for the culture of the microalgae for scale up. Working volume was 4L per container and the inoculum size was still 10% (v/v) at an absorbance of 0.05 with sterile air supply. The culture was monitored by spectrophotometric analysis at 680nm and harvested after 14 days. Nitrogen limitation to enhance carbohydrate accumulation Two 5L plastic containers were used to prepare 4L of modified BBM medium (MBBM) each. The medium was modified by omitting sodium nitrate and was inoculated with 10% of microalgae and left to grow for 14 days. Harvesting microalgae and acid hydrolysis The biomass was collected by centrifugation method at 8000rpm for 10 minutes at 4oC in round bottom corning tubes. The pellet was oven dried for 24 hours at 80oC. 0.25 g of dried sample was added to 50mL of 2N sulphuric acid solution and autoclaved at 121oC for 15 min according to parameters (except for biomass

added) established by Miranda et al., (2012). 3M NaOH was used to neutralize the hydrolysate to pH 5.5-6.0. The solution was then centrifuged at 8000rpm for 10 min. The supernatant was filtered with a filter paper to remove residues and then filter-sterilised using a 0.2μm sterile syringe filter. Total sugar analysis (Following Dubois et al., (1956) Fermentation and qualitative tests for bioethanol The yeast, Saccharomyces cerevisiae, was used for fermentation as described by Wang et al., (2014). The fermented samples were subjected to iodoform test, ester test and litmus paper test to investigate the presence of alcohol. DNA extraction and quantification Microalgal cells were cultured in BBM for 7 days. For the DNA extraction, a phenol/chloroform extraction method was used as described by Cheng and Jiang (2006). However, the protocol was modified by including sterile sand at the lysis step. Various mass of sand were tested in replicates of (g) 0.025, 0.05, 0.075, 0.1 and 0.0 as control. A 1.0% agarose gel was used to check the efficacy of the extraction protocol using sand. The gel was stained with ethidium bromide and viewed under UV light. For DNA quantification, analysis was carried out at 230, 260 and 280 nm. Polymerase Chain Reaction Three sets of primers were used for the amplification of the extracted DNA namely (forward – reverse); ITS1 – ITS4, fw1- rev1 and P45P47 (Abou-Shanab et al., 2011; Sonnenberg et al., 2007; Bérard et al., 2005). The products were run on 1.5% agarose gel and were sequenced. Sequences were then run on NCBI BLAST. RESULTS AND DISCUSSION Morphological attributes Figure 1 shows the isolated microalgae and Table 1 shows its morphological traits. The morphological traits of the isolated microalgae were relatively similar to microalgae such as Eudorina elegans and more specifically Tetrabaena socialis and Tetraspora gelatinosa. These microalgae belong to the Chlorophyta division but considering that the isolated microalgae lacked flagella and was greenish-yellow in colour, it was

Isolation and Characterisation of Microalgae from University of Mauritius Farm for Bioethanol 1067 1.8 1.6

Absorbance at 680nm

1.4 1.2 1.0 0.8 0.6 0.4 0.2 0.0 0

2

4

6

8

10

12

14

16

No of Days

Fig. 2. Growth curve of isolated microalgae after 14 days.

Fig. 1. Isolated microalgae (x400) Table 1. Morphological characteristics of the isolated microalgae Characteristics

Observations

Shape Colour

Spherical Yellow greenish to pale green Description 4 cells enclosed in transparent mucilage Presence of flagellate No Motile cells No Growth or arrangement pattern Colonial

assumed to belong to the Xantophyceae group (Alam et al., 2012). Growth analysis of microalgae Figure 2 shows the growth pattern that was observed with the isolated microalgae. A short lag phase was observed for the first 2 days with the samples followed by an exponential growth phase. Similar results were shown by AlShatri et al., (2014), where the growth curve established for Scenedemus dimorphus in various media and showed that among them, BBM broth was a suitable medium for the growth of microalgae. Ilavarasi et al., (2011) had same growth patterns with Chlorella sp and Monoraphidium sp, but the growth patterns differed from microalgae to microalgae which could, hypothetically, be due to different metabolic activities or other factors. Biomass production A significant difference was obtained in microalgae mean dry weight isolated from the BBM 1.1849±0.05g and Modified BBM 0.5450±0.06g

treatments for a volume of 4 Lculture were (p