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BIODEGRADATION OF PERSISTENT ORGANIC POLLUTANTS USING ALGAE AND CYANOBACTERIA:

AROMATIC AMINES DEGRADATION AFTER AZO DYE DECOLORIZATION

ALEJANDRO BRAVO HERMIDA

200611209

DEGREE PROJECT FOR BACHELOR’S DEGREE IN ENVIRONMENTAL ENGINEERING

SUPERVISORS:

SERGIO FERNANDO BARRERA TAPIAS

BO MATTIASSON

LUND, SWEDEN

UNIVERSIDAD DE LOS ANDES

Depto. de Ingeniería Civil y Ambiental

LUNDS TEKNISKA HÖGSKOLA (LUNDS UNIVERSITET)

Dept. of Biotechnology

2010

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ACKNOWLEDGEMENTS

I sincerely acknowledge:

In the first place, God, who has guided me and illuminated me, and who has made all this possible.

My parents, whose effort and support have been indispensable for me to complete this work.

Professors Sergio Fernando Barrera and Bo Mattiasson, my project supervisors, for their wise

advice and guidelines let me complete a good thesis work.

Adriano Cano Cuervo, coordinator for undergraduate programs at the Department of Civil and

Environmental Engineering, Universidad de los Andes, for all the help provided.

And all the people, who work in the laboratories at Uniandes and at LTH (Dept. of Biotechnology),

who patiently helped me with my work. In special, to Aida Juliana Martínez, Leonela Silva, Olga

Gómez, Marisa Punzi, David Svensson, Cecilia Orellana, Martin Hedström, Malik Badshah, and all

those people who were there ready to share their knowledge and help.

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CONTENTS

1. INTRODUCTION …………………...…………………………………………………………………………………………… 4

2. OBJECTIVES OF THE STUDY ……………………………………………………………………………………………………. 5

2.1 GENERAL OBJECTIVE …………………………………………………………………………………………………………… 5

2.2 SPECIFIC OBJECTIVES ............................................................................................................... 5

3. METHODOLOGY …………………………………………………………………………………………………………………….. 6

3.1 METHYL ORANGE DECOLORIZATION …………………………………………………………………………………… 6

3.2 AROMATIC AMINES DEGRADATION PART I ………………………………………………………………………….. 8

3.2.1 ANILINE EXPERIMENT ……………………………………………………………………………………………………….. 8

3.2.2 P-PHENYLENE DIAMINE EXPERIMENT ……………………………………………………………………………… 11

3.2.3 2,5-DIMETHYL ANILINE EXPERIMENT …………...……………………………………………………...…………. 11

3.2.4 ANILINE DUPLICATE EXPERIMENT …………………….…………………………………...…………………….... 14

3.2.5 2,5-DIMETHYL ANILINE DUPLICATE EXPERIMENT ................................................................ 14

3.3 AROMATIC AMINES DEGRADATION PART II ........................................................................... 14

3.3.1 DMA EXPERIMENT I ............................................................................................................. 14

3.3.2 DMA EXPERIMENT II ............................................................................................................ 15

3.3.3 DMA EXPERIMENT III ........................................................................................................... 15

4. RESULTS, ANALYSIS AND DISCUSSION ..................................................................................... 16

4.1 METHYL ORANGE DECOLORIZATION ..................................................................................... 16

4.2 AROMATIC AMINES DEGRADATION PART I ........................................................................... 19

4.2.1 ANILINE EXPERIMENT .......................................................................................................... 19

4.2.2 P-PHENYLENE DIAMINE EXPERIMENT ................................................................................. 22

4.2.3 2,5-DIMETHYL ANILINE EXPERIMENT .................................................................................. 24

4.2.4 ANILINE DUPLICATE EXPERIMENT ....................................................................................... 26

4.2.5 2,5-DIMETHYL ANILINE DUPLICATE EXPERIMENT ............................................................... 27

4.3 AROMATIC AMINES DEGRADATION PART II ........................................................................... 28

4.3.1 DMA EXPERIMENT I ............................................................................................................. 28

4.3.2 DMA EXPERIMENT II ............................................................................................................ 30

4.3.3 DMA EXPERIMENT III ........................................................................................................... 31

4.4 CELL IDENTIFICATION .............................................................................................................. 33

5. CONCLUSIONS ............................................................................................... ............................ 41

BIBLIOGRAPHY .................................................................................................. ............................ 42

ANNEXES ....................................................................................................................................... 45

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1. INTRODUCTION

The decolorization of dyes has been widely studied during the past years. Special focus has been

put on azo dyes (those whose molecular structure contains a –N=N- bond), because these kind of

compounds are ones of the most commonly used among industries, and are often toxic and

carcinogenic [5, 7-10, 12]. Several treatments have been proposed, regarding the importance of

color removal and detoxification of industrial wastewaters, since these kind of pollutants go

through conventional treatments unaffected [1, 9]. Such treatments, include photocatalysis

(usually mediated by TiO2), UV radiation, bacterial, algal and fungal biodecolorization, among

others. Recently, much research has been made about the ability of algae and cyanobacteria to

degrade azo dyes.

All of the above mentioned alternatives have shown excellent, or at least, relatively good results in

dye decolorization. However, to the best of our knowledge, studies have not gone any further

than identifying the decolorization metabolites, the aromatic amines, and do not give much

importance to the latter compounds, which are often equally or even more toxic than the dye

itself [9]. In fact, knowledge about degradation of such compounds by the different alternatives,

and specially, by algae and cyanobacteria, is scanty and more research about that is worth doing.

Therefore, the present work focuses on the capacity of algae and cyanobacteria to degrade

aromatic amines, rather than to decolorize an azo dye, which was, nevertheless, also done.

Decolorization of azo dyes occurs when there is a cleavage of the azo bond (-N=N-), which is the

chemical structure responsible for color in the molecule [24]. In addition, almost without

exception, azo dyes also contain at least one aromatic ring at each side of the azo bond.

Regardless on the degradation pathway the dye undergoes (photocatalysis, biodegradation), the

immediate metabolites generated are, therefore, aromatic amines [5, 8-10, 12, 19]. These

compounds are of equal importance since, as stated before, they can eventually be more toxic

than the parent compound, and should be removed as well before releasing the effluent into

natural water bodies.

Algae and cyanobacteria have shown to be capable of decolorizing azo dyes with at least relatively

good efficiency [6, 12]. Parikh et al. 2005, found about 40 different species of cyanobacteria living

in very polluted effluents from several dye stuff industries in India [13], while many other

researchers found that different species of algae and cyanobacteria can degrade azo dyes [6, 11,

12, 16, 19, 20, 21], aniline [12, 15, 17, 19], and other colorants [12], indicating that these

microorganisms can be able to adapt to such compounds. Three different degradation pathways

have been proposed when decolorizing with these microorganisms: biodegradation by azo

reductase enzymes [12, 19], and photo-generated reactive species by the cells [15, 17, 18]. The

latter pathway is surprising, since it is exactly the same principle as that of traditional

photocatalysis. Cáez and Barrera 2005, suggested that chlorophyll is the molecule responsible for

the photo-generation of reactive species in the cell [1].

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2. OBJECTIVES OF THE STUDY

2.1 General objective

The main objective of the present work is to evaluate the ability of algae and cyanobacteria to

degrade aromatic amines after decolorizing an azo dye.

2.2 Specific objectives

Evaluate the capability of previously unexposed algae and cyanobacteria to decolorize an

azo dye, Methyl Orange.

Evaluate the response of the same algae and cyanobacteria to the presence of primary

aromatic amines, namely, aniline, p-phenylene diamine and 2,5-dimethyl aniline, and their

capability to remove such compounds from water.

State whether there is true biodegradation, or else, if removal is only due to adsorption of

the aromatic amines to the biomass.

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3. METHODOLOGY

Although the methods employed for all experiments were nearly the same, each experiment was

carried on with specific variations, which are explained in the next sections.

3.1 Methyl orange decolorization

The Dye

Decolorization of an azo dye was the first step to take for the present study. Methyl orange is an

azo dye, often used as a pH indicator at laboratory. Thereby, it is easily obtained. It has got a

classical and simple molecular structure, according to Parshetti et al. 2010: one aromatic ring with

a sulphonate group (SO3) on the one side of the azo bond, and another aromatic ring with a N-

dimethyl group on the other side [21] (see Annex 1).

The algae

The algae and cyanobacteria were collected from stones surrounding a small mountain stream

nearby the city of Bogotá, Colombia. Their natural environment was rather cold and poorly

illuminated, due to the height of the place (nearly 2700 meters over the sea level), and the

abundant vegetation, which blocks much of the sunlight. Because it is a natural stream, coming all

the way down from the unspoiled mountain, the microorganisms certainly had never been

exposed to such pollutants before.

Use of consortia of microorganisms was advisable; when to be applied for major scale treatment,

it is practically impossible to maintain a single strain of microorganisms without being

contaminated, as well as the proper conditions for it to grow and degrade. Also, it has been

reported in literature that active consortia of microorganisms (although bacteria were used for

this case) has better results in degradation than single strains [9].

The growth medium

The growth medium used for growing the algae is described below in Table 1:

Table 1: BG-11 medium for growing cyanobacteria

Nutrient Amount (grams per liter of distilled water)

MgSO4•7H2O 0.075 CaCl2•2H2O 0.036 K2HPO4•3H2O 0.03 Na2CO3 0.02 Citric Acid 0.006

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H3BO3 * 2.86 MnCl2•4H2O* 1.84 ZnSO4•7H2O* 0.222 Na2MoO4•2H2O* 0.39 CuSO4•5H2O* 0.08

*Micro nutrients. These were poured in the amounts specified into a separate liter of water. 1 mL

of this solution was added to the main growth medium solution.

The preparation of this growth medium was done with some variations with respect to the

commonly used one: no nitrogen source of any kind was added, in order to force the growth of

nitrogen fixing cyanobacteria. Neither ferric EDTA was added, as the EDTA may constitute a carbon

source for non-photosynthetic organisms, whose presence was not desired for this study. Even

though this was done, some of these compounds (nitrogen and organic carbon containing) may

have come within the algae samples, allowing subsequent growth of many undesired

microorganisms, especially protozoa. Algal and cyanobacterial biomass, living or dead, itself, may

have served as a carbon source, as well as the citric acid, which was not used for the following

experiments. But, in spite of all this, the biomass was predominantly constituted by algae and

cyanobacteria, which was good enough for the study. Trying to kill everything that is not

cyanobacteria, would have probably been worthless, time consuming, and would have, perhaps,

misleaded the project from its main goal, so it was not done.

Procedures

Two glass bottles, of 1 liter capacity, were filled with 200 mL of growth medium. Samples were

collected and released into both bottles. Thereafter, the dye was added to both solutions in

different concentrations. One bottle had an initial concentration of 25 mg/L of methyl orange

(MO), and the other one had 5 mg/L of the same chemical. The bottle containing 25 mg/L MO was

named M1 and the one containing 5 mg/L MO was named M2.

The concentration of MO in each solution was monitored by color measuring through

spectrophotometry. Three mL of solution was taken and poured into a 3 mL volume, 1 cm path

length, glass made cubette for spectrophotometry, and analyzed with a visible-light

spectrophotometer. Scan was made from 315 nm to 700 nm, and the maximum absorbance peak

was found at 460 nm, being this consistent with literature [1, ]. The removal efficiency was

calculated as follows:

Where:

%E is the removal efficiency, expressed as a percentage;

Acl is the absorbance at 460 nm of the solution before adding the MO;

A0 is the absorbance at 460 nm of the solution just after having added the MO; and

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Ai is the absorbance at 460 nm of the solution at the time i of monitoring.

Monitoring was done every two or three days. The pH was kept around 7.1, since it is known to

affect the color intensity of a MO solution. Temperature was not controlled. It was subject to

environmental conditions. The Bottles were placed outside, in a terrace of the 8th floor of the

Mario Laserna Building, part of the main campus of the Universidad de los Andes, in Bogotá,

Colombia. The purpose of this was to approach the whole sunlight available, so that the

experiment would be conducted under more realistic conditions than those of a laboratory scale

experiment. The bottles were covered with plastic paper, to avoid insects and other external

things (like dust or other things that could eventually fall into the solutions, altering the process)

to get in, but small holes were opened for the air to circulate. The experiment was completed

within three weeks.

3.2 AROMATIC AMINES DEGRADATION PART I

All the following experiments were carried on in the laboratories of the Department of

Biotechnology, at Lunds Tekniska Högskola (Lunds Universitet), in Lund, Sweden. Since the climate

conditions there were not the same, and the sunlight availability is much less due to the time of

the year (autumn – winter), the experiments were run indoors, at room temperature (about 20°C)

and with permanent artificial fluorescent light (no light – dark periods). These experiments were

performed in order to evaluate if the same algae and cyanobacteria used in the previous

experiment can also survive in presence of aromatic amines, remove them from water, and if the

removal pathway is adsorption or true biodegradation. Three primary aromatic amines were used:

aniline (ANL), p-phenylene diamine (PFDA) and 2,5-dimethyl aniline (DMA).

3.2.1 ANILINE EXPERIMENT

Cell gathering

Two samples of cells were taken from each bottle after the MO experiment. The cells were

separated from the dye containing water (none of the solutions was 100% decolorized, as shown

further in the Results section) by centrifuging at 4000 rpm for 20 minutes. After separation, cells

corresponding to M1 and M2 were grown in two glass flasks apart, each one of 200 mL capacity.

Each flask had been previously filled with 200 mL of growth medium. The flasks were shaken in

order to spread the cells all over the space. The cells were grown for a week under permanent

artificial light, without adding any chemical or pollutant.

Growth medium

The growth medium used for this, and the following experiments, was somehow different to the

first one, shown above. No organic substance was added, and the chemicals used changed

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according to availability in the laboratory. For those which were not found, the closest chemicals

available were used, and a source of iron was also added. Table 2 shows the growth medium

prepared for these experiments:

Table 2: varied BG-11 medium used for the aromatic amines experiments.

Nutrient Amount (grams per liter of distilled water)

MgSO4•7H2O 0.075 CaCl2•2H2O 0.036 K2HPO4 * 0.03 Na2CO3 0.02 Fe(SO4)3 * 0.003 H3BO3 0.003* MnCl2•4H2O 0.002* ZnSO4•7H2O 0.0002* Na2MoO4•2H2O 0.0004* CuSO4•5H2O 0.0001*

*Chemical or amount changed, depending on availability. For the micronutrients, there was no

solution apart prepared, but they were poured directly into the main solution in the amounts

specified.

Procedures

After a week of growing, the two solutions M1 and M2 were injected with aniline. In order to state

if the aniline removal was truly due to the presence of algal and cyanobacterial cells, or else, to the

presence of growth medium, or either a spontaneous reaction in water, two control solutions

were prepared: one containing only growth medium and aniline (no cells), named B, and another

containing only distilled water and aniline, named W. All solutions were prepared in 200 mL glass

flasks. (Recalling, M1 is the solution containing the cells which were exposed to 25 mg/L of MO,

and M2 is the solution containing the cells exposed to 5 mg/L of MO).

The aniline was added to the solutions this way: 1 mL of pure aniline (approximately equal to 1

gram) was dissolved into 100 mL of distilled water. From this solution, 2 mL were taken and

poured into each flask. That makes 1 mL of aniline aqueous solution per 100 mL of solution.

The concentration of aniline in the solutions was monitored through UV-spectrophotometry, by

following the UV absorbance peak, which is found at 280 nm for this compound. Samples of 1 mL

were taken and centrifuged at 3500 rpm for 20 minutes (in this experiment, it was only done for

the final spectrophotometry test. In all the next experiments, it was done for all tests). A 1 mL

volume, 1 cm path length, quartz made cubette was used for the analysis. The removal efficiency

was calculated in terms of the dilution, as follows:

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Where:

Do is the initial dilution (mL ANL aqueous solution per 100 mL water), calculated in terms of the

initial absorbance;

Di is the dilution at the time i of monitoring, calculated in terms of the absorbance read at the

same time;

Ao, Acl, Ai and %E are the same parameters explained above, in the MO experiment.

Equations 2 and 3 correspond to the general equation obtained empirically by a calibration curve

for aniline; into 100 mL of distilled water, 0.1 mL of the aniline aqueous solution were poured, the

UV spectrum was obtained and the absorbance at 280 nm was read. 0.15 mL of ANL aqueous

solution was then added, and the same was done. This procedure was repeated until a dilution of

1.5 mL of ANL aqueous solution per 100 mL of distilled water was reached. The data were

collected and analyzed through Microsoft Excel. A linear regression was done, and the resulting

general equation was:

Where Y was the absorbance and X the dilution. The same was done with DMA, obtaining, of

course, different expressions. The Graph which plots the calibration curve (Absorbance vs.

Dilution) for aniline in water can be seen in Annex 2-a.

HPLC analysis

In order to confirm the results obtained by UV spectrophotometry, HPLC was done. An initial and a

final test were done in order to give more reliability. In addition, HPLC was also performed to

analyze the biomass, so that it could be stated if aniline had been adsorbed to the cells, and in that

case, how much of it was adsorbed.

HPLC procedures

Samples of 1 mL were taken from the solution flasks and centrifuged at 13000 rpm for 20 minutes.

The supernatant (water) was separated with a 2 mL syringe, and filtered through 0.45µm pore

nylon filters, and dropped into the HPLC vials. A C18 column, 250 mm x 4.6 mm, was used. The

mobile phase was isocratic, 60/40 water/acetonitrile in volume. The flow rate was set in 1 mL/min,

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the detection wavelength set at 280 nm and the retention time was of 25 minutes. The standard

chromatogram for aniline in water is shown in Annex 3-a.

Cell extraction

Possible aniline adsorbed to the cells was extracted as follows: 1 mL of the each solution, rich in

cells, was taken and centrifuged at 13000 rpm for 20 minutes. The supernatant was removed and

methanol was added. Cells were mashed in presence of methanol, and the methanol-cells solution

was again centrifuged at the same speed for the same time. The supernatant (methanol with cell

extracts) was removed and filtered through a 0.45µm pore nylon filter and dropped into the HPLC

vials. The HPLC parameters were exactly the same as the ones specified above. All the above

mentioned procedures were done according to those employed by Wang et al. 2007 [15]. The

standard chromatogram for aniline in methanol is shown in Annex 3-b.

3.2.2 P-PHENYLENE DIAMINE EXPERIMENT

After the experiment with Aniline, M1 and M2 were to be exposed to a some more complex

compound. Para-phenylene diamine (PFDA) was selected. New control solutions B and W were

prepared. 0.1 grams of PFDA were added to 200 mL of distilled water, and 2 mL of the latter

solution were poured into each solution flask. The growth media used, as well as the procedures,

were exactly the same as those employed for the ANL experiment. However, as it will be shown in

the Results section, calculations were not necessary.

3.2.3 2,5-DIMETHYL ANILINE EXPERIMENT

Once concluded the PFDA experiment, the same M1 and M2 were exposed to a different, also

more complex aromatic amine. 2,5-dimethyl aniline (DMA) was used. Again, new solutions B and

W were prepared. 0.05 mL (approximately equal to 0.05 g) of DMA were poured into 100 mL of

distilled water. 2 mL of this DMA aqueous solution were poured into each solution flask. Once

again, the same growth media and procedures were employed, except that the DMA peak is found

at 284 nm. Thereby, this was the detection wavelength for the HPLC analysis. The retention time

was also changed to 15 minutes. The calculations, nonetheless, are different.

Calculations

The calibration curve was obtained exactly in the same way as with the ANL. The general equation

which describes the Absorbance at 280 nm in terms of the Dilution (in mL per 100 mL) is the

following:

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Where Y is the Absorbance, and X is the Dilution. The percentage of removal or efficiency,

however, could not be calculated following the same methods used for aniline, because, since the

concentration is much lower, the UV peak is susceptible of changes in the base spectrum (the

spectrum of the solution, obtained before addition of DMA), and cannot be easily monitored, at

least in a numerical sense. Thus, it was necessary to develop a calculation algorithm, simple

although slanted, in order to estimate the removal percentage. The algorithm is now explained:

Figure 1-a. shows the spectrum of DMA at a dilution of 0.75 mL/100 mL in distilled water, while

Figure 1-b. shows the spectrum of DMA diluted in a solution containing algae plus growth medium

(upper curve), and the spectrum of the same solution before adding the compound (lower curve):

0

0,02

0,04

0,06

0,08

0,1

0,12

0,14

0,16

0,18

235 245 255 265 275 285 295 305

Fig. 1-a.

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By comparing both graphs, it is clear that the DMA spectrum is affected by the solution’s

spectrum; the resulting spectrum (upper curve, Fig 1-b.) is the sum of absorbances of both spectra.

Hence, when an important quantity of DMA is removed from water, the peak gradually moves

from 282 nm (where it initially appears) towards approximately 276 nm. From the calibration

curves, it was determined that the real DMA UV peak was found at 284 nm. A reference point was

set at 261 nm, where the lowest point, or valley, is situated. The difference in absorbance between

the peak at 284 nm and the point at 261 nm was measured from the calibration curves, and was

also found to be linearly correlated to the Dilution. Equation 7 expresses the absorbance in terms

of the dilution:

Where dA is the absorbance at 284 nm minus the absorbance at 261 nm and D is the dilution. The

difference between both points was then measured for the spectra of each solution before the

addition of DMA, which is negative. It was then assumed that such difference would be constant

throughout the experiment, which is not necessarily correct, but could give an approximation

good enough. Then, every time the UV spectrum was monitored, this difference was measured,

and the dilution (in mL/100mL) was calculated this way:

0

0,05

0,1

0,15

0,2

0,25

0,3

0,35

0,4

0,45

0,5

235 245 255 265 275 285 295 305

Fig. 1-b.

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Where dAr is the “real” difference in absorbance, dAi is the difference in abs. measured at the day

i and dAo is the difference measured before adding DMA, assumed as a constant (< 0). The result

was then replaced in equation 7, and D was left in terms of dAr and calculated. The removal

percentage was calculated according to Equation 4. The calibration curves for DMA in terms of

absorbance at the peak point and for the difference in absorbance between the peak and the low

point, are shown in Annexes 2-b and 2-c respectively.

3.2.4 ANILINE DUPLICATE EXPERIMENT

During the first ANL experiment, it had happened that, in the control solution B, after some 10

days, new algae had surprisingly grown in it and removed all the aniline. The cells of solutions M1

and M2 had been affected by the presence of PFDA and seemed to have trouble for living and

growing. Hence, these new algae, grown in B, were taken for carrying on a second experiment

with ANL. In this case, three 100 mL glass flasks were used. One was filled with 50 mL of growth

medium and 50 mL of the B solution (plenty of algae) and named M1, the next, filled with 100 mL

of growth media and named B, and the third one, with 100 mL of distilled water and named W.

The latter two, were kept again as control solutions. The procedures followed were exactly the

same as in the first ANL experiment, except for the HPLC column, which was changed for a 150

mm x 4.6 mm. The HPLC retention time was set at 15 minutes. The new standard chromatograms

for aniline in water and methanol are shown in Annexes 4-a and 4-b respectively.

3.2.5 2,5-DIMETHYL ANILINE DUPLICATE EXPERIMENT

Since the results obtained in the first were not clear, a duplicate of the DMA experiment was

found to be necessary. The same procedures as those described in the ANL duplicate and the first

DMA experiments were followed. Annexes 5-a and 5-b show the standard chromatograms for

DMA in water and methanol respectively.

3.3 AROMATIC AMINES DEGRADATION PART II

After the DMA duplicate, research was deepened in DMA removal by algae, in order to test the

capacity of the latter to remove such compound. The procedures followed were the same as those

described above, unless otherwise stated.

3.3.1 DMA EXPERIMENT I

Solutions M2 from the first DMA experiment and M1 from the DMA duplicate experiment were

taken for the following experiments, and renamed DMA1M2 and DMA2M1 respectively. The same

amount of DMA was added. Meanwhile, solutions M1 and B from the ANL duplicate experiment

were taken and renamed ANL2M1 and ANL2B respectively, and re-injected with a higher dilution

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(1.5 Ml/100Ml) of aniline, in order to grow them faster. No HPLC tests were carried on, either in

this, or in the 2nd DMA experiment.

3.3.2 DMA EXPERIMENT II

Once completed the last experiment, samples DMA1M2, DMA2M1, ANL2B and ANL2M1 were re-

filled with 1 mL/100mL DMA aqueous solution. 1 mL/100mL was injected to solution DMA2M1.

Before spectrophotometry analysis, samples were centrifuged at a speed of 7000 rpm for 10

minutes. The solutions were monitored for one week.

3.3.3 DMA EXPERIMENT III

This experiment was quite similar to the latter two, but with some differences: the centrifugation

speed was set at 13000 rpm for 15 minutes, before spectrophotometry; samples were not taken

daily; those solutions showing the fastest degradation were immediately refilled with DMA. HPLC

cell tests were done by the end of the experiment.

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4. RESULTS, ANALYSIS AND DISCUSSION

4.1 METHYL ORANGE DECOLORIZATION

As it was expected, the decolorization percentage was much higher in M2 than in M1, as the MO

initial concentration was less in the first one than in the second one. Absorbances were measured

nine times during the three weeks for both solutions. As it can be seen from Figure 2-a, M1

showed a very small removal at the beginning. Such removal can be due only to adsorption of the

dye to the cells. In fact, important removal rates were only registered by the end of the

experiment, as shown in Figures 2-b and 2-c.

0

0,2

0,4

0,6

0,8

1

1,2

1,4

1,6

1,8

310 360 410 460 510 560

Fig. 2-a: Spectra of M1 in time

Abs0

Abs (cl)

Abs1

Abs2

Abs3

Abs4

Abs5

Abs6

Abs7

Abs8

Abs9

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It is likely to have happened that, since the algae and cyanobacteria had never been exposed to an

azo dye before, MO removal in the first week might have probably been a consequence of

adsorption of the dye to the cells, not to any kind of degradation. Something different occurs,

nevertheless, with M2. A particularly high decolorization was registered only two days after

exposure to MO, up to around 17%. However, after five days, the removal had only been of an

additional 3% of the initial absorbance. The removal rate was kept low and then shifted again,

showing a very irregular behavior, as seen in Figures 3-a, 3-b and 3-c.

0

0,2

0,4

0,6

0,8

1

1,2

1,4

1,6

0 5 10 15 20

Fig. 2-b: Absorbance at 460 nm vs time (days) in M1

Abs.

0

5

10

15

20

25

30

35

40

0 5 10 15 20

Fig 2-c: % removal vs time (days) in M1

%Rem

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0

0,05

0,1

0,15

0,2

0,25

0,3

0,35

0,4

0,45

310 360 410 460 510

Fig. 3-a: spectra of M2 in time

Abs (cl)

Abs0

Abs1

Abs2

Abs3

Abs4

Abs5

Abs6

Abs7

Abs8

Abs9

0

0,05

0,1

0,15

0,2

0,25

0,3

0 5 10 15 20

Fig 3-b: absorbance of M2 vs time (days)

Abs.

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Such an irregular behavior can be explained by changing environmental conditions. Since the color

was much less intense in M2 than in M1, sunlight could have somehow been responsible for some

of the decolorization, either directly (UV light action on the MO molecules) or indirectly (possible

cell-generated reactive species). This could hence explain the high initial removal, combined with

cell adsorption. But when comparing Figs. 2-c and 3-c, a very similar behavior is noticeable,

indicating similar responses, which agrees with the fact that no cell had had any contact with a

pollutant like MO, and thus, no enzymatic reaction should be expected. Photosynthesis did not

seem to have any correlation with color removal; pH always shifted to high values (between 8 and

9) and, as a photosynthesis indicator, did not have any clear relation with the absorbance

measured. Recalling, pH was adjusted to 7 before spectrophotometry analysis.

4.2 AROMATIC AMINES DEGRADATION PART I

4.2.1 ANILINE EXPERIMENT

Wang et al. 2007 had achieved about 70% degradation of aniline by microscopic algae within only

four hours [15]. The present experiment did not show, nevertheless, the same results. After four

days of being exposed to aniline, both M1 and M2 algae did not show further removal than nearly

30%. However, a significant shift in removal was presented: control HPLC test was done, and

almost no aniline was found in either one solution containing algae (only around 1.1% of the initial

aniline was still present in M1 solution, and less than 1% in M2), while the aniline peak was still

present in B and W. Figures 4-a to 4-d, show the removal of aniline in each solution.

0

10

20

30

40

50

60

70

0 5 10 15 20

Fig 3-c: percentage removed vs time (days) by M2

%Rem

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0

0,5

1

1,5

2

2,5

3

3,5

4

4,5

210 230 250 270 290 310

Fig. 4-a: Spectra of M1 in time

Before Add.

After Add.

Day 2

Day 4

Day9

0

0,5

1

1,5

2

2,5

3

3,5

4

4,5

210 230 250 270 290 310

Fig. 4-b: Spectra of M2 in time

Before Add.

After Add.

Day 2

Day 4

Day 9

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As it can be seen, the yield for both cell solutions is quite similar. The initial relatively high aniline

removal might be due to reactions or adsorption between the compound and the growth media,

since the three solutions containing it, present the same curve shape. All these three solutions

presented an eventual change of color; at the beginning, the water was colorless (except for the

green in M1 and M2 due to chlorophyll), while a couple of days after addition, they acquired a

slight reddish-orange color. Thereafter, by the 7th day, M1 and M2 had this color no more, but

0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

0,9

1

0 2 4 6 8 10

Fig. 4-b: ANL dilution in time (days)

M1

M2

B

W

0

10

20

30

40

50

60

70

80

90

100

0 2 4 6 8 10

Fig. 4-c: ANL removal percentage in time (days)

M1

M2

B

W

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they had clearly turned green (much greener than at the beginning), which evidences cell growth.

B kept the reddish color for much longer, but, an interesting formation of flocs at the bottom of

the flask, as well as a very slim film on the water surface, were noticed. Flocks seemed to grow in

size and number within time and after two weeks of exposure, new algae had grown inside,

removing as well, all the aniline present. 7 days after addition, indeed, a considerable change in

aniline concentration was noticed. Since the spontaneous growth of algae coincided with a sudden

ANL disappearance, it can therefore be said, that cell growth and aniline removal are directly

correlated. Also, because the change in ANL concentration was shown faster in M1 and M2 than in

B, and indeed, a lot more than in W, it can be concluded that aniline is effectively removed from

water by algae and cyanobacteria. The small change in concentration in solution W, might be due

to molecule destruction by light, by oxygen in water, or else, to a physical process which might not

be volatilization of the compound. Annexes 6-a and 6-b show the chromatogram obtained after

one week of exposure for M1 and M2, respectively.

In order to state if aniline had been removed only due to adsorption to the cell biomass, cell

extraction with methanol was done, and analyzed through HPLC. Less than 1% of the initial aniline

was found in the cells, assuming 100% extraction by methanol. Nonetheless, even assuming lower

extraction values (~90% [15]), the percentage of aniline remaining in the cells would not be

greater than the above mentioned. Thus, it can be also concluded that aniline disappeared by

degradation pathways, and not by adsorption. The chromatograms obtained from cell extractions

of M1 and M2 can be seen in Annexes 7-a and 7-b respectively.

4.2.2 P-PHENYL DIAMINE EXPERIMENT

The experiment with PFDA was not successful at all. Apparently, when solid PFDA is poured into

water, it undergoes a spontaneous reaction; less than one hour before adding PFDA, water starts

acquiring a purplish wine color, which gets more and more intense with time. The color of the

PFDA aqueous solution (see section 3.2.2, page 10) was so intense, that no light could pass

through it. Besides, numerous small crystals of the same color were formed and stuck to the glass

walls of the flask. The same phenomenon happened in all four solutions. Indeed, the color was

much less intense, and no visible crystals were formed in the flask walls. Nevertheless, it was

clearly visible that the purple substance was adsorbed to the biomass, and showed to be

recalcitrant to the latter. In addition, the same phenomenon as that presented in the ANL

experiment’s B solution started to take place at the beginning, but the flocs were also covered

completely by the purple substance after some days (see Figure 6), and the cell growth was

completely inhibited. To support the hypothesis that PFDA reacts, deriving in another substance,

spectra of the solutions was obtained and showed in Figure 5:

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The spectrum of W after 5 days is difficult to interpret, but it is evident that, since it was only

water and PFDA, a spontaneous reaction takes place, as there is no clear UV peak. A slight peak

was detected at 442 nm (visible region), indicating absorbance due to visible color. The product of

the reaction is, presumably, a complex azo dye, due to the color presented, and which might be

recalcitrant to photosynthesis. It is also worth mentioning that standard HPLC tests were to be

done with PFDA. When filtering the samples through the nylon filter, the colored substance was

adsorbed to the nylon fibers, and water came out absolutely transparent. The latter is another

reason to think that the purple product is an azo dye. No PFDA peak was found in HPLC tests. PFDA

was therefore, not suitable for the present study.

0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

0,9

200 220 240 260 280 300 320 340 360

Fig 5: Spectra of PFDA in different solutions and time

W after add.

W Day 5

M1 Day 5

PFDA std.

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Figure 6: photograph taken to a floc formed in B solution with the microscope. The huge dark spot

on the cells, corresponds to that purplish substance which adsorbed to the cells, and could be

seen by naked eye.

4.2.3 2,5-DIMETHYL ANILINE EXPERIMENT

DMA seemed to be a more reliable and challenging compound for this work, so it was selected for

the next experiment. The results were, however, not very clear as with aniline. In fact, only a small

DMA removal was registered, and was attributed to adsorption to the cells. Figures 6-a to 6-c

show, respectively, the spectra of M1 and M2 and the DMA removal yield:

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0

0,02

0,04

0,06

0,08

0,1

0,12

0,14

210 230 250 270 290 310

Fig 7-a: Spectra of M1 in time

After Add.

Day 1

Day2

Day 3

Day 7

-0,02

0

0,02

0,04

0,06

0,08

0,1

0,12

0,14

210 230 250 270 290 310

Fig 7-b: Spectra of M2 in time

After Add.

Day 1

Day2

Day3

Day 7

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A better response from M1, however, could be noticed by telling for the spectra, than that

estimated numerically. But, anyway, the total DMA removal was not good. This was expectable,

since, in first place, DMA is a more complex compound than aniline, and second, cells had

previously been negatively affected throughout the PFDA experiment. At the end, the results from

M1 and M2 were not different from those of W. It was thus, necessary to make a duplicate

experiment.

4.2.4 ANILINE DUPLICATE EXPERIMENT

The duplicate experiment with aniline showed even better results than the first one: four days

after exposure, the algae had completely degraded the aniline, while a considerable cell growth

was noticed. Again, the reddish color was present in the solution, but disappeared some days

later. Figure 7 shows the spectra in time of M1. The same phenomenon presented in the first ANL

experiment took place in B.

0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

0,9

1

0 1 2 3 4 5 6 7 8

Fig 7-c: DMA removal by M1 and M2

M1

M2

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Again, very few aniline was found among the cell extracts, confirming aniline degradation. The

chromatogram obtained can be seen in Annex 8.

4.2.5 2,5-DIMETHYL ANILINE DUPLICATE EXPERIMENT

The results obtained by the present experiment, are far different from those obtained in the first

one. Significant removal rates were registered, while an important percentage of DMA seemed to

be degraded. After four days of exposure to DMA, the algae could remove up to 85%

approximately. Around 9% of DMA was found in the cells, after extraction with methanol (10% if

90% extraction capacity is assumed; see Annex 9). It means that more than 75% of the DMA was

presumably degraded by algae presence. Figure 8 shows the spectra of M1 within the four days:

0

0,5

1

1,5

2

2,5

3

210 230 250 270 290 310

Fig. 8: Spectra of M1 in time

Before Add.

Day 1

Day 3

Day 4

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Unlike with aniline, there was no noticeable cell growth when degradation occurred. Neither the

water turns reddish. The blank B also showed floc formation and some cell growth, and registered

some more than 50% removal. The DMA concentration in solution W remained nearly the same.

4.3 AROMATIC AMINES DEGRADATION PART II

4.3.1 DMA EXPERIMENT I

In first place, it was found that, after about two weeks, DMA had disappeared from solution

DMA1M2. However, the DMA removal rate was much lower in the latter than in DMA2M1, as

seen in Figure 9-c. The first solution achieved around 55% DMA removed, while the second one

attained more than 90% in one week. That could evidence that DMA1M2 cells were probably still

negatively affected. Again, no important cell growth was observed. Instead, the green color of cells

suddenly lost intensity, as evidence of some adverse effect of DMA on the biomass. However,

photosynthetic activity was still present, as the pH of the solution was always found at around 8.

The spectra of both solutions and the removal rate are plotted in Figures 9-a to 9-c.

0

0,1

0,2

0,3

0,4

0,5

0,6

210 230 250 270 290 310

Fig 9: Spectra of M1 in time

Before add.

After add.

Day 1

Day 3

Day 4

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0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

0,9

210 230 250 270 290 310

Fig 10-a: Spectra of DMA1M2 in time

Before Injection

After Injection

Day2

Day4

0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

210 230 250 270 290 310

Fig. 10-b: Spectra of DMA2M1 in time

Before Injection

After Injection

Day2

Day5

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Cells from DMA2M1 probably removed DMA better because of their previous exposure to the

compound, while those from the other solution were, perhaps, recovering from the adverse

conditions they had faced with PFDA and DMA. The small shift in the removal graph (Fig. 9-c) can

be explained by an abnormal spectrum curve flattening. As the dilution was calculated based in

the difference between absorbances read at 261 nm and 284 nm, a particular change in the base

absorbances (those assumed as constant, read before adding the compound), originated perhaps

in centrifugation, or a possibly contaminated baseline, could be the reason for this strange datum

registered in both solutions. Another explanation could be an unusual shift in the water

evaporation rate, big enough to heighten the DMA concentration. This is however, very unlikely to

happen, and was not noticed by naked eye. Nevertheless, regardless of this abnormality, a decay

tendency (perhaps, exponential) could explain the rest of the data, especially in the case of

DMA2M1.

4.3.2 DMA EXPERIMENT II

The two solutions were then refilled with DMA, and so were the solutions ANL2B and ANL2M1.

Again, solution DMA1M2 showed the lowest removal, while DMA2M1 had the highest, since 1.5

mL of DMA aqueous solution was injected to that solution. The second best removal was

presented in solution ANL2B. The results obtained, are plotted in Figure 10:

0

0,2

0,4

0,6

0,8

1

1,2

0 1 2 3 4 5 6 7 8

Fig. 10-c: DMA removal in time (days)

DMA1M2

DMA2M1

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The results went according to the expected, as DMA2M1 had had the largest exposure to the

compound. Nevertheless, the removal presented in ANL2B was surprising, since it was expected to

have a much lower one, as those cells had had the shortest exposure to aromatic amines. The

results obtained for both solutions, as well as for ANL2M1 can be explained by an initial adsorption

of DMA to the biomass, followed by similar degradation rates, which is evidenced in Figure 10.

It was interesting to observe that, while ANL2M1 cells remained totally green and abundant, those

of ANL2B seemed to be a little diminished and their color turned from green to a reddish brown.

Such color could be evidence of DMA adsorption to the cell walls, or otherwise, an adverse effect

of the compound on the cells, or perhaps, a combination of both options. As an interesting

observation, those two solutions whose color had changed, had better removal rates than that of

ANL2M1, whose cells remained green. Different results would be attained in the third and last

experiment.

4.3.3 DMA EXPERIMENT III

The idea of this final experiment was to push cells to remove more and more DMA, in order to see

if they were capable of dealing with frequent discharges of the pollutant. In order not to force the

cells so much that they would die, only those showing the best removal rates were refilled with

DMA.

0

0,2

0,4

0,6

0,8

1

1,2

1,4

1,6

0 1 2 3 4 5 6 7 8

Fig. 11: DMA remotion yields in each solution

DMA1M2

DMA2M1

ANL2B

ANL2M1

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As seen from Figure 11, the removal yields of all solutions, was different. DMA2M1 could not

degrade enough for refilling to be made; only ANL2B and ANL2M1 were refilled with DMA and

showed significant removal rates, being the first one’s lower than that of the second one. By the

13th day, only ANL2M1 could achieve nearly 100% DMA removal after refilling, showing thus the

best removal yield. DMA2M1 also achieved nearly 100% removal, but without being refilled. This

goes according to the expected at the beginning of DMA experiment II, since ANL2M1 cells

seemed healthier, as well as larger in population. This also gives support to the idea that DMA

might have some adverse effect on algal and cyanobacterial cells.

An alternative explanation for the results obtained, is that DMA is only adsorbed to the biomass,

and since DMA2M1 cells had been exposed in repeated times to relatively large amounts of the

compound, and besides, no significant cell growth was presented, the adsorption rate decreased

over time. Something similar could have happened to ANL2B cells, although they were much less

exposed to DMA. This hypothesis could also explain the behavior presented by solution DMA1M2,

whose cells would have adsorbed huge amounts of DMA compared to all the others, and to the

purple PFDA product. ANL2M1’s behavior is not very well explained by this hypothesis, though. In

order to have evidence that could support or deny such hypothesis, extraction with methanol was

once again made. As confirming evidence, a huge peak at around minute 7 of retention time

would be expected, which would mean large amounts of DMA. Otherwise, a very small peak, or,

indeed, no peak at all, would prove that removal is due to degradation, thus, giving support to the

first hypothesis and denying the second one.

0

0,2

0,4

0,6

0,8

1

1,2

0 2 4 6 8 10 12 14

Fig 12: removal yields of all solution in 13 days

DMA1M2

DMA2M1

ANL2B

ANL2M1

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The cell extracts were then analyzed through HPLC, but practically no DMA was found in them (see

Annexes 10-a to 10-d). Therefore, it can be said that DMA is removed from water through

degradation pathways, and not by adsorption to biomass. Evidence good enough to support the

hypothesis of degradation of primary aromatic amines, mediated by algae, was thus provided by

performing cell extraction with methanol, and analyzing such extracts through HPLC.

Whether enzymatic action or cell generated reactive species are responsible for the degradation

was not stated in the present work, since no additional experiments were performed, which would

give evidence of one thing or another. According to Wang et al. 2007, aniline removal from water

is a consequence of photo-generated reactive species into the cells, some of which are eventually

released by the latter to the external environment, being these responsible for the degradation of

aniline [15]. Such theory was supported by other studies [17, 18]. This makes sense for the present

study, since previously unexposed algae were able to degrade MO, ANL and DMA. But the action

of enzymes is also a valid explanation, at least for the case of aniline, as it could be proved that its

presence favors and stimulates algal growth; it happened 100% of the times it was done, and even

each time faster. Besides, algae would not be greatly effective as a photocatalyst, since they might

also produce antioxidants to protect themselves from the reactive species [15].

4.4 CELL IDENTIFICATION

Cell identification was considered to be worth doing; knowing which species or at least, families of

algae and cyanobacteria were responsible for removal of the different compounds studied. It

could also provide additional information which could explain the results obtained. The

identification process was very simple: samples of biomass were taken and seen with the

microscope. Pictures of the images were taken and, comparing to literature [22, 23], some families

and species were identified. Those are listed next:

Chlorophyceae filamentosa sp.

Monoraphydium sp.

Oocystis sp.

Cyanophyceae sp.

Oscillatoria sp.

Pseudoanabaena sp.

Pseudoanabaena cf. planctonica

Sphaerocystis sp.

Johannesbaptistia cf. pellucida

Surirella sp.

Scytonema sp.

Lyngbya sp.

Nitzchia spp.

Elkanothryx sp.

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Scedenesmus sp.

Phacus sp.

Anabaena sp.

Aphanocapsa sp.

Microcystis sp.

Synechococcus sp.

Synechocystis sp.

Chlorococcales sp.

Chrococcus sp.

The most abundant species were the Chrococcus sp., Oocystis sp. and Chlorococcales sp.

Nevertheless, different combinations of species were noticeable in each solution. For example,

Oocystis sp. and Chrococcus sp. were the most numerous species in solutions M1 and M2 when

the first experiments (MO, ANL, PFDA, DMA) were run, although many different species could be

seen (Figures 14-a and 14-b). Bacterial growth would have been, after all, also expected.

Moreover, in the DMA experiments II and III, solution ANL2M1 was rich in Scedenesmus species,

while in ANL2B, DMA2M1 and DMA1M2, Chrococcus sp. were the dominant species, as it can be

seen in Figures 13-a to 13-d. Such a variation could provide an extra explanation for the results

obtained in the DMA experiment III; The high removal yield registered in ANL2M1 compared to

the other solutions, could be influenced by the abundant presence of Scedenesmus sp., which is

scanty in the other three solutions. Chrococcus species, could be negatively affected (lowered

response capacity) after several repeated times of being exposed to DMA, while Scedenesmus

species, on the other hand, could probably increase their response capacity to DMA presence.

That would be evidence of better adaptability of the latter species than that of the Chrococcus

The variation in species diversity could have been an additional factor that affected pollutant

removal, in as much as the adaptability varies from one species to another and so do aspects like

cell wall composition, response capacity and velocity and enzyme production, among others.

IAMB 201020-06

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Fig. 13-a: cells from ANL2M1. Scedenesmus sp. is the most abundant, although not the only one.

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Fig. 13-b: cells from ANL2B. Chrococcus sp. are the most abundant, but more diversity is seen.

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Fig. 13-c: Cells from DMA2M1. Chrococcus sp. dominance is evident. Some Anabaena filaments are

seen by the left lower side of the picture.

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Fig. 13-d: cells from DMA1M2. Chrococcus sp. are the most common, but other cyanobacteria

(probably Cyanophyceae or Pseudoanabaena cf. planctonica) are frequently found.

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Fig. 14-a: Cells from M1 during the first ANL experiment.

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Fig 14-b: cells from M2 during the first ANL experiment.

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5. CONCLUSSIONS

Previously unexposed algae and cyanobacteria were able to remove partially methyl

orange, an azo dye. The removal yield depended, as expected, on the initial dye

concentration, and probably on the climatic conditions (sunlight intensity due to cloud

presence or absence, temperature, etc). The decolorization of MO could have been, in

part, due to adsorption to the cells, and probably to photo-production of reactive

oxidative species mediated by algal and cyanobacterial cells. There is no evidence of

enzymatic reaction, which was expected, since the algae had never been exposed to

pollutants like MO before.

Aniline is removed from water in presence of algal and cyanobacterial cells and, in a small

part, to growth media, but there is no doubt that the main cause for ANL removal is the

presence of the first ones. Aniline significantly stimulates algal and cyanobacterial cell

growth, hence, stimulating photosynthesis as well. Action of enzymes produced by cells is

most likely to be the main cause of aniline removal and, and slightly, adsorption at the

beginning. Photo-produced reactive species by cells does not seem to play an important

role in removing ANL from water. At last, aniline is completely consumed by the cells, not

adsorbed or stored inside them.

P-phenylene diamine undergoes an spontaneous reaction in contact with water, and was

therefore useless for the purposes of this study. Such reaction is unknown, but there are

hints which indicate that the product might be a dye, perhaps azo type. Such product is

highly recalcitrant to algal activity.

2,5-dimethyl aniline was effectively removed from water by algae and cyanobacteria,

although with lower yields than those presented with aniline, which is expected, since this

compound is chemically more complex. Unlike aniline, DMA does not favor cell growth. On

the opposite, it could have some adverse effect on cells, although photosynthetic activity

did not seem to be affected. In addition, since very few or no DMA at all was found in the

cell extracts, removal could be attributed to photo-produced reactive species by the algal

and cyanobacterial cells, and not to enzymatic action.

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REFERENCES

1. Cáez Pérez, M.A., Barrera Tapias, S.F. (2005). Decoloración del anaranjado de metilo a través de

un proceso de fotocatálisis con clorofila activada. Universidad de los Andes, Bogotá D.C.,

Colombia.

2. García, A., Amat, A.M., Arques, A., Sanchís, R., Gernjak, W., Maldonado, M.I., Oller, I., Malato, S.

(2006). Detoxification of aqueous solutions of the pesticide ‘Sevnol’ by solar photocatalysis.

Environmental Chemistry Letters. Volume 3, Number 4.

3. Bumpus, J.A., Tricker, J., Andrzejewski, K., Rhoads, H., Tatarko, M. (1999). Remediation of water

contaminated with an Azo Dye: an undergraduate laboratory experiment utilizing an inexpensive

photocatalytic reactor. Journal of Chemical Education. Volume 76, Issue 12.

4. Hong, J., Ma, H., Otaki, M. (2005). Controlling algal growth in photo-dependent decolorant

sludge by photocatalysis. Journal of Bioscience and Bioengineering. Volume 99, Issue 6.

5. Xingzu, W., Xiang, C., Dezhi, S., Hong, Q. (2008). Biodecolorization and partial mineralization of

Reactive Black 5 by a strain of Rhodopseudomonas palustris. Journal of Environmental Sciences.

Volume 20, Issue 10.

6. Parikh, A., Madamwar, D. (2004). Textile dye decolorization using cyanobacteria. Biotechnology

Letters. Volume 27, Issue 5.

7. Ayed, L., Achour, S., Khelifi, E., Cheref, A., Bakhrouf, A. (2010). Use of active consortia of

constructed ternary bacterial cultures via mixture design for Congo Red decolorization

enhancement. Chemical Engineering Journal. Volume 162, Issue 2.

8. Kolekar, Y.M., Pawar, S.P., Gawai, K.R., Lokhande, P.D., Shouche, Y.S., Kodam, K.M. (2008).

Decolorization and degradation of disperse blue 79 and acid orange 10 by Bacillus fusiformis KMK5

isolated from the textile dye contaminated soil. Bioresource Technology. Volume 99, Issue 18.

9. Khehra, M.S., Saini, H.S., Sharma, D.K., Chadha, B.S., Chimni, S.S. (2005). Comparative studies on

potential of consortium and constituent pure bacterial isolates to decolorate azo dyes. Water

research. Volume 39, Issue 20.

10. Guang-fei Liu, Ji-ti Zhou, Jing-Wang, Zhi-jong Song, Yuan-yuan Qv. (2006). Bacterial

decolorization of azo dyes by Rhodopseudomonas palustris. World Journal of Microbiology and

Biotechnology. Volume 22, Issue 10.

IAMB 201020-06

43

11. Karacakaya, P., Koçberber Kiliç, N., Duygu, E., Dönmez, G. (2009). Stimulation of reactive dye

removal by cyanobacteria in media containing triacontanol hormone. Journal of Hazardous

Materials. Volume 172, Issue 2-3.

12. El-Sheekh, M.M., Gharieb, M.M., Abou-El-Souod, G.W. (2009). Biodegradation of dyes by some

green algae and cyanobacteria. International Biodeterioration and Biodegradation. Volume 63,

Issue 6.

13. Parikh, A., Shah, V., Madamwar, D. (2005). Cyanobacterial flora from polluted industrial

effluents. Environmental Monitoring and Assessment. Volume 116, Issue 1-3.

14. Khataee, A.R., Dehghan, G., Ebadi, A., Zarei, M., Pourhassan, M. (2010). Biological treatment of

a dye solution by Macroalgae Chara sp.: Effect of operational parameters, intermediates

identification and artificial neural network modeling. Bioresource Technology. Volume 101, Issue

7.

15. Lei Wang, L., Zhang, C., Wu, F., Deng, N. (2007). Photodegradation of aniline in aqueous

suspension of microalgae. Journal of Photochemistry and Photobiology. Volume 87, Issue 1.

16. Ertuğrul, S., Bakir, M., Dönmez, G. (2008). Treatment of dye-rich wastewater by an immobilized

thermophilic cyanobacterial strain: Phormidium sp. Ecological Engineering. Volume 32, Issue 3.

17. Zepp, R.G. (US EPA), Schlotzhauer, P.F. (1983). Influence of algae on photolysis rates of

chemicals in water. Environmental Science and Technology. Volume 17, Number 18.

18. Liu, X., Wu, F., Deng, N. (2004). Photoproduction of hydroxyl radicals in aqueous solution with

algae under High Pressure Mercury Lamp. Environmental Science and Technology. Volume 38,

Number 1.

19. Acuner, E., Dilek, F.B. (2004). Treatment of Tectilon Yellow 2G by Chlorella Vulgaris. Process

Biochemistry. Volumen 39, Issue 5.

20. Wan-Loy Chu, Yike-Chu See, Siew-Moi Phang. (2008). Use of immobilised Chlorella vulgaris for

the removal of colour from textile dyes. Journal of Applied Phycology. Volumen 21, Number 6.

21. Parshetti, G.K., Telke, A.A., Kalyani, D.C., Govindwar, S.P. (2010). Decolorization and

detoxification of sulfonated azo dye methyl orange by Kocuria rosea. Journal of Hazardous

Materials. Volume 176, Issues 1-3.

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22. Pérez-G, D., Rivera-Rondón, C. (2007).Estructura de las comunidades fitoplanctónicas y

perifíticas en sistemas de inundació del Río Orinoco (Puerto Carreño, Vichada) en época de sequía.

Pontificia Universidad Javeriana-Fundación Omacha.

23. Shah, V., Garg, N., Madamwar, D. (2000). Record of the cyanobacteria present in the Hamisar

pond of Bhuj, India. Acta Botánica Malacitana. Volume 25, Issue 85.

24. Needles, H.L. (2001). Textile fibers, dyes, finishes and processes. University of California, Davis,

California. First Indian Edition, 2001. Standard Publishers Distributors, Delhi, India.

IAMB 201020-06

45

ANNEXES

Annex 1: Molecular structure of Methyl Orange and its degradation pathway, according to

Parshetti et al. 2010. [21]

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Annex 2-a: calibration curve for aniline dissolved in water, based on the measurement of the UV

absorbance peak at 280 nm.

Annex 2-b: Calibration curve for DMA in water, based on the measurement of the UV absorbance

peak at 284 nm.

y = 1,4223x + 0,0658R² = 0,9926

0

0,5

1

1,5

2

2,5

0 0,2 0,4 0,6 0,8 1 1,2 1,4 1,6

Abs. 280 nm

Abs.

Lineal (Abs.)

Lineal (Abs.)

y = 0,0943x + 0,0062R² = 0,9983

0

0,02

0,04

0,06

0,08

0,1

0,12

0,14

0,16

0 0,5 1 1,5

Abs. (284 nm)

Abs. (284)

Lineal (Abs. (284))

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Annex 2-c: Calibration curve for the difference between absorbance at 284 nm and absorbance at

261 nm in DMA UV spectrum.

Annex 3-a: Standard chromatogram for aniline in water with a 250 mm long column.

y = 0,062x + 0,0007R² = 0,9977

0

0,02

0,04

0,06

0,08

0,1

0,12

0 0,5 1 1,5

A284-A261

A284-A261

Lineal (A284-A261)

1.76

5.99

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

-5

0

5

10

15

20

25

Intensity (mV)

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Annex 3-b: Standard chromatogram for aniline in methanol with 250 mm long column.

Annex 4-a: Standard chromatogram for aniline in water with 150 mm long column.

2.68

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

0

100

200

300

400

Intensity (mV)

3.64

0 2 4 6 8 10 12 14 16 18

Retention Time (min)

0

20

40

60

80

100

120

140

160

Intensity (mV)

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Annex 4-b: Standard chromatogram for aniline in methanol with 150 mm long column.

Annex 5-a. Standard chromatogram for DMA in water with 150 mm long column.

1.51 2.72

3.76

7.460 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

0

100

200

300

400

Intensity (mV)

5.77

6.91

9.84

0 2 4 6 8 10 12 14 16 18

Retention Time (min)

0

20

40

60

80

100

120

Intensity (mV)

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Annex 5-b: Standard chromatogram for DMA in methanol with 150 mm long column.

Annex 6-a: chromatogram for M1 solution after 7 days of exposure to aniline.

1.51 2.71 4.67 6.887.54

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

0

100

200

300

400

500

600

700

800

Intensity (mV)

1.76

6.61

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

-7.4

-7.2

-7.0

-6.8

-6.6

-6.4

-6.2

-6.0

Intensity (mV)

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Annex 6-b: chromatogram for M2 solution after 7 days of exposure to aniline.

Annex 7-a: chromatogram for cell extracts of M1 after aniline removal.

1.76

6.29

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

-7.4

-7.2

-7.0

-6.8

-6.6

-6.4

-6.2

Intensity (mV)

1.061.66

2.213.02

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

-1.5

-1.0

-0.5

0.0

0.5

Intensity (mV)

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Annex 7-b: chromatogram for cell extracts of M2 after aniline removal.

Annex 8: chromatogram for cell extracts of ANL2 M1 after aniline removal.

1.05

2.21

7.45

0 2 4 6 8 10 12 14 16 18 20 22 24

Retention Time (min)

-1.5

-1.0

-0.5

0.0

0.5

Intensity (mV)

3.71 7.46

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-1

0

1

2

3

4

5

6

7

Intensity (mV)

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Annex 9: chromatogram for cell extracts of DMA2 M1.

Annex 10-a: chromatogram for cell extracts of ANL2B.

Annex 10-b: chromatogram for cell extracts of ANL2M1.

3.70 7.40

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-2

0

2

4

6

8

10

12

Intensity (mV)

2.77

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-2

0

2

4

6

8

10

12

Intensity (mV)

2.77 7.84

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-2

0

2

4

6

8

10

12

Intensity (mV)

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Annex 10-c: chromatogram for cell extracts of DMA1M2.

Annex 10-d: chromatogram for cell extracts of DMA2M1.

2.77 7.85

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-2

0

2

4

6

8

10

12

Intensity (mV)

2.77

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14

Retention Time (min)

-2

0

2

4

6

8

10

12

Intensity (mV)