Photosynth Res DOI 10.1007/s11120-016-0294-2
ORIGINAL ARTICLE
Metabolism of xenobiotics by Chlamydomonas reinhardtii: Phenol degradation under conditions affecting photosynthesis Theocharis T. Nazos1 • Emmanouel J. Kokarakis1 Demetrios F. Ghanotakis1
•
Received: 15 June 2016 / Accepted: 6 July 2016 Ó Springer Science+Business Media Dordrecht 2016
Abstract In the present work, the biodegradation of phenol by axenic cultures of the unicellular microalga Chlamydomonas reinhardtii was investigated. Biodegradation proved to be a dynamic bioenergetic process, affected by changes in the culture conditions. Microalgae biodegraded defined amounts of phenol, as a result of the induced stress caused at high concentrations, despite the fact that this process proved to be energy demanding and thus affected growth of the culture. High levels of biodegradation were observed both in the absence of an alternative carbon source and in the presence of acetate as a carbon source. Biodegradation of phenol by Chlamydomonas proved to be an aerobic, photoregulated process. This is the first time that Chlamydomonas reinhardtii has been used for bioremediation purposes. This study has demonstrated that the most important factor in the biodegradation of phenol is the selection of the appropriate culture conditions (presence or absence of alternative carbon source, light intensity, and oxygen availability) that provide the best bioenergetic balance among growth, induced stress, and biodegradation of phenol.
CO2 Acetate ? CO2 Limit C t X0 X S0 S l YX/S
F0
Fmax Keywords Chlamydomonas reinhardtii Phenol Biodegradation Photosynthesis Bioenergetics Stress Abbreviations PCV Acetate
Packed cell volume Treatment with acetate as carbon source carried out into light
& Demetrios F. Ghanotakis
[email protected] 1
Department of Chemistry, University of Crete, Vasilika Voutes, 70013 Heraklion, Crete, Greece
Fv Fv/Fmax ABS/RC DI0/R C RC/CS0 TR0/ABS Ca Cb
Treatment with carbon dioxide as carbon source carried out into light Treatment with both acetate and CO2 as carbon sources carried out into light Treatment with limited carbon source carried out into light Time in days Total initial cell biomass in the culture (mg) at the beginning of the experiment Total cell biomass in the culture (mg) at time t Total initial amount (mg) of phenol at the beginning of the experiment Total amount (mg) of phenol at time t Specific growth rate Observed growth yield calculated according to the equation YX/S = (X – X0)/(S0 - S) Minimum fluorescence that corresponds to the time that all photosynthetic reaction centers are open Maximal fluorescence that corresponds to the time that all reaction centers are closed Variable fluorescence (Fmax - Fo) Photosynthetic efficiency Size of the functional antenna per active reaction center Dissipation energy per active reaction center Active reaction center density Quantum yield of primary photochemistry Concentration of chlorophyll-a Concentration of chlorophyll-b
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Cx?c nd
Concentration of total carotenoids Not detected
Introduction Chlamydomonas reinhardtii is a unicellular microalga that has been used as a model organism in photosynthesis research. Chlamydomonas cells are haploid and consist of a nucleus, one chloroplast, a cell wall, a photosynthetic antenna, and two flagella (Rochaix et al. 1998). The presence of high concentrations of phenolic compounds in the environment is the result of anthropogenic activity. These compounds penetrate ecosystems as the result of drainage from municipal or industrial sewage into surface water (Jaromir et al. 2005). Phenol is used as an intermediate in the chemical industry for the synthesis of more complex compounds (Keith and Telliard 1979). The toxicity of phenolic compounds stems mainly from their hydrophobic character as well as their ability to form free radicals (Hansch et al. 2000). Many microorganisms, mainly bacteria and fungi, have been used for the biodegradation of phenolic compounds (Varsha et al. 2011). The idea of using microalgae in biodegradation processes was initially presented by Oswald and Gotaas (Oswald and Gotaas 1957), who proposed the use of engineered photosynthesis as a method for simultaneous oxygen production and treatment of wastes. There are many advantages of using algal species in bioremediation processes for wastewater treatment: (i) cost effectiveness; (ii) low energy requirements; (iii) utilization of a cheap and abundant energy source; (iv) reduction in sludge formation; and (v) production of algal biomass for biofuel production (de-Bashan and Bashan 2010; Park et al. 2011). In addition, algae have the advantage of being easily grown, requiring only light and a carbon source (Tikoo et al. 1997). In another study, various species of fresh-water algae were used for the biodegradation of phenol and catechol. A remarkable observation was the carbon dioxide production when biodegradation was taking place (Ellis 1977). Since then, a large number of studies have been carried out demonstrating that microalgae are capable of biodegrading phenolic compounds. Regarding the mechanistic studies, the green microalga O. danika was found to biodegrade phenol through the meta-cleavage pathway (Semple and Cain 1996), while in Chlorella species, there is involvement of cytochrome P450 in the biodegradation of phenolic compounds (Thies et al. 1996). In addition, NADH-dependent reactions were found to take place during the biodegradation of these compounds by the marine diatom Thalassiosira sp. (Lovell et al. 2002).
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Pinto et al., used two green microalgae, Ankistrodesmus braunii and Scenedesmus quadricauda, in order to biodegrade phenols found in olive-oil mill wastewaters (Pinto et al. 2002). Chlorella and Scenedesmus species have been widely used for the biodegradation of phenolic compounds. A number of publications have demonstrated that Chlorella species are capable of biodegrading a variety of phenolic compounds, such as phenol, bisphenol-A, 4-nitrophenol, 4-chlorophenol, 2,4- dinitrophenol, and 2,4- dimethylphenol (Klekner and Kosaric 1992; Hirooka et al. 2003; Tsuji et al. 2003; Lima et al. 2004). In the abovementioned studies, the common characteristic was the presence of an alternative carbon source under illumination. Scenedesmus species have been shown to be capable of removing acylated phenols and bisphenol-A (Nakajima et al. 2007; Zhou et al. 2013). Although there are many studies regarding the biodegradation of phenolic compounds by microalgae, only few focus on the bioenergetic aspect of this procedure. Cultures of Scenedesmus obliquus treated with various compounds showed an increased growth in the presence of glucose as a carbon source, and, at the same time, exhibited higher levels of phenolic compound biodegradation as compared to treatments with inorganic carbon or a limited carbon source. This study proposed that the most important factor in the biodegradation of these compounds is the selection of the appropriate conditions (Papazi and Kotzabasis 2007). Two years later, a mechanistic dynamic energy budget model for aerobic degradation was proposed. This model suggests that biodegradation inhibition may occur in the presence of a growth-enhancing carbon source, such as glucose, because of a competition for oxygen, while at the same time its presence increases the specific growth rate (Lika and Papadakis 2009). Recently Chlamydomonas mexicana was found to be able to remove bisphenol-A, while increasing the concentration of this substance caused an increase of the carbohydrate levels in these cells as a response to stress effects (Ji et al. 2014). In the present study, the biodegradation of phenol by axenic cultures of Chlamydomonas reinhardtii was investigated. Various phenol concentrations were examined, ranging from 0.15 to 4.00 mM. The effect of various conditions, such as the presence or absence of an alternative carbon source in the culture medium, light intensity during growth, and oxygen availability, on phenol biodegradation was investigated. We also investigated the effects of phenol on the growth and photosynthetic activity of the microalgae, and how the sensitivity/tolerance of Chlamydomonas to different concentrations of phenol correlates with the biodegradation of this compound.
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Materials and methods Organism and growth conditions In this study, the CC-125 strain (wild type) of Chlamydomonas reinhardtii was used. Cultures of Chlamydomonas were grown heterotrophically in TAP medium (Harris 1989) at 25 °C, under a light intensity of 70–80 lE m-2 s-1 for five days. The cultures were grown in Erlenmeyer flasks shaken at a rate of 140 min-1. The cells were first collected by centrifugation at 1000 g for 3 min, resuspended into the new medium and, after following the same procedure twice, were finally distributed into tightly capped glass bottles, with septa (diameter 5 cm, height 9.5 cm), with 100 mL volume capacity. The final volume added was 50 mL. The experiments were performed in a temperature-controlled room (25 °C) under a light intensity of 50–60 lE m-2 s-1. Four different phenol concentrations were examined, 0.15 mM (14.1 mg/L), 0.50 mM (47.1 mg/L), 2.00 mM (188.2 mg/L), and 4.00 mM (376.4 mg/L). Phenol was dissolved in twice distilled water, and the solution was filtered through a 0.2lm syringe filter. The phenol stock solution’s concentration was 0.5 M. Apart from phenol, four different combinations of an exogenously supplied carbon source were tested, acetate (1 g/L) as an organic carbon source, carbon dioxide (20 %) as an inorganic carbon source, a combination of the two abovementioned carbon sources, and limitation of alternative carbon source (acetate and carbon dioxide). All experiments were repeated at least three times. All nutrient media and glassware were autoclaved at 120 °C for 20 min, in order to avoid contamination. Cultures were shaken at 140 min-1on a rotary shaker. Experiments were carried out in a laminar flow hood sterilized with ethanol and a UV lamp.
Determination of growth Cell growth was determined by measuring the packed cell volume (PCV) of the culture. The PCV of a cell suspension was determined by centrifugation at 1000 g for 5 min using hematocrit TPP tubes and expressed as lL PCV(mL culture)-1 (Senger and Brinkmann 1986). PCV was converted to cell dry weight according to the following equation that was determined experimentally: Dry Weight (mg/mL) = 0.340 PCV (lL/mL) - 0.020 (R2 = 0.991). Samples of 500lL were collected every 24 h with a sterile syringe. Sample collection was carried out in a laminar flow hood sterilized with ethanol and a UV lamp. From the growth measurements, the specific growth rate (l) was calculated for the first 48 h of culture according to the
equation l = ln[(X – X0)/(t – t0)], where X is the cell biomass at time t and X0 is the initial cell biomass at time t0. The observed growth yield (YX/S) was also calculated at the last day of the experiments (day 5) according to the equation YX/S = (X – X0)/(S0 - S), where X is the cell biomass at the last experimental day, X0 is the initial cell biomass, S0 is the initial phenol amount (mg), and S is the phenol amount (mg) left in the culture medium at the last experimental day (Basak et al. 2014). Fluorescence induction measurements The fluorescence induction measurements were carried out using the Handy Plant Efficiency Analyser PEA (Hansatech Instruments, Kings’ Lynn, Norfolk, UK). The parameters were measured according to the JIP-test, and included the maximal photosynthetic efficiency (Fv/Fmax), the antenna size per reaction center (ABS/RC), the dissipation energy per reaction center (DI0/RC), the density of active photosynthetic centers (RC/CS0), and the quantum yield of primary photochemistry (uP0 or TR0/ABS) (Strasser and Strasser 1995). This method is based on the measurement of a fast fluorescence transient with a 10 ls resolution for 1 s and allows the dynamic measurement of a photosynthetic sample at a given physiological state. Fluorescence was measured at 12-bit resolution and excited by six lightemitting diodes providing a saturating red (650 nm) light intensity of 3000 lmol m-2 s-1. Quantification of photosynthetic pigments For the quantification of photosynthetic pigments, 1 mL of culture was centrifuged at 1000 g for 5 min, and after supernatant disposal, the cell pellet was dissolved in 1 mL of 80 % acetone. Then the absorbance at specific wavelengths (470.0, 646.8 and 663.2 nm) was carried out by a Shimadzu UV-2700 UV–Vis spectrophotometer and formulae described previously in literature (Wellburn 1994). High-Performance Liquid Chromatography (HPLC) for the analysis of phenol For phenol analysis, culture samples were centrifuged for 3 min at 1000 g and 50 lL of the filtered supernatant were injected into HPLC. The analyses were performed using an HPLC system, equipped with a Gyncotec high-precision pump model 480, a Linear UV–Vis 200 UV detector, and a narrow-bore column (Grace Smart RP18, 250 mm l and 4.6 mm ID, 5 lm particle size); an isocratic HPLC method, using as a mobile phase methanol:water:acetic acid (50:49:1), and a flow rate of 1.0 mL min-1 (Lovell et al. 2002). Detection was carried out by measuring absorbance
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at 279 nm and quantification by the integration of known quantities of phenolic compounds.
Results and discussion High concentrations of phenol inhibit the growth of Chlamydomonas reinhardtii Kinetics of biomass production (Fig. 1) clearly showed that the carbon source used, in combination with the concentration of phenol used, exerted a strong influence on culture growth. More specifically, independently of the presence or absence of phenol, best growth was achieved in the presence of acetate. In the absence of an alternative carbon source (Limit C), no growth was observed. This result is expected because we have to
Fig. 1 Growth curves of Chlamydomonas reinhardtii CC-125 cultures under various carbon source treatments (CO2, Acetate, Acetate ? CO2, Limit C) under a light intensity of
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take into consideration two factors; (i) the organic carbon source provides the microalgae with facile energy reserves and (ii) CO2 was added in the culture medium only in the beginning of the experiment and thus growth stopped after it was consumed. An inhibition of growth was observed as the phenol concentration increased. As shown in Table 1, the specific growth rate (l) decreased from 29 to 42 % in the presence of 4.00 mM phenol. Under all conditions, even at low concentrations of phenol, growth higher than that of the control culture was not observed. This is shown more clearly in the Acetate treatment, where only the 0.15 mM phenol culture reaches the control biomass in the last experimental day. In CO2 treatment, the same happens for the 0.50 mM culture. The above results show that phenol acts as a growth inhibitor in Chlamydomonas cultures, especially at high concentrations.
50–60 lE m-2 s-1 in the presence of various concentrations of phenol. Initial cell concentration was 2.0 lL PCV/mL
Photosynth Res Table 1 Expression of the microalgal growth in terms of specific growth rate (l) and observed growth yield (YX/S) under various carbon source treatments (CO2, Acetate, Acetate ? CO2), under light intensity of 50–60 lE m-2 s-1, in the presence of different concentrations of phenol CO2 a
Acetate -1
l (days )
YbX/S
(mg cells/mg phenol)
-1
l (days )
Acetate ? CO2 YX/S (mg cells/mg phenol)
l (days-1)
YX/S (mg cells/mg phenol)
Control
0.46
–
0.65
–
0.66
–
0.15 mM
0.46
nd
0.60
4521.30
0.65
2260.65
0.50 mM 2.00 mM
0.46 0.28
nd nd
0.55 0.46
387.15 71.84
0.65 0.49
470.97 80.26
4.00 mM
0.28
38.28
0.46
19.87
0.38
17.77
a
Specific growth rate (l) was calculated for the first 48 h of culture according to the equation l = ln[(X – X0)/(t - t0)], where X is the cell biomass at time t (48 h) and X0 is the initial cell biomass at the beginning of the experiment (t0)
b
Observed growth yield (YX/S) was calculated at the last day of the experiments (day 5) according to the equation YX/S = (X - X0)/(S0 - S), where X is the cell biomass at the last experimental day, X0 is the initial cell biomass, S0 is the initial phenol amount (mg) and S is the phenol amount (mg) left in the culture medium at the last experimental day
Impact of phenol on the molecular structure and function of the photosynthetic apparatus To address the stress effects caused in the presence of phenol in the first 24 h, fluorescence induction measurements were carried out, and JIP test parameters were estimated (Table 2). From the results, it seems that in the presence of CO2 there are no stress effects. In Acetate cultures, gradually lower values of photosynthetic efficiency, density of the reaction centers, and primary photosynthetic yield are followed by an increase of the functional photosynthetic antenna and dissipation energy. A similar behavior was observed in the Limit C treatment as well. These changes in the parameters have been previously used to provide a typical evaluation of stress responses in the presence of high ozone levels (Navakoudis et al. 2003), thermal stress (Strasser and Strasser 1995), and the presence of substituted phenolic compounds (Papazi and Kotzabasis 2007). It is interesting to note that when acetate is the only alternative carbon source these effects are more intense, while at the same time the combination of organic and inorganic carbon sources in the culture medium results in milder stress effects, verifying the results observed in the presence of CO2 that were mentioned above. In all cases, there was a significant decrease in the density of fully active reaction centers (RC/CS0), regardless of whether the development of other stress responses was observed. It is important to note that when acetate was present, the maximum photosynthetic efficiency on the last experimental day returned to high levels, especially at 4.00 mM phenol. On the other hand in the absence of acetate, photosynthetic efficiency decreases significantly on the last experimental day. This fact underscores the significance of the organic carbon source as a facile energy reserve. In all cases, phenol treated cultures showed lower Fv/Fmax ratios
than the corresponding controls on the last experimental day. This result is also confirmed by the chlorophyll-a/b ratio (Table 3) in Chlamydomonas cells on the same day. The chlorophyll-a/b ratio appeared to be lower in phenol treated cultures and this can be attributed to a decrease in the density of the reaction centers and an increase of the functional photosynthetic antenna (chlorophyll-a is a pigment bound in the reaction center, while chlorophyll-b is mainly an antenna pigment).
Exogenously supplied carbon source, initial phenol concentration, and stress effects influence the levels of phenol biodegradation The biodegradation of phenol after five days of incubation is illustrated in Fig. 2. Figure 2a represents the absolute phenol biodegradation per culture, expressed in lmol, while Fig. 2b shows the phenol biodegradation expressed per packed cell volume (PCV). It is clear that in each separate condition, as the initial concentration of phenol increases so does its biodegradation. In the presence of inorganic carbon as the sole carbon source, biodegradation is observed only in the highest phenol concentration tested, and it is significantly lower compared to other treatments. So it can be assumed that the presence of an inorganic carbon source does not facilitate phenol biodegradation. This is supported by the fact that in the mixotrophic treatment (Acetate ? CO2), biodegradation is exactly the same as in the heterotrophic (Acetate) treatment. In terms of absolute phenol biodegradation, the presence of acetate seems to enhance this procedure. This observation is not actually true because in the presence of acetate, the cell biomass was at much higher levels compared to the other treatments. Since in Limit C conditions no growth was observed, it would be more appropriate to compare the
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Photosynth Res Table 2 JIP test photosynthetic parameters of Chlamydomonas reinhardtii cultures in the four experimental treatments of carbon source in the presence of different concentrations of phenol 24 h after the initialization of the experiments. Numbers in parentheses show the corresponding values in the 5th experimental day
Fv/Fmax
ABS/RC
DI0/RC
RC/CS0
TR0/ABS
Control
0.768 (0.635)
2.036
0.529
44.538
0.740
0.15 mM
0.777 (0.578)
1.891
0.455
42.002
0.759
0.50 mM
0.765 (0.577)
1.937
0.500
40.858
0.741
2.00 mM
0.766 (0.575)
2.181
0.572
37.977
0.738
4.00 mM
0.774 (0.568)
2.060
0.526
36.049
0.745
Control
0.773 (0.750)
1.855
0.437
53.901
0.747
0.15 mM
0.759 (0.738)
2.048
0.496
47.126
0.741
0.50 mM
0.734 (0.728)
2.163
0.572
43.814
0.704
2.00 mM
0.700 (0.714)
2.261
0.669
38.521
0.674
4.00 mM
0.536 (0.650)
3.109
1.365
30.643
0.511
Control
0.760 (0.765)
1.944
0.496
50.910
0.745
0.15 mM 0.50 mM
0.726 (0.758) 0.712 (0.753)
1.991 2.088
0.594 0.621
50.747 46.209
0.711 0.692
2.00 mM
0.719 (0.741)
1.958
0.599
42.468
0.686
4.00 mM
0.697 (0.741)
2.319
0.755
39.987
0.674
Control
0.681 (0.640)
2.138
0.698
30.715
0.663
0.15 mM
0.677 (0.552)
2.287
0.772
29.671
0.649
0.50 mM
0.653 (0.540)
2.877
1.246
26.484
0.590
2.00 mM
0.600 (0.506)
3.385
1.470
23.286
0.581
4.00 mM
0.580 (0.491)
3.489
1.533
24.802
0.554
CO2
Acetate
Acetate ? CO2
Limit C
Fv/Fmax photosynthetic efficiency (STDEV 0.005–0.027), ABS/RC functional antenna size (STDEV 0.044–0.309), DIo/RC dissipation energy per reaction center (STDEV 0.001–0.087), RC/CS0 density of active reactions centers (STDEV 0.181–3.551), TR0/ABS quantum yield of primary photochemistry (STDEV 0.001–0.043)
amounts of phenol degradation expressed in terms of PCV. In such a case, the highest biodegradation levels are observed in the absence of an alternative carbon source, especially at low concentrations. The importance of this fact is that in carbon limited conditions cells are under greater pressure to adapt their metabolism in order to either gain carbon reserves and/or to detoxify their microenvironment. Similar observations made by Papazi and Kotzabasis working on Scenedesmus cultures (Papazi and Kotzabasis 2007). They observed biodegradation only in limited carbon cultures, and thus, they concluded that phenol was used by the microalga as an alternative carbon source. This scenario changes in the case of 4.00 mM phenol. In acetate-treated cultures biodegradation per culture was significantly increased. In any case, the growth inhibition caused by phenol at high concentrations should not be underestimated. For this reason, the observed growth yield (YX/S) was calculated (Table 1). It is clear that phenol biodegradation slows down the cell growth. According to the Pirt theory (Pirt 1965), this can be attributed to
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maintenance, implicating that the extra substrate consumption is not used for growth purposes or in other words ‘‘the energy is used for functions other than the production of new cell material’’. Rusell and Cook (Russell and Cook 1995) proposed that since maintenance is a function that detracts from growth, the contribution of maintenance is more pronounced when the growth rate is low as it is observed for the specific growth rates in our data (Table 1). Liu et al. (1998) took into account the energy uncoupling coefficient and proposed that this parameter features reduction in the efficiency of converting energy into cellular biosynthesis under substrate-sufficient conditions. The substrate is consumed by microorganisms to form various intracellular metabolites and energy (ATP). The metabolites and energy are then used for biomass formation, maintenance and product formation. Meanwhile, the metabolites and energy may also be consumed by energy spilling or through futile cycles (Liu et al. 1998). In addition, a substrate-sufficient culture exhibits higher substrate consumption than a limiting one (Forrest 1969). In our case, these hypotheses are supported by the fact that in
Photosynth Res Table 3 Photosynthetic pigment content of Chlamydomonas reinhardtii cultures in the four experimental treatments of carbon source in the presence of various phenol concentrations in the 5th experimental day Ca
Cb
Cx?c
Ca/Cb
CO2 Control
4.10
1.69
1.54
2.43
0.15 mM
4.23
1.80
1.59
2.36
0.50 mM
4.30
1.82
1.72
2.36
2.00 mM
4.26
1.85
1.78
2.30
4.00 mM
5.09
2.54
1.92
2.00
Acetate Control
4.23
1.99
1.62
2.12
0.15 mM
4.07
1.92
1.60
2.12
0.50 mM
3.89
1.88
1.59
2.07
2.00 mM
5.25
2.67
2.21
1.97
4.00 mM
7.24
3.98
2.88
1.82
Control 0.15 mM
4.55 4.45
2.02 2.00
1.67 1.68
2.25 2.22
0.50 mM
3.86
1.84
1.46
2.10
2.00 mM
5.50
2.68
2.14
2.06
4.00 mM
5.34
2.80
1.99
1.90
Acetate ? CO2
Chlamydomonas cultures that lack an alternative carbon source, no growth was observed despite the high phenol biodegradation efficiency, so microalgal metabolism utilizes energy production for phenol degradation rather than for biomass production. Taking into account the biodegradation of phenol in acetate-treated cultures (Fig. 3a) as well as the Fv/Fmax parameter (Table 2) under the same conditions on the last experimental day, we can assume that the biodegradation of phenol present in cultures (in concentrations higher than 2.00 mM) takes place for detoxification purposes, as the respective acetate-sufficient cultures exhibit increased Fv/Fmax ratios after the effective removal of phenol from the culture medium. Although the same approach seems to be followed in the carbon limited cultures, the fact that a decrease in photosynthetic efficiency is observed demonstrates the importance of the presence of a high-energy source in the culture medium. Thus, the role of acetate should not be underestimated, because it is clear that it has a prominent role in allowing the microalgae to overcome the stress effects imposed by the high concentrations of phenol, independently of the cell growth. It is clear that in all cases, increased stress leads to increased biodegradation of phenol.
Limit C Control
6.11
2.79
2.26
2.19
0.15 mM
5.76
2.82
2.24
2.05
0.50 mM
4.89
2.39
1.94
2.05
2.00 mM
4.28
2.29
1.47
1.86
4.00 mM
4.01
2.51
1.56
1.60
Light intensity and oxygen availability contribute significantly to the biodegradation of phenol
Ca Chlorophyll-a (STDEV 0.02–0.50), Cb Chlorophyll-b (STDEV 0.01–0.28), Cx?c Total carotenoids (STDEV 0.01–0.20), Ca/Cb Chlorophyll-a over chlorophyll-b ratio
Since higher phenol biodegradation per cell volume unit was observed in acetate deprived cultures, we conducted further experiments on the Limit C treatment. In order to verify the results of increased phenol biodegradation in the limited carbon conditions, experiments in Sueoka’s high salt (HS) minimal medium (Sueoka 1960) were carried out.
Fig. 2 Biodegradation of phenol under various carbon source treatments (CO2, Acetate, Acetate ? CO2, Limit C) measured after 5 days of incubation time. Biodegradation is expressed as a Absolute
amount (lmol) of phenol biodegraded and b Amount biodegraded per unit of PCV [lmol of phenol (mL PCV)-1]. Cultures were incubated under a light intensity of 50–60 lE m-2 s-1
All pigment concentrations are expressed in mg pigment/mL PCV
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The results are presented in Fig. 3a. No significant difference is observed in phenol biodegradation compared to the acetate-limited TAP medium (phenol was the only carbon source present). It has been proposed that the biodegradation of phenolic compounds is a photoregulated process (Papazi and Kotzabasis 2007). For this reason, the same series of experiments were carried out under a higher light intensity (70–80 lE m-2 s-1) and in absolute darkness as well. As shown in Fig. 3b, a significant increase in the phenol biodegradation was observed at high initial phenol concentrations at higher illumination conditions, while a negligible biodegradation of phenol was observed in the dark at higher initial phenol concentrations. It is important to mention that at 0.15 mM of phenol a higher biodegradation was observed in the dark; this phenomenon though changes
as the phenol concentration increases. We could attribute this to the fact that higher light intensities (that is more energy provided for photosynthetic activity) facilitate phenol biodegradation under increased stress conditions. Most probably, microalgae achieve a higher phenol biodegradation by using light, a cheap and abundant energy source. In the dark, Chlamydomonas has to use phenol as an alternative carbon source as it has no energy reserves provided by photosynthesis. It is easier for the cells to perform this function at low concentrations of the substance but not at high concentrations that are inhibitory. It has been proposed that biodegradation of phenol by microalgae is an aerobic process with high oxygen demands (Lika and Papadakis 2009) as the enzymes implicated (e.g., phenol monooxygenase, catechol 2,3dioxygenase, etc.) use oxygen as a substrate (Semple and
Fig. 3 Biodegradation of phenol expressed in lmol of phenol (mL PCV)-1 at the Limit C conditions at different experimental factors. a Comparison between TAP medium (dark gray) and HS (minimal) medium (light gray) under a light intensity of 50–60 lE m-2 s-1. b Comparison between absolute dark (black square) and increased light intensity from 50 to 60 lE m-2 s-1 (light gray circle) to 70–80 lE m-2 s-1 (dark gray triangle). c Comparison between air
(dark gray) and nitrogen atmosphere (light gray) at the start of the experiment. d Comparison among air (dark gray), nitrogen atmosphere (light gray), and oxygen atmosphere (gray) at the start of the experiment, under a light intensity of 50–60 lE m-2 s-1. All experiments in b, c and d were carried out in acetate-limited TAP medium
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Cain 1996). When we flushed the bottles with nitrogen in order to replace the air and repeated the experiments under light conditions (Fig. 3c), we observed no significant differences in phenol biodegradation. This result was expected because upon illumination oxygen is produced photosynthetically. On the other hand, when the same experiment was carried out in complete darkness (Fig. 3d), no biodegradation of phenol was observed even at low concentrations. In order to confirm this observation, the same experiment was performed, but this time the bottles were flushed with oxygen; a significant biodegradation of phenol at high concentrations was observed in the dark, and this fact leads to the conclusion that biodegradation of phenol by Chlamydomonas reinhardtii is an oxygen-dependent process. It is important to note that under all conditions parallel control experiments were carried out in order to check whether any abiotic removal of phenol (due to photooxidative chemical reactions) was taking place; no decrease in the concentration of phenol was observed in these control experiments (data not shown). The biodegradation of phenol is a dynamic bioenergetic process that strongly depends on the selection of the appropriate growth conditions that assist microalgae to activate appropriate mechanisms in order to metabolize the compound. This work is not one more routine study about another microalgal strain capable of biodegrading phenol. The present study emphasizes mostly on the bioenergetic aspects of this issue, thus providing information on the mechanism a photosynthetic microorganism under stress conditions is using in order to regulate its energy balance to biodegrade phenol, while at the same time investigating other parameters that regulate phenol biodegradation.
Conclusions The present study has demonstrated that the biodegradation of phenol by Chlamydomonas is a dynamic bioenergetic process, affected by changes in culture conditions. It seems that microalgae biodegrade high amounts of phenol, although this process is energy demanding, probably for detoxification purposes. Biodegradation of phenol by Chlamydomonas is an aerobic and photoregulated process. This is the first time that Chlamydomonas reinhardtii has been used for bioremediation purposes. The present study has provides information regarding the bioenergetic aspects of the phenol biodegradation process that will facilitate our efforts to further elucidate the biodegradation mechanism. Acknowledgments The work was funded by the Greek General Secretariat of Research and Technology (Greece-Romania bilateral program 2011–2012).
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