Open access peer-reviewed chapter

Selection of Promising Bacterial Strains as Potential Tools for the Bioremediation of Olive Mill Wastewater

Written By

Daniela Campaniello, Antonio Bevilacqua, Milena Sinigaglia and Maria Rosaria Corbo

Submitted: September 30th, 2014 Reviewed: May 25th, 2015 Published: October 14th, 2015

DOI: 10.5772/60896

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The main objective of this paper was the selection of promising bacterial strains to be used as potential tools to remove phenols in olive mill wastewater (OMW) or in other food wastes. Therefore, 12 OMW samples were analyzed and 119 isolates were collected. After a preliminary screening on a medium containing vanillic and cinnamic acids, three isolates were selected to evaluate their viability in presence of different compounds (cinnamic, vanillic and caffeic acids, rutin, tyrosol and oleuropein) and a possible bioremediation effect. The isolates generally survived with phenols added and exerted a significant bioremediation activity in some samples (reduction of phenols by 20%). The last step was focused on the evaluation of the combined effects of pH, cinnamic and vanillic acids on the viability of a selected isolate (13M); the combination of the acids exerted a strong effect on the target, but alkaline pH played a protective role.


  • Bioremediation
  • phenol degradation
  • phenolic compounds
  • olive mill wastewater

1. Introduction

Olive oil production is one of the most important food sectors in the Mediterranean area as olive processing is considered a traditional industry for its countries since ancient times [1]. It is mainly produced in Spain (36% of the global production), Italy (24%), Greece (17%) followed by Portugal, France, Cyprus, Croatia, Turkey, Syria, and Tunisia. New producers are Argentina, Australia, and South Africa.

Olives are processed through two methods: pressing (discontinuous process) and centrifuging (continuous process, two/three phase centrifugation). The main inconvenience of these methods is the production of a polluting by-product that is a dark effluent known as olive mill wastewater (OMW) [2].

The environmental impact of olive oil production is strong due to the use of large quantities of water and the production of OMW: 1,000 kg of olives produce 0.5 m3–1.5 m3 of OMWs [1]. They are by-products generally considered undesirable but inevitable for every olive processing.

As defined in reference [3], OMW is “a stable emulsion constituted by vegetation waters of the olives, water from the processing, olive pulp and oil.” It is characterized by a particular color (intensive violet-dark brown up to black color), odor (strong specific olive oil smell), high degree of organic pollution (expressed as biological and chemical oxygen demand (BOD and COD) values), acidic pH, high electrical conductivity, high content of polyphenols, high buffer capacity, and high content of solid matter.

OMWs are generally composed of water (83%–96%) and organic fraction (3.5%–15%) composed of 1%–8% carbohydrates, 0.5%–2.4% nitrogen compounds, 0.5%–1.5% organic acids, 0.02%–1% of fatty acids such as propionic, butyric, etc., and 1%–1.5% of phenolic compounds consisting of a hydroxyl group (-OH) bound directly to an aromatic hydrocarbon group and pectins [4].

Concerning phenols, they comprise low molecular weight compounds and polyphenols. Low molecular weight compounds are represented by caffeic, cinnamic, 2,6-dihydroxybenzoic, p-hydroxybenzoic, syringic, 3,4,5-trimethoxybenzoic, vanillic, and veratric acids; they have phytotoxic effects and antibacterial activity. Polymeric phenols (lignins, tannins, etc.) cause the typical brownish-black color of OMW and are the most recalcitrant fraction of this effluent.

The quantitative and qualitative composition of OMWs are variable due to climatic conditions, variety, ripeness of olives, and extraction processes; generally, they are produced in high quantities in a short time, thus their disposal represents an important problem.

As OMWs are rich in nutrients they could be used to remediate arid or semi-arid regions but their phytotoxicy affect plant growth [5]. OMWs have the highest polluting rate within the food industry due to the fact that they are recalcitrant to traditional biodegradative methods. The reduction of COD and BOD values represent an important goal for many industries but the high content in phenols complicate waste management; they exert an antimicrobial activity towards wastewater microflora thus biodegradation is slowed [6].

For these reasons phenols are considered as undesirable compounds; thus, physical, chemical, or biological treatments are used to reduce their pollutant load.

Waste remediation has been traditionally performed through some expensive methods (incineration, pyrolysis, landfill, etc.). In recent years, the increasing trends towards green economy and friendly approaches for the environment are the background to design alternative ways. According to this point of view, numerous researchers proposed bioremediation, defined as “the process whereby organic wastes are biologically degraded under controlled conditions to an innocuous state or to level below the limits established by regulatory authorities” [7]. According to the Environmental Protection Agency (EPA), bioremediation is a “treatment that use naturally occurring organisms to break down hazardous substances into less toxic or nontoxic substances.”

Thanks to their ubiquity and metabolic pathways (aerobic, anaerobic fermentation, and co-metabolism) microorganisms are able to degrade and utilize various toxic compounds as energy source. Generally, the aerobic biodegradation has a higher efficiency than anaerobic processes and it is widely used. Nevertheless, in many cases, aerobic and anaerobic processes can also be used in series to reduce the complexity or the toxicity of the contaminants.

Numerous bacteria such as Bacillus pumilus[8], Pediococcus pentosaceus[9], Lactobacillus plantarum[10], Arthrobactersp. [11], Azotobacter vinelandii[12–14], Azotobacter chroococcum[15], Pseudomonas putida, and Ralstoniasp. [16, 17] were able to degrade and/or remove phenols from OMW.

Yeasts and molds are also able to degrade phenols, namely, Candida tropicalis, Candida cylindraceaand Yarrovia lypolitica[18, 19, 3, 20, 4], and white-rot fungi such as Phanerochaete chrysosporiumor the genus Pleurotus[2125]. In addition, Trametes versicolor, Funalia trogii, Lentinus edodes, Aspergillus niger, and Aspergillus terreushave been also mentioned as phenol-degrading organisms [26]. The main objective of this paper was the selection of promising bacterial strains to be used as potential tools for bioremediation; namely, after the isolation of some strains from OMWs, they were studied in relation to their ability to grow in a medium containing two secondary phenols. Then, a validation on a lab scale was performed.


2. Materials and methods

2.1. Isolation and phenotyping of potential phenol-degrading strains

Twelve different samples of OMW were analyzed. Aliquots of 100 ml of each OMW sample were mixed with 900 ml of sterile Ringer solution (0.25×; Oxoid, Milan, Italy) and shaken at 100 rpm for 30 min at room temperature. Then, this homogenate was serially diluted with a sterile saline solution (0.9% NaCl) and plated onto a Plate Count Agar (PCA; Oxoid) at 30°C for 48–72 h (for mesophilic bacteria and Bacillus) and Pseudomonas Agar Base + Pseudomonas C-F-C supplement (Oxoid) at 25°C for 72 h for Pseudomonadaceae. The analyses were performed in duplicate over two different batches. From each batch, some colonies were randomly selected from plates, and stored at 4°C on Tryptone Soya Agar (TSA) slants (Oxoid, Milan, Italy). Phenotyping of isolates was carried out through different tests (Gram, catalase activity, oxidase test, proteolytic activity, and oxido-fermentation).

2.2. Selection of potential phenol-degrading strains

The ability of the isolates to grow with phenols added was evaluated using Mineral Salt Medium (MSM), a synthetic medium containing K2HPO4 (1.6 g/l), KH2PO4 (0.4 g/l), NH4NO3 (0.5 g/l), MgSO4*7H2O (0.2 g/l), CaCl2 (0.025 g/l), FeCl2 (0.005 g/l), Agar (12 g/l), cinnamic (C9H8O2) or vanillic acids (C8H8O4) (0.5 or 1 g/l; Sigma-Aldrich, Milan, Italy). After streaking the isolates onto the surface of this modified medium, the plates were incubated at 30°C for mesophilic bacteria and Bacillusand at 25°C for Pseudomonadaceaefor 72 h. MSM without phenolic compounds was used as control. For a second assay, the isolates were preliminary grown in MSM broth added of 0.025 g/l and 0.05 g/l of cinnamic and vanillic acids and incubated for 24 h; thereafter they were streaked onto the surface of MSM with phenols, as reported above.

2.3. Effect of phenolic compounds on the viable cell count

This step was performed on three selected strains (6P, 13M, and 44M); they were grown in TS broth incubated at 25°C (strain 6P) or at 30°C (strains 13M and 44M) for 48 h. Each culture was centrifuged at 4,000 rpm for 10 min and the pellet was re-suspended in sterile saline solution (0.9% NaCl); ca. 6–7 log cfu/ml were inoculated in MSM medium (1, 2, and 3 g/l), added with phenols (cinnamic acid; vanillic acid; caffeic acid-C9H9O4; rutin hydrate-C27H30O16 x H2O; tyrosol-C8H10O2; oleuropein-C25H32O13; phenols were purchased from Sigma-Aldrich). MSM without phenols was used as control.

The samples were stored at 25°C–30°C for 33 days and periodically analyzed to evaluate the viable count on TSA and the content of phenols through the Folin-Ciocalteau method [27]. The analyses were performed in duplicate and the results analyzed through one-way Analysis of Variance (one-way ANOVA), using Tukey’s test as the post-hoc comparison test, or t-student test for paired comparisons. The statistical analysis was performed using the software Statistica for Windows version 10.0 (Statsoft, Tulsa, OK, USA).

2.4. Combined effects of pH, cinnamic, and vanillic acids on the viable count of the strain 13M

The strain 13M was used as target; it was inoculated into MSM broth to 6–7 log cfu/ml. The amounts of cinnamic and vanillic acids and pH varied according to a 2k experimental (Table 1). The samples were stored at 30°C and periodically analyzed to evaluate the viable count and phenol content (up to 12 days).

CombinationspHCinnamic acid (g/l)Vanillic acid (g/l)

Table 1.

Combinations of 2k design.

The analyses were performed in duplicate and the results of viable count analyzed through a multiple regression approach by using the option DoE/2k design of the software Statistica for Windows.


3. Results

3.1. Isolation and screening on MSM

The viable count of mesophilic and spore-forming bacteria and Pseudomonaswas fairly high (7–8 log cfu/ml) (Table 2); thus, we selected 119 isolates (46 labeled as mesophilic bacteria, 44 and 29 belonging to Bacillusand Pseudomonasgenera, respectively).


Table 2.

Viable count (log cfu/g) of mesophilic bacteria (M), Bacillus(B), and Pseudomonas(P) in OMW samples. Data are the average (n=2).

The isolates were streaked on MSM with cinnamic or vanillic acids; Figure 1 shows the results for vanillic acid. Namely, 12 isolates were able to grow in presence of 0.5 g/l of this compound (Figure 1a) and 9 with 1g/l (Figure 1b). None of the isolates grew with cinnamic acid.

Figure 1.

Screening of the isolates on MSM with vanillic acid (0.5 g/l and 1.0 g/l). The numbers on the pictures indicate if the targets are able or not able to grow.

The screening was also performed after isolate growing in MSM broth with low amounts of phenols; this step could be referred as an induction phase, aimed at inducing the resistance to phenols. Figure 2 shows the results with 1 g/l of vanillic acid. There were 32 out of 119 isolates that acquired the ability to grow in MSM with vanillic acid; at the lowest concentration (0.5 g/l) 49 strains were able to grow. The same protocol was also used for cinnamic acid, but only a single isolate was able to grow after induction both at 0.5 g/l and 1 g/l (the isolate 26M).

Figure 2.

Screening of the isolates on MSM with vanillic acid (0.5 g/l and 1.0 g/l) after the induction. The numbers on the pictures indicate if the targets are able or not able to grow.

3.2. Viability and phenol reduction by some selected isolates

Three isolates (6P, 13M, and 44M; see Table 3 for their phenotypic traits) were selected and used as targets to assess their viability in the presence of various phenolic compounds (caffeic, cinnamic and vanillic acids, oleuropein, and rutin and tyrosol) at different concentrations (1 g/l, 2 g/l, and 3 g/l); moreover, we focused also on microorganism ability to reduce phenol content. Figure 3 reports the viability of the isolate 6P in the presence of caffeic acid; the initial cell number was 7 log cfu/ml. Then, it underwent a strong reduction within 5 days (ca. 1.5 log cfu/ml at 1 g/l and 2 log cfu/ ml at 3 g/l); in the last days of storage we found a tailing effect, with a residual cell count of 5 log cfu/ml. Similar results were found in the presence of cinnamic acid, tyrosol, rutin, and oleuropein (data not shown).

Vanillic acid at 2 g/l reduced the viable count by 3 log cfu/ml in 5 days with a final tailing effect and a residual cell count of 3–4 log cfu/ml. The lowest concentration (1 g/l) resulted in a slower death kinetic, with a similar residual viable count (Figure 4).

Proteolitic activity-++

Table 3.

Phenotyping of the isolates selected for the second step of the research. F, metabolism under aerobic and anaerobic conditions.

Figure 3.

Viability of the isolate 6P in MSM+caffeic acid (mean values ± standard deviation). *, viable count in MSM+caffeic acid are significantly different from control.

Figure 4.

Viability of the isolate 6P in MSM+vanillic acid (mean values ± standard deviation). Letters indicate significant differences (one-way ANOVA and Tukey’s test; P<0.05).

Phenols did not affect the viability of the strain 44M and the viable count was at 6–7 log/ml for the entire running time (data not shown). Some interesting results were found for the isolate 13M. Tyrosol at 3 g/l reduced the viable count by 3 log cfu/ml within 5 days, thereafter cell number increased up to 6–7 log cfu/ml (Figure 5); this trend could be the result of a kind of induction and adaptive evolution with phenols added. Caffeic acid at 1 g/l and 2 g/l caused a slight viability loss (1–2 log cfu/ml), while cell number was below the detection limit after 25 days at the highest concentration (3 g/l), thus suggesting a possible dose-dependent bactericidal effect (Figure 6). The other phenols did not affect the viable count (data not shown).

Figure 5.

Viability of the isolate 13M in MSM+tyrosol (mean values ± standard deviation). *, significantly different from control (t-student test, P<0.05).

Figure 6.

Viability of the isolate 13M in MSM+caffeic acid (mean values ± standard deviation).

We focused also on phenol content; Figure 7 shows the removal of vanillic acid by the isolate 13M and 44M (the initial content of the compound was 1 g/l). Both the strains were able to reduce its concentration in the broth, although the isolate 13M showed higher removal efficiency (ca. 18% after 33 days). The isolate 44M was also able to reduce cinnamic and caffeic acids by 24%–27% after 33 days (Figure 8). Removal was always found at 1g/l of phenols; higher initial amounts completely depleted bioremediation, probably due to a possible saturating action on the membrane.

Figure 7.

Removal of vanillic acid (initial content, 1 g/l) in MSM inoculated with the isolates 13M and 44 M; data are reported as absorbance fall. Mean values ± standard deviation.

Figure 8.

Removal of cinnamic and caffeic acids (initial content, 1 g/l) in MSM inoculated with the isolate 44 M; data are reported as absorbance fall. Mean values ± standard deviation.

3.3. Combined effects of cinnamic and vanillic acids and pH on cell count of the isolate 13M

This phase focused on the evaluation of pH, cinnamic and vanillic acids (combined through a 2k experimental design) on the viability of the isolate 13M. Figure 9 shows the evolution of cell count in some selected combinations of the design; when MSM medium was adjusted to pH 7.0 in presence of 2 g/l of the phenols (combination D) the viable count (7 log cfu/ml) was drastically reduced to ca. 4.73 log cfu/ml. On the other hand, an alkaline pH played a protective role, as the viable count was not reduced both in absence and with phenol added (combinations E and H).

Figure 9.

Viable count of the isolate 13M in some selected combinations of 2k design (seeTable 1).

Finally, Figure 10 shows the increase/decrease of cell count after 12 days; a positive value indicates a significant increase in the viable count, while a negative value indicates a death kinetic. These values were used as input data to run a multiple regression procedure and pinpoint the weight of each factor of the design; the results of this statistical analysis are listed in Table 4. The individual effects of phenols were not significant, while their interactive term played a negative role, i.e., it was the leading factor of death kinetic; on the other hand, the statistical analysis pinpointed a positive mathematical effect of pH, thus confirming its protective role toward viable count.

TermsStatistical effect
Cinnamic acidns
Vanillic acidns
pH/Cinnamic acidns
pH/Vanillic acidns
Cinnamic acid/vanillic acid-3.06
R2 ad0.776

Table 4.

Standardized effects of vanillic and cinnamic acids and pH on the reduction of the viable count of the isolate 13M after 12 days in MSM. Ns, not significant. R2ad, determination coefficient corrected for multiple regression.

Figure 10.

Increase/decrease of the viable count of the isolate 13M in MSM with phenols after 12 days. Mean values ± standard deviation (for the combinations seeTable 1).


4. Discussion

The disposal of wastewaters represents a major problem. Namely, OMWs have the highest polluting rate within the food industry, especially for the high concentration of phenolic compounds. An increased interest in environmental issues has favored the introduction of new technologies as alternative ways to traditional methods. A promising approach is represented by bioremediation, which reduces the pollution load of various by-products using the phenol-degrading ability of some microorganisms. In a previous research, we selected some promising yeasts (Pichia holstiiand Candida boidinii) able to reduce phenol content in OMW [26, 28]. Hereby, we evaluated bacterial ability to grow in a phenol-enriched medium; after isolation and phenotyping, we preliminary identified the most promising strains as Bacillusand Pseudomonasspp. These bacteria are normal constituents of wastewater microflora [29, 30]. After the screening with vanillic and cinnamic acids (chosen as representative of the most important phenolic moieties: coumaric and hydroxybenzoic compounds), we focused on viability in the presence of these compounds, as well as in a caffeic acid, rutin, oleuropein, and tyrosol enriched medium. Caffeic acid and rutin are representative of secondary phenolic compounds, while oleuropein is a high molecular weight phenolic glycoside responsible for the bitter taste in olive fruits. Oleuropein is present in olive mill wastewater as a result of debittering treatments and the extraction process of table and olive oil, respectively. Tyrosol is a product of hydrolysis of oleuropein. We used higher concentrations (1 g/l, 2 g/l, and 3 g/l) than those generally present in OMWs to evaluate bacterial viability in extreme conditions.

In some samples, phenols exerted a bactericidal action, due probably to their ability to form hydrogen bonds with proteins and/or enter cells. The resistant microorganisms generally convert phenols into carbolic acids; these latter compounds are transported through a trans-membrane shuttle system into the cytoplasm. Carbolic acid is converted to catechol, and after three enzymatic steps, oxaloacetate is formed; the final step is the conversion of oxalacetate to acetaldehyde and pyruvate. The enzymes involved are oxygenase, hydroxylase, peroxidase, tyrosinase, and peroxidase [31, 32]. These products, finally, follow the main metabolic pathway up to their complete mineralization by mitochondrial chain [33, 34]. The oxalacetate can also be used in other cellular activities. The isolates 13M, 44M, and 6P strains were catalase positive and this trait is an important requirement as it is related to phenol degradation [28].

The isolates showed a good metabolic capacity towards simple phenolic compounds, as they were generally able to survive, with some exceptions to this generalized statement. In addition, some isolates (e.g., 13M) significantly reduced the concentration of some compounds in the broth and these are desired traits to select a promising microorganism acting as a bioremediation tool [35].

Concerning the second step of the research (combined effects of phenols and pH), the most important result was the effect of alkaline pH, as it seemed to exert a protective role on cell viability, thus we could suggest that phenolic metabolism at basic pHs is favored because the enzymes might have an optimal pH of 9. In these conditions a high presence of hydroxyl ions is ensured and it is very important as they represent a fundamental substrate used in the first step of the catabolic pathway of phenol to obtain catechol. These assumptions, however, require a confirmation. Finally, the protective effect of alkaline pH suggests the potential use of these isolates for the bioremediation of alkaline washing water of table olives.


5. Conclusions

Bioremediation could be considered as the promising solution for numerous food industry wastes, and to date, several works are in progress to isolate new phenol-degrading strains. This work concurs to confirm the importance of microorganisms to degrade pollutants; we selected some promising bacterial isolates, showing some desired traits in lab media. Further investigations are required to improve our work, i.e., evaluation of waste, evaluation of the effect on BOD and COD, and a focus on the role of alkaline pH on the removal.



This paper was supported by the Italian Ministry of Education, University and Research through the grant ECO_P4 Promotion of ECO-friendly processes for the enhancement of quality of apulian food productions (P.O.N. RICERCA E COMPETITIVITA' 2007-2013 per le Regioni della Convergenza - Avviso n. 713/Ric. del 29/10/2010 - Titolo II - "Sviluppo/Potenziamento di DAT e di LPP").


  1. 1. La Cara F, Ionata E, Del Monaco G, Marcolongo L, Gonçalves MR, Marques IP. Olive Mill Wastewater Anaerobically Digested: Phenolic Compounds with Antiradical Activity. Chemical Engineering Transactions. 2012; 27 325–330.
  2. 2. Dias A, Bezerra RM, Periera AN. Activity and Elution Profile of Laccase During Biological Decolorization and Dephenolization of Olive Mill Wastewater. Bioresource Technology 2004; 92 (1) 7–13.
  3. 3. Lanciotti R, Gianotti A, Baldi D, Angrisani R, Suzzi G, Mastrocola D, Guerzoni ME. Use ofYarrowia lipolyticaStrains for the Treatment of Olive Mill Wastewaters. Bioresource Technology 2005; 96 (3) 317–322.
  4. 4. Martinez-Garcia G, Johnson AC, Bachmann RT, Williams CJ, Burgoyne A, Edyvean RGJ. Anaerobic Treatment of Olive Mill Wastewater and Piggery Effluents Fermented withCandida tropicalis. Journal of Hazardous Materials 2009; 164 (1–2) 1398–1405.
  5. 5. Paraskeva P, Diamadopoulos E. Technologies for Olive Mill Wastewater (OMW) Treatment: A Review. Journal of Chemical Technology and Biotechnology 2006; 81 (9) 1475–1485.
  6. 6. Bevilacqua A, Augello S, De Stefano F, Campaniello D, Sinigaglia M, Corbo MR. Bioremediation of Wastes of Table Olives and Olive Oil: Environment-friendly Approaches. Industrie Alimentari 2014; 547 5–11.
  7. 7. Mueller J.G., Cerniglia C.E., Pritchard P.H. Bioremediation of Environments Contaminated by Polycyclic Aromatic Hydrocarbons. In: Crawford R.L., Crawford D.L. (eds.) Bioremediation: Principles and Applications, Cambridge: Cambridge University Press 1996; 125–194.
  8. 8. Ramos-Cormenzana A, Juarez-Jimenez B, Garcia-Pareja MP. Antimicrobial Activity of Olive Mill Wastewaters (Alpechin) and Biotransformed Olive Oil Mill Wastewater. International Biodeterioration and Biodegradation 1996; 38 (3–4) 283–290.
  9. 9. Ben Othman N, Ayed L, Assas N, Kachouri F, Hammami M, Hamdi M. Ecological Removal of Recalcitrant Phenolic Compounds of Treated Olive Mill Wastewater byPediococcus pentosaceus.Bioresource Technology 2008; 99 (8) 2996–3001.
  10. 10. Ayed L, Hamdi M. Fermentative Decolorization of Olive Mill Wastewater byLactobacillus plantarum.Process Biochemistry 2003; 39 (1) 59–65.
  11. 11. Knupp G, Rucker G, Ramos-Cormenzana A, Hoyos SEG, Neugebauer M, Ossenkop T. Problems of Identifying Phenolic Compounds During the Microbial Degradation of Olive Mill Wastewater. International Biodeterioration and Biodegradation 1996; 38 (3–4) 277–282.
  12. 12. Constantinos E, Papadopoulou K, Kotsou M, Mari I, Constantinos B. Adaptation and Population Dynamics ofAzotobacter vinelandiiDuring Aerobic Biological Treatment of Olive Mill Wastewater. FEMS Microbiology Ecology 1999; 30 (4) 301–311.
  13. 13. Balis C, Chatzipavlidis J, Flouri F. Olive Mill Waste as a Substitute for Nitrogen Fixation.International Biodeterioration and Biodegradation 1996; 38 (3–4) 169–178.
  14. 14. Piperidou CI, Chaidou CI, Stalikas D, Soulti K, Pilidis GA. Balis C. Bioremediation of Olive Mill Wastewater: Chemical Alterations Induced byAzotobacter vinelandii.Journal of Agricultural and Food Chemistry 2000; 48 (5) 1941–1948.
  15. 15. Borja R, Martin A, Alonso V, Garcia I, Banks CJ. Influence of Different Aerobic Pretreatments on the Kinetics of Anaerobic Digestion of Olive Mill Wastewater. Water Research 1995; 29 (2) 489–495.
  16. 16. Di Gioia D, Bertin L, Fava F, Marchetti L. Biodegradation of Hydroxylated and Methoxylated Benzoic, Phenylacetic and Phenylpropenoic Acids Present in Olive Mill Wastewaters by Two Bacterial Strains.Research Microbiology 2001; 152 (1) 83–93.
  17. 17. Di Gioia D, Fava F, Bertin L, Marchetti L. Biodegradation of Synthetic and Naturally Occurring Mixtures of Mono-Cyclic Aromatic Compounds Present in Olive Mill Wastewaters by Two Aerobic Bacteria.Applied Microbiology and Biotechnology 2001; 55 (5) 619– 626.
  18. 18. D’Annibale A, Brozzoli V, Crognale S, Gallo AN, Federici F, Petruccioli M. Optimisation by Response Surface Methodology of Fungal Lipase Production on Olive Mill Wastewater.Journal of Chemical Technology and Biotechnology 2006; 81 (9) 1586–1593.
  19. 19. D’Annibale A, Sermanni GG, Federici F, Petruccioli M. Olive-Mill Wastewaters: A Promising Substrate for Microbial Lipase Production. Bioresource Technology 2006; 97 (15) 1828–1833.
  20. 20. Martinez-Garcia G, Johnson AC, Bachman RT, Williams CJ, Burgoyne A, Edyvean RGJ. Two-Stage Biological Treatment of Olive Mill Wastewater with Whey as Co-substrate. International Biodeterioration and Biodegradation 2007; 59 (4) 273–282.
  21. 21. Aloui F, Abid N, Roussos S, Sayadi S. Decolorization of Semisolid Olive Residues of “Alperujo” During the Solid State Fermentation byPhanerochaete chrysosporium, Trametes versicolor, Pycnoporus cinnabarinusandAspergillus niger. Biochemical Engineering Journal 2007; 35 (2) 120–125.
  22. 22. Garcìa Garcìa I, Jiménez Pena PR, Venceslada B, Martìn Martìn A, Martin Santos MA, Ramos Gomez E. Removal of Phenol Compounds from Olive Mill Wastewater UsingPhanerochaete chrysosporium, Aspergillus niger, Aspergillus terreusandGeotrichum candidum. Process Biochemistry 2000; 35 (8) 751–758.
  23. 23. Jaouani A, Sayadi S, Vanthournhout M, Penninckx MJ. Potent Fungi for Decolourisation of Olive Mill Wastewaters.Enzyme and Microbial Technology 2003; 33 (7) 802–809.
  24. 24. Kissi M, Mountadar M, Assobhei O, Gargiulo E, Palmieri G, Giardina P, Sannita G. Roles of Two White-rot Basidiomyces Fungi in Decolorisation and Detoxification of Olive Mill Wastewater.Applied Microbial Biotechnology 2001; 57 (1-2) 221–226.
  25. 25. Tsioulpas A, Dimou D, Iconomou D, Aggelis G. Phenolic Removal in Olive Oil Mill Wastewater by Strains ofPleurotusspp. in Respect to their Phenol Oxidase (Laccase) Activity. Bioresource Technology 2002; 84 (3) 251–257.
  26. 26. Bevilacqua A., Petruzzi L., Corbo M.R., Sinigaglia M. Bioremediation of Olive Mill Wastewater by Yeasts – A Review of the Criteria for the Selection of Promising Strains. In: Patil Y. B., Rao P. (eds.) Applied Bioremediation - Active and Passive Approaches. Rijeka: Intech; 2013; 53–68.
  27. 27. Bray H.G., Thorpe W.V. Analysis of phenolic compounds of interest in metabolism. In: Blick D. (ed.) Methods of Biochemical Analysis. New York: Interscience, 1954; 27–52.
  28. 28. Sinigaglia M, Di Benedetto N, Bevilacqua A, Corbo MR, Capece A, Romano P. Yeasts Isolated from Olive Mill Wastewaters from Southern Italy: Technological Characterization and Potential Use for Phenol Removal. Applied Microbial Biotechnology 2010; 87 (6) 2345–2354.
  29. 29. Ntougias S, Bourtzis K, Tsiamis G. The Microbiology of Olive Mill Wastes. BioMed Research International 2013; (accessed 13 February 2015).
  30. 30. Prakash B, Irfan M.Pseudomonas aeruginosais Present in Crude Oil Contaminated Sites of Barmer Region (India). Journal of Bioremediation & Biodegradation 2011; 2:5 (accessed 1 July 2014).
  31. 31. Nordlund I, Powlowski J, Shingler V. Complete Nucleotide Sequence and Polypeptide Analysis of Multicomponent Phenol Hydroxylase fromPseudomonassp. Strain CF600. Journal of Bacteriology 1990; 172 (12) 6826–6833.
  32. 32. Nair CI, Jayachandran K, Shashidhar S. Biodegradation of Phenol. African Journal of Biotechnology 2008; 7 (25) 4951–4958.
  33. 33. Zouari H, Moukha S, Labat M, Sayadi S. Cloning and Sequencing of a Phenol Hydroxylase Gene ofPseudomonas pseudoalcaligenesStrain MH1. Applied Biochemistry and Biotechnology 2002; 102–103 (1–6) 261–276.
  34. 34. Kirchner U, Westphal AH, Müller R, van Berkel WJ. Phenol Hydroxylase fromBacillus thermoglucosidasiusA7, a Two-protein Component Monooxygenase with a Dual Role for FAD. Journal of Biological Chemistry 2003; 278 (48) 47545–47553.
  35. 35. Murínová S, Dercová K. Response Mechanisms of Bacterial Degraders to Environmental Contaminants on the Level of Cell Walls and Cytoplasmic Membrane. International Journal of Microbiology 2014;

Written By

Daniela Campaniello, Antonio Bevilacqua, Milena Sinigaglia and Maria Rosaria Corbo

Submitted: September 30th, 2014 Reviewed: May 25th, 2015 Published: October 14th, 2015