Isolation and Characterization of Phenol-Degrading Psychrotolerant Yeasts

  • Natalia Filipowicz
  • Malwina Momotko
  • Grzegorz Boczkaj
  • Tomasz Pawlikowski
  • Marta Wanarska
  • Hubert Cieśliński
Open Access


In this study, the potential of selected psychrotolerant yeast strains for phenol biodegradation was studied. From 39 strains isolated from soil and water samples from Rucianka peat bog, three psychrotolerant yeast strains, A011, B021, and L012, showed the ability to degrade phenol. The result shows that all three yeast strains could degrade phenol at 500 and 750 mg l−1 concentration, whereas strains A011 and L012 could degrade phenol at 1000 mg l−1 concentration. The time needed for degradation of each phenol concentration was no longer than 2 days. Strains A011, B021, and L012 were identified based on 26S rDNA and ITS sequence analysis as belonging to species Candida subhashii, Candida oregonensis, and Schizoblastosporion starkeyi-henricii, respectively.


Biodegradation Phenol Psychrotolerant Yeast Peatland 

1 Introduction

Phenol is one of the most widely distributed environmental pollutants which can be found mostly in wastewaters and industrial effluents. World annual production of this chemical compound is around 10 million tons, while 3.3 million tons are produced in European countries, 2.4 million tons in the USA, and 0.2 million tons in Russia (Federal State Statistics 2011). By the end of 2016, global phenol production is expected to increase by 2.1 million tons (ICIS 2016). Main sources of this hazardous compound are chemical industry, coal conversion processes, crude oil refineries, textile industry, and production of pesticides (Aleksieva et al. 2002; Boczkaj et al. 2016a, b, 2014). Also, it is an ingredient in many customer products like mouthwashes or antiseptic lotions. Phenol is present in food, especially in smoked, grilled, and fried meat, and is also detected in honey and coffee. Moreover, phenol is also formed during the decomposition of biomass and is released into the atmosphere during volcanic eruptions (ATSDR 2015). It is also formed during chemical reactions which lead to cloud formation. Some levels of phenols are also produced during UV irradiation (sunlight) from amino acids in plant’s hemicellulose and tyrosine transformation in digestive track (Davidson 1996; Tsuruta et al. 2000).

The important fact, in respect of environmental aspects, is that phenol is stable and resistant to self-degradation in water solutions. Moreover, it tends to accumulate in soil and groundwater and is able to move from soil to water. Phenol can be also detected in rainwater, surface water, and drinking water.

Due to its toxicity, phenol could become life-threatening to living organisms if proper precautions are not taken. When discharged into water, it can become a danger for fish life at low concentration 5–25 mg l−1 (Chung et al. 2003; Kibret et al. 2000; Kumar et al. 2005). Phenol at the concentration level of 1 g/1 kg of body weight is lethal to humans and animals but individual tolerance for this chemical compound can be higher (ATSDR 2015). Exposure to phenol occurs through breathing, skin contact, or ingestion.

The water or soil treatment of this toxic compound is focused on physical and chemical methods such as solvent extraction, adsorption, ion exchange, and advanced oxidation processes (AOPs) (preferably by ozonation) (Aksu 2001; Aksu et al. 1999; Banat et al. 2000; Gimeno et al. 2005; Hameed and Rahman 2008; Lin and Juang 2009; Rengaraj et al. 2002; Roostaei and Tezel 2004). For this purpose, adsorption of phenol on activated carbon is one of the most common methods. However, the major problems associated with this method are: (i) limited mechanical strength of activated carbon, (ii) high regeneration cost of this sorbent, and (iii) intraparticle resistance (Aravindhan et al. 2009; Siva Kumar et al. 2009). Besides, in all mentioned above methods, there is a risk of formation of hazardous by-products (Wang et al. 2000). In comparison with these methods, biological treatment of phenol has two major advantages: (i) low cost of mineralization process (Zümriye et al. 1999; Bux et al. 1999) and (ii) rare possibility of toxic secondary metabolite production (Wang et al. 2000). Hence, biodegradation of phenol is an attractive alternative for recently used physical and chemical methods of phenol removing. It could be carried out using biofilters for gaseous pollutions (García-Peña et al. 2008; Qi et al. 2002; Woertz et al. 2001), bioreactors for industrial effluents treatment (Jarvinen et al. 1994; Juárez-Ramírez et al. 2001), bioremediation techniques for contaminated waters (Puhakka et al. 1995) and soil (Cordova-Rosa et al. 2009), and natural attenuation for both water and soil samples (Broholm and Arvin 2000; Holder et al. 1999). Removal of phenol from contaminated soil and industrial effluents by bioremediation techniques can be achieved through aerobic biodegradation, by microorganisms utilizing this toxic aromatic compound as a sole source of carbon and energy. Pseudomonas putida has been the most extensively studied bacterium for aromatic compound biodegradation (Morasch et al. 2002; Reardon et al. 2002; Wang et al. 2008; You et al. 2013). Alcaligenes eutrophus (NRRL B 75940) (Hill et al. 1996), Bacillus stearothermophilus BR219 (Gurujeyalakshmi and Oriel 1989), and some bacterial strains belonging to Acinetobacter spp., Citrobacter spp., and Shigella spp. (Kafilzadeh et al. 2010) are also capable of degrading phenol at low concentration. Among yeast, Candida tropicalis is known to degrade high initial phenol concentration (beyond 1700 and 2000 mg l−1, respectively) in a relatively short period of time (Jiang et al. 2008; Yan et al. 2005). Moreover, C. tropicalis strains, for, e.g., strain CC1 (Galíndez-Mayer et al. 2008) and strain LB-L20 (Juárez-Ramírez et al. 2001), were successfully used in bioreactors for the removal of phenol. Yeast strains belonging to Rhodotorula spp. and Trichosporon spp. are also capable of phenol degradation (Kurtz and Crow 1997; Sampaio 1999). Advantages of using yeasts in bioremediation include rapid growth and ability to resist unfavorable environmental conditions (Yu and Wen 2005). Cold-adapted yeasts, with higher metabolic versatility than bacteria to biodegradation of petroleum hydrocarbon fractions (Margesin et al. 2003), are excellent candidates in studies on phenol bioremediation for contaminated soil and water in cold climatic regions.

On the other hand, most of the studied hydrocarbon-degrading microorganisms are mesophiles (Boroujeni et al. 2014; Chakraborty et al. 2010). However, the temperature of groundwater and soils in most regions of the world is lower than optimum of the growth temperature for mesophilic microorganisms (Bergauer et al. 2005). Therefore, in recent years, the interest in research on the use of psychrotolerant microorganisms to biodegradation of monoaromatic compounds has increased. Kotturi et al. (1991) showed the ability to degrade phenol by psychrophilic P. putida strain Q5. Margesin et al. (2004) described cold-tolerant Arthrobacter sp. strain AGG31 with the ability to use as sole source of carbon: phenol, o-cresol, m-cresol, catechol, hydroquinone, and salicylate. But there is still little known about psychrotolerant yeasts and their potential in aromatic compound biodegradation (Bergauer et al. 2005). Bergauer et al. (2005) described the ability to degrade monoaromatic compounds by cold-tolerant and psychrophilic yeast strains. Phenol and 18-phenol-related compounds were tested. None of 32 isolated strains were able to utilize any of the highly volatile monoaromatic compounds. Compounds which were easily utilized were phenol (100% of strains), hydroquinone (91% of strains), resorcinol (88% of strains), and catechol (75% of strains). However, in comparison to analogous research on bacteria, all compounds were utilized for yeast strains by growth at low concentration (50–200 mg l−1, depending on the compound).

Another significant factor, which should be considered, is the origin of yeast strain isolation. Microorganisms with the ability to degradation of phenol and other monoaromatic compounds are usually isolated from hydrocarbon contaminated sites caused by human activity, such as petroleum hydrocarbon-contaminated alpine soils (Margesin et al. 2005), sewage (Sivasubramanian and Namasivayam 2015; Wang et al. 2012), activated sludge (Jiang et al. 2008; Yan et al. 2005), oil-shale mine and railway area (Bergauer et al. 2005), oil refinery wastewaters (Rocha et al. 2007), or wastewater from coking plants (Karimi et al. 2016). Some investigations about ability of microorganisms from natural sources to aromatic compounds degradation were made (Kotturi et al. 1991; Margesin et al. 2004; Wang et al. 2008). Presumably, screening of microorganisms with potential for phenol and its derivatives biodegradation from other natural sources can be an alternative solution. In this study, yeast strains were isolated from peatland. Peatlands are rich in polyphenols and phenol derivatives which are products of lignocellulosic biomass decomposition (Thormann et al. 2007). Furthermore, it has been shown that yeast species which are involved in the process of biomass decomposition are also able to degrade simple and complex sugars, selected sugar alcohols, organic acids, and glycosides (Thormann and Rice 2007). Hence, we assumed that the soil and water samples from peat bogs could be an attractive source of microorganisms, especially psychrotolerant yeast species which are capable of biodegrading this class of monoaromatic compounds. However, there is still a lack of information about studies on yeasts isolated from peatlands with the ability to degrade phenol and its derivatives (Thormann et al. 2007).

Therefore, the purpose of this study was to isolate and identify psychrotolerant yeast strains from water and soil samples derived from Rucianka peatland, which are capable of utilizing phenol (at a wide range of concentration 500–2000 mg l−1) as the sole source of carbon.

2 Materials and Methods

2.1 Sample Collection

Soil and water samples were collected from Rucianka raised bog, located in northern-east Poland (54° 15′ 22″ N, 19° 44′ 14″) on January 5 and on April 7, 2015. Rucianka bog covers an area of 240 ha. It was formed during Vistulian glaciation and was filled by a lake. The bog was formed on sandy deposition and consists of two materials: fen and sphagnum peat. The exploration of peat started before Second World War, and by now, 60% of bog area consists of peat mine (Pawłat 1996).

Sterile 50-ml conical centrifuge tubes (Sarstedt, AG & Co., Germany) were used to collect samples. Soil samples were collected from 30 locations and each sample was collected only once from each place from depth about 5 cm. Water samples were collected from 19 locations such as drainage ditches, pipelines, and puddles, and each sample was collected only once from each place from depth about 2 cm. All samples were transported straight after a collection to the laboratory in the ice box to maintain the temperature around 4–8 °C.

2.2 Isolation of Yeast Strains

One gram of each soil sample was transferred to sterile 50-ml conical centrifuge tube and mix vigorously with 9-ml sterile 0.9% saline solution for 10 s. After soil sedimentation, 100 μl of each sample was collected from 2 to 3 cm under water meniscus and spread with a sterile glass rod on YPD plates supplemented with chloramphenicol (stock solution = 34 mg ml−1) and ampicillin (stock solution = 100 mg ml−1). For water samples, 100 μl of each sample was transferred directly on YPD plate and spread with sterile glass rod. All plates were incubated at 18 °C for 6 days. After this period, colonies which differed from the other by morphological appearance (color, size, and shape of the colony) and microscopic examination were three times streak plating on YPD plates and incubated at 18 °C for 6 days to obtain a pure colony. Strain’s ability to grow on different temperature (4, 18, 30, and 37 °C) was tested.

Yeast extract peptone dextrose (YPD) contained (per liter) glucose 20 g, yeast extract 10 g, casein peptone 20 g, and bacteriological agar 20 g supplemented with 1 ml of each antibiotic: chloramphenicol (stock solution = 34 mg ml−1) and ampicillin (stock solution = 100 mg ml−1).

2.3 Assimilation Tests—Screening for Yeast Strain Capable of Utilizing Phenol

Screening for yeast strains capable of utilizing phenol was done using a slant culture method (Middelhoven et al. 1991) with some modifications. Phenol was a sole source of carbon for yeast growth and the agar slants inoculated with tested yeast strains were incubated at 18 °C. If tested yeast strain was capable of utilizing phenol, a growth of yeast appeared on a slant surface. The growth intensity of tested strain on a slant surface depended on the toxicity of phenol towards this strain and the efficiency of utilization of phenol by this strain. Criteria used for evaluation the slant culture method were (++) for an intensive growth of yeast strain without inhibition zone, (+) for a weak growth of yeast strain with small inhibition zone (5–7 mm), and (−) for no growth or weak growth of yeast strain with inhibition zone >10 mm.

2.4 Phenol Biodegradation

The first step of an experiment was a preparation of inoculation for each yeast isolate selected from assimilation test. This was achieved by inoculating 3-ml YPD medium with the single colony of examined yeast strain from actively growing YPD plate. For all yeast isolates, test tubes were incubated at 18 °C on the rotary shaker at the speed 170 rpm for 1 day. After this period, 80 μl of each yeast culture was transferred to 20 ml of fresh YPD medium in 200-ml Erlenmeyer flasks. Flasks were incubated at 18 °C on the rotary shaker at the speed 170 rpm for 3 days. Next, yeast cells were harvested and used as an inoculum. In all the experiments, 5% of subculture (1 ml) was inoculated into 20 ml of mineral salt medium (MSM) (Yan et al. 2005) in 200-ml Erlenmeyer flasks, supplemented with phenol at concentration 500, 750, 1000, 1500, and 2000 mg l−1, respectively. Mineral salt medium contained (per liter) (NH4)2SO4 0.4 g, K2HPO4 0.4 g, KH2PO4 0.2 g, NaCl 0.1 g, MgSO4 0.1 g, MnSO4·H2O 0.01 g, Fe2(SO4)3·nH2O 0.01 g, Na2MoO4·2H2O 0.01 g, and casein peptone 0.25 g, supplemented with 1 ml of each antibiotics: chloramphenicol (stock solution = 34 mg ml−1) and ampicillin (stock solution = 100 mg ml−1). The pH of the medium was adjusted to 6.0. At the start of incubation, OD of cultures were 0.2–0.3. All flasks were incubated at 18 °C on a rotary shaker at the speed 170 rpm for 5 days.

2.5 Analytical Methods

The optical density of growing yeast cultures was measured spectrophotometrically by measuring the absorbance at wavelength 600 nm (Feng et al. 2002) using 96-well plate VIS 96/F-PS (Eppendorf) and Eppendorf PlateReader AF2200. One hundred microliters of each yeast culture was placed on a well plate. For every well, five measurements were done according to parameters: type, xy; size, 3 × 3; and border, 1350 μm.

2.6 Process Control of Phenol Concentration During Biodegradation

To measure the concentration of phenol during biodegradation, simultaneously during optical density measurements, samples of suspended cultures were collected and centrifuged at 10,000 rpm for 10 min. One milliliter of each sample supernatant was placed in a 2-ml glass vial and 25 μl of 4-chlorophenol internal standard (IS) solution was added. The concentration of phenol in cell-free supernatant samples was determined by gas chromatography with a flame ionization detector (Clarus 580 GC, PerkinElmer, USA). An internal standard method was used. Detailed conditions of analysis are listed below.
  • Column: 60.0 m × 0.32 mm (ID) × 1.8 μm (DB-624) (Agilent, USA)

  • Carrier gas: nitrogen

  • Flow rate (a constant flow mode): 2.0 ml min−1.

  • The temperature program: initial temperature 60 °C, then a constant rate of 10 °C min−1 to final temperature 250 °C and then isothermally for 10 min

  • Temperature of injector and detector (FID): 275 °C

  • Sample injection: by autosampler, injection volume 2.0 ml, in splitless mode (for 1 min and after split mode 25:1)

2.7 Identification of Yeast Strains by Sequencing of the D1/D2 Region and ITS Region

For genomic DNA isolation from selected yeast strains, one colony of each strain was used to inoculate 4-ml sterile YPD medium. All yeast cultures were grown at 18 °C on a rotary shaker at the speed 170 rpm for 4 days. Afterward, each yeast culture was harvested, and the cell pellets were used for genomic DNA isolation with ExtractMe DNA yeast kit according to manufacturer’s instructions (Blirt S.A., Poland).

For yeast identification, two different genomic DNA regions were amplified by PCR, using DNA polymerase Pwo HyperNova (Blirt S.A., Poland). First was D1/D2 rDNA region which was amplified using the following primer pair: forward NL-1 (5′-GCATATCAATAAGCGGAGGAAAAG-3′) and reverse NL-4 (5′-GGTCCGTGTTTCAAGACGG-3′) (Hesham et al. 2006).

PCR was performed for 30 cycles including an initial denaturation at 95 °C for 60 s, subsequent denaturation at 95 °C for 30 s, annealing at 59 °C for 30 s, and extension at 72 °C for 45 s followed by final extension at 72 °C for 5 min and holding at 4 °C.

The second region was 5.8S-ITS rDNA region which was amplified using the following primer pair: forward ITS-1 (5′-TCCGTAGGTGAACCTGCGG-3′) and reverse ITS-4 (5′-TCCTCCGCTTATTGATATGC-3′) (Kurtzman and Robnett 1998).

PCR was performed for 30 cycles including an initial denaturation at 95 °C for 60 s, subsequent denaturation at 95 °C for 30 s, annealing at 52 °C for 60 s, and extension at 72 °C for 45 s followed by final extension at 72 °C for 5 min and holding at 4 °C.

Next, PCR amplification products were purified using ExtractMe DNA Clean-up Kit (Blirt S.A., Poland) according to manufacturer’s instructions. PCR products were sent to Genomed S.A., Poland and directly sequenced using 3730xl DNA Analyzer (Applied Biosystems, Life Technologies). PCR primers NL-1 and NL-4 of the D1/D2 26S rRNA gene and ITS1 and ITS4 of 5.8S-ITS region were used in the sequencing reactions to reading both DNA strands. The resulting DNA sequences were compared to the appropriate DNA sequences of the GenBank database using the basic local alignment search tool (BLAST) at the National Center for Biotechnology Information ( Phylogenetic trees for studied yeast strains were constructed with MEGA version 6.0 using a neighbor-joining algorithm and bootstrap analysis (1000) (Tamura et al. 2013).

3 Results and Discussion

3.1 Isolation of Yeast Strains

A total of 39 psychrotolerant yeasts were isolated from water and soil samples of Rucianka raised bog. Twenty-one isolates were obtained from water samples and 18 isolates were isolated from soil samples. For each isolate, a morphology of colony and cell morphology were studied. The pure cultures of all isolates were deposited in yeast strain collection of the Department of Molecular Biotechnology and Microbiology, Gdańsk University of Technology. All of the isolates were cryoconserved and kept under −80 °C.

3.2 Isolation of Phenol Utilizing Yeasts

After 1 week of slant incubation, growth density was estimated. For 6 from 39 examined yeast strains, intensive growth on the whole surface of slant was observed. Moreover, for these strains, there was no zone of growth inhibition on slants and turbidity of water-phenol solution present above slant surface was observed. For 23 yeast strains, growth on slants was classified as weak. Intensity was low and zone of growth inhibition was 5–7 mm. For 10 yeast strains, no growth or growth of few single yeast colonies was observed. For those strains, the zone of inhibition was greater than 10 mm (Table 1).
Table 1

Growth intensity on slants

Strain number

Collection number

Growth intensity

A023, A025, B012, F032, G021, J021, M033, P021, S012, S013

373, 374, 375, 353, 376, 358, 366, 367, 377, 378

A031, C021, C022, C031, D011, D014, D015, D021, F031, F034, I022, J022, K011, K012, L011, M031, M032, N021, P031, R011, R012, R013, S011

326, 328, 329, 379, 380, 381, 351, 350, 352, 382, 385, 359, 360, 361, 362, 364, 365, 383, 368, 369, 370, 371, 384


A011, B021, I021, L012, S014, A021

323, 324, 357, 363, 372, 325


Strain number: number assigned to pure culture after isolation from water or soil sample. Collection number: number assigned to strain (pure culture) deposited in the collection of Department of Molecular Biotechnology and Microbiology, Gdańsk University of Technology. Growth intensity: according to 1.2.3 in “Materials and Methods

Based on this method, six strains were selected for further studies. These strains were resistant to toxic properties of phenol, and presumably, these strains were able to use phenol as a sole carbon source.

4 Phenol Biodegradation—Results and Discussion

As mentioned above, most investigations are focused on phenol degradation by mesophilic microorganisms. Among them, bacteria are the most extensively examined. Bacteria are able to mineralize phenol at 100–1200 mg l−1 concentration (Polymenakou and Stephanou 2005) but the greater part of publications is focused on study of ability to use of phenol at 200 mg l−1 concentration. Also, several yeast strains were reported as capable of phenol biodegradation. The highest phenol concentration was degraded by Candida maltosa (1700 mg l−1 concentration) (Fialová et al. 2004) and C. tropicalis (up to 2000 mg l−1) (Yan et al. 2005). Therefore, for our selected psychrotolerant yeast strains, we decided to test their ability to degrade phenol at 500, 750, 1000, 1500, and 2000 mg l−1 concentration, respectively.

Figure 1a shows growth of selected six yeast strains during incubation time (76 h). For three of six examined strains, a significant increase of OD600 values and a decrease of residual phenol concentrations were observed. Strains A011, B021, and L012 showed an increase in the values of OD600 between 10 and 48 h of culture incubation (logarithmic phase of growth). After this period, OD600 values remained stable on the same level or decreased slightly with time. Strains B021 and L012 exhibited an adaptation phase which lasted for around 10 h when for strain A011 OD600 values started to increasing since of the beginning of its culture incubation. For strains A021 and I021, slightly increasing OD600 values were observed but it was not associated with decreasing of residual phenol concentration. For strain S014, no connection between OD600 values and residual phenol concentration was observed and it is likely that phenol is toxic for this microorganism. Strains A021, I021, and S014 were excluded from further studies.
Fig. 1

Growth curves for yeast strains in MSM medium supplemented with phenol at 500 mg l−1 concentration (a) and residual phenol concentration curves (b). Each strain has defined marker pointed out on figure

Analysis of residual phenol concentration chart (Fig. 1b) leads to conclusions that strains A011, B021, and L012 were able to reduce phenol from 500 mg l−1 concentration to below the detection limits in less than 30 h of the culture incubation. Reduction of phenol corresponded with increasing OD600 values. After utilization of phenol as a sole source of carbon, no significant changes of OD600 values were observed.

The next examined phenol concentration was 750 mg l−1. Only three strains, selected on 500 mg l−1 phenol concentration, were tested. Isolate A011 exhibited an adaptation phase which lasted around 24 h, whereas, for strains B021 and L012, OD600 values started to increase directly after starting of the culture incubation (Fig. 2a). Moreover, during the culture incubation time, the largest increase of OD600 values was observed for strain A011 (logarithmic growth phase between 24 and 48 h), whereas for strains B021 and L012, the increase of OD600 values was less significant and lower yeast biomasses were yielded. Highest OD600 values were reached around 48 h of culture cultivation for strains A011 and L012 and around 70 h for strain B021. After this period, OD600 values remained stable on the same level or decreased slightly.
Fig. 2

Growth curves for yeast strains in MSM medium supplemented with phenol at 750 mg l−1 concentration (a) and residual phenol concentration curves (b). Each strain has defined marker pointed out on figure

Chart of residual phenol concentration (Fig. 2b) reveals that all three strains were able to reduce phenol below detectable concentration. For isolate A011, reduction of phenol followed after 24 h of incubation and for strains B021 and L012 after around 48 h, respectively. Isolates B021 and L012 used phenol from the beginning of incubation, what corresponded with changes of OD600 values. For strain A011, the OD600 values increased faster and reached much higher levels, and along with these changes, phenol degradation was faster than for the other strains. In comparison to results obtained from previous chart of residual phenol concentration, for strain A011, incubation time needed for degradation of phenol at 500 and 750 mg l−1 was similar, whereas for strains B021 and L012, reduction of phenol at 750 mg l−1 concentration below detectable level took twice as much time as for 500 mg l−1 concentration.

The next tested phenol concentration was 1000 mg l−1. Only two strains, A011 and L012 (Fig. 3a), were able to grow in the presence of high concentration of phenol. For both strains, A011 and L012, OD600 values at the beginning and the end of the incubation, were comparable. For isolate A011, similarly to 750 mg l−1 concentration, adaptation phase lasted for 24 h whereas for isolate L012 OD600 values started to increase immediately after starting of incubation. Chart of residual phenol concentration (Fig. 3b) revealed that curves for each strain were similar. Both strains were able to reduce phenol concentration below the detectable level in around 48 h.
Fig. 3

Growth curve for yeast strains in MSM medium supplemented with phenol at 1000 mg l−1 concentration (a) and residual phenol concentration curves (b). Each strain has defined marker pointed out on figure

To sum up, the analysis of Figs. 1 and 2 leads to main conclusions. Three psychrotolerant yeast strains A011, B021, and L012 exhibited an effective degradation of phenol at 500 and 750 mg l−1 concentration. Two strains (Fig. 3), A011 and L012, were able to grow in the presence of phenol at 1000 mg l−1 concentration. In all experiments, complete degradation of phenol was possible in around 2 days. For all analyzed yeast strains, higher initial phenol concentration required longer incubation time what could be associated with potential toxic phenol influence on this yeast metabolism. None of the selected strain was able to grow in the presence of phenol concentration (at 1500 or 2000 mg l−1, respectively). Presumably, phenol at these concentrations had a toxic influence on a growth rate of all examined strains. Besides, the production and accumulation of other intermediates of phenol degradation pathways may be responsible for decreasing cell mass yield and toxicity of phenol (Allsop et al. 1993).

Among psychrotolerant microorganism, some strains of Rhodococcus spp. were able to mineralize phenol in a concentration ranging from 1 to 12.5 mM (around 94–1176 mg l−1) (Margesin et al. 2005). Degradation was done within 5 days while higher concentration was done in 14 days. Considering yeast strains, 400 mg l−1 concentration was investigated (Bergauer et al. 2005). Moreover, Margesin et al. (2005) pointed out yeast isolates able to degrade phenol concentration as high as 15 mM (around 1412 mg l−1). Phenol up to 5 mM (around 470 mg l−1) was degraded within 3 days and 7.5 mM (around 705 mg l−1) within 7 days. The time which was needed to effectively degrade was longer than the results obtained for yeast strains presented in this study. Strains isolated from peat bog exhibited effective phenol degradation in less than 2 days. These comparisons should be considered only as an overview, because specific methodology of each investigation effect on the time and specific rate of phenol degradation.

4.1 Identification of Yeast Strains

Appropriate PCR products (for ITS region or D1/D2 region) were purified, sequenced, and analyzed for nucleotide matching by BLAST (Nucleotide BLAST, high similar sequences—megablast).

For D1/D2 rDNA region of strain A011 (613 bp fragment), the highest DNA sequence identity (99%) and cover (100%) was found with Candida subhashii strain UAMH 10744 (GenBank accession number EU836708.1). Strain UAMH 10744 is a type strain, which was deposited in the University of Alberta Microfungus Collection and Herbarium, Edmonton, Canada. It was also deposited in CBS-KNAW (Centraalbureau voor Schimmelcultures, Utrecht, the Netherland) as CBS 10753 and in Mycobank as MB 512099. For this C. subhashii strain, both conventional fermentation and assimilation tests in liquid were performed (Adam et al. 2009). Analysis of D1/D2 rDNA-based phylogenetic tree (Fig. 4) revealed that strain A011 clustered not only with C. subhashii strain UAMH 10744 but also with C. subhashii strain ATY945 (AB985632.1). However, the characterization of strain ATY945 with biochemical tests was not done (Tanimura et al. 2015).
Fig. 4

Phylogenetic tree based on the D1/D2 region of rDNA for strain A011

For analyzed ITS sequence (565 bp fragment), this strain showed highest DNA sequence identity (96%) and cover (100%) with C. subhashii type strain UAMH 10744 (NR_073356.2). Also a phylogenetic tree based on ITS sequences analysis shows that analyzed strain clustered with strain UAMH 10744 (Fig. 5).
Fig. 5

Phylogenetic tree based on the ITS region of rDNA for strain A011

To sum up, results obtained after above-mentioned analysis show that the strain A011 might belong to C. subhashii. Although the distance and composition of trees branches and clusters led to the hypothesis that clustered might be connected with different yeast species than Candida spp. (Fig. 4 and Fig. 5). Cluster contained strain A011 adjoined with major clusters gathered strains of different species. It was also suggested by Adam et al. (2009) where C. subhashii strain UAMH 10744 belonged to a different branch than other Candida sp. strains.

For D1/D2 rDNA region of strain B021 (558 bp fragment), the highest DNA sequence identity (100%) and cover (96%) was found with Candida oregonensis strain (KY106624.1). This strain was deposited in CBS-KNAW culture collection as CBS 5036. Phylogenetic tree based on D1/D2 region (Fig. 6) shows that selected strain clustered with strain Candida sp. Ht-gut 6-8-01 (AY242332.1) and C. oregonensis (U44815.1), where both sequences show 100% identity between each other. Identification of strain Ht-gut 6-8-01 based on D1/D2 loop sequence of the LSU rRNA gene. Only some of isolates from each LSU genotype were deposited in collections and morphological observations and metabolic tests were performed (Suh et al. 2004). In case of type strain deposited in GenBank as U44815.1, it was maintained in the Agricultural Research Service Culture Collection, National Center for Agricultural Utilization Research, Peionia, III (NRRL Y-5850) and was also deposited in CBS-KNAW as CBS 5036 (Kurtzman and Robnett 1997).
Fig. 6

Phylogenetic tree based on the D1/D2 region of rDNA for strain B021

For ITS region (381 bp fragment), analyzed DNA sequence of strain showed 99% identity and 100% cover also with C. oregonensis strain CBS 5036. ITS region-based phylogenetic tree shows that selected strain clustered also with C. oregonensis strain CBS 5036 (Fig. 7).
Fig. 7

Phylogenetic tree based on the ITS region of rDNA for strain B021

According to these results, strain B021 may be classified as C. oregonensis.

For the D1/D2 rDNA region of strain L012 (613 bp fragment), the highest DNA sequence identity (99%) and cover (100%) were found with Nadsonia starkeyi-henricii culture collection strain CBS 5288 (KY109593.1). For this yeast species, authority name is N. starkeyi-henricii and synonymic names are Schizoblastosporion starkeyi-henricii and Schizoblastosporion starkeyihenricii (Kurtzman and Robnett 2013). Phylogenetic tree generated from the analysis of other closely related D1/D2 rDNA nucleotide sequence revealed that this strain clustered with S. starkeyi-henricii strain NRRL YB-3963 (KC254859.1) and also above-mentioned N. starkeyi-henricii strain CBS 5288 (KY109593.1) (Fig. 8).
Fig. 8

Phylogenetic tree based on the D1/D2 region of rDNA for strain L012

For ITS region BLAST analysis, the L012 strain (571 bp fragment) shows 99% identity of DNA sequence with 99% of cover with analogous DNA sequence N. starkeyi-henricii culture collection strain CBS 5288 (KY109593.1), the same strain which revealed the highest DNA sequence identity for D1/D2 region DNA sequence analysis. ITS region-based phylogenetic tree shows that strain L012 clustered with two N. starkeyi-henricii strains—strain CBS 5288 (KY109593.1) and CBS 2159 (KY105364.1) (Fig. 9).
Fig. 9

Phylogenetic tree based on the ITS region of rDNA for strain L012

To sum up, analysis of D1/D2 rDNA and ITS region led to the hypothesis that strain L012 may be a member of species S. starkeyi-henricii (N. starkeyi-henricii). Moreover, reference strain of S. starkeyi-henricii (ATCC 24615) was also isolated from soil collected from a peat bog. Furthermore, this yeast genus appeared in multiple studies on peats and fens in Europe, Canada, Asia, and New Zealand and in most reports about peat bog microorganisms (Babjeva and Blagodatskaia 1971; Hong et al. 2006; Polyakova et al. 2001). It is considered as a rare genus with not fully classified taxonomy and isolated mainly on peatlands (Golubev and Pfeiffer 2014; Polyakova et al. 2001).

To date, there is no information about studies focused on the evaluation of the usefulness of strains belonging to C. subhashii (Adam et al. 2009; Fricova et al. 2010; Tanimura et al. 2015; Valach et al. 2011; Watanabe et al. 1998) and C. oregonensis (Dohet et al. 2016; Lou et al. 2014; Phaff and do Carmo-Sousa 1962, Rivera et al. 2007) to biodegradation of aromatic hydrocarbons. In this connection, some yeast strains (G28 and G38) belonging to species S. starkeyi-henricii exhibited a potential of assimilation of benzene derivative compounds. According to culture slant method, strain G28 assimilated phenol, catechol, resorcinol, hydroquinone, gentisic acid, and 3-hydroxybenzoic acid at 25 °C. Yeast strain G38 was able to degrade phenol at 10 °C based on yeast enrichment cultures on solid media (Middelhoven et al. 1992).

5 Conclusions

In this study, the potential of psychrotolerant yeasts isolated from peatland to biodegrade phenol was studied for the first time. Phenol as the only carbon source was degraded by three yeast strains isolated from soil and water samples collected from Rucianka peat bog. Each strain was able to degrade phenol at 500 and 750 mg l−1 concentration and two strains at 1000 mg l−1 concentration in the mineral salt medium during incubation at 18 °C. All phenol concentrations were degraded below detectable levels in less than 48 h. None of the strains were able to degrade higher phenol concentration (1500 or 2000 mg l−1).

The data obtained in this study shows that the contribution of cold-adapted yeasts to biodegradation of phenol is underestimated. Most investigations on low temperatures are focused on bacteria, whereas knowledge about yeasts is limited. Moreover, according to literature, biodiversity of yeast isolated from peatlands has received little attention. This study pointed out a new direction to search of psychrotolerant yeasts with biodegradation of organic compound potential. Selected strains, especially strain L012 and strain A011, may be useful for examination of degradation for other monoaromatic toxic compounds, like BTEX group.


Compliance with Ethical Standards


This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.


  1. Adam, H., Groenewald, M., Mohan, S., Richardson, S., Bunn, U., Gibas, C. F. C., Poutanen, S., & Sigler, L. (2009). Identification of a new species, Candida subhashii, as a cause of peritonitis. Medical Mycology, 47(3), 305–311. doi: 10.1080/13693780802380545.CrossRefGoogle Scholar
  2. Aksu, Z. (2001). Equilibrium and kinetic modelling of cadmium(II) biosorption by C. vulgaris in a batch system: effect of temperature. Separation and Purification Technology, 21(3), 285–294. doi: 10.1016/S1383-5866(00)00212-4.CrossRefGoogle Scholar
  3. Aksu, Z., Akpinar, D., Kabasakal, E., & Köse, B. (1999). Simultaneous biosorption of phenol and nickel(II) from binary mixtures onto dried aerobic activated sludge. Process Biochemistry, 35(3), 301–308. doi: 10.1016/S0032-9592(99)00072-2.CrossRefGoogle Scholar
  4. Aleksieva, Z., Ivanova, D., Godjevargova, T., & Atanasov, B. (2002). Degradation of some phenol derivatives by Trichosporon cutaneum R57. Process Biochemistry, 37(11), 1215–1219. doi: 10.1016/S0032-9592(01)00336-3.CrossRefGoogle Scholar
  5. Allsop, P. J., Chisti, Y., Moo-Young, M., & Sullivan, G. R. (1993). Dynamics of phenol degradation by Pseudomonas putida. Biotechnology and Bioengineering, 41(5), 572–580. doi: 10.1002/bit.260410510.CrossRefGoogle Scholar
  6. Aravindhan, R., Rao, J. R., & Nair, B. U. (2009). Application of a chemically modified green macro alga as a biosorbent for phenol removal. Journal of Environmental Management, 90(5), 1877–1883. doi: 10.1016/j.jenvman.2008.12.005.CrossRefGoogle Scholar
  7. ATSDR, Agency for Toxic Substances and Disease Registry. (2015). Toxicological profile for phenol. Accessed 20 December 2016.Google Scholar
  8. Babjeva, I. P., & Blagodatskaia, V. M. (1971). Physiological characteristics and ecology of the yeast Schizoblastosporion starkeyi-henricii Ciferri. Mikrobiologiia, 41(1), 99–104.Google Scholar
  9. Banat, F., Al-Bashir, B., Al-Asheh, S., & Hayajneh, O. (2000). Adsorption of phenol by bentonite. Environmental Pollution, 107(3), 391–398. doi: 10.1016/S0269-7491(99)00173-6.CrossRefGoogle Scholar
  10. Bergauer, P., Fonteyne, P.-A., Nolard, N., Schinner, F., & Margesin, R. (2005). Biodegradation of phenol and phenol-related compounds by psychrophilic and cold-tolerant alpine yeasts. Chemosphere, 59(7), 909–918. doi: 10.1016/j.chemosphere.2004.11.011.CrossRefGoogle Scholar
  11. Boczkaj, G., Makoś, P., & Przyjazny, A. (2016a). Application of dispersive liquid-liquid microextraction and gas chromatography with mass spectrometry for the determination of oxygenated volatile organic compounds in effluents from the production of petroleum bitumen. Journal of Separation Science, 39(13), 2604–2615. doi: 10.1002/jssc.201501355.CrossRefGoogle Scholar
  12. Boczkaj, G., Makoś, P., & Przyjazny, A. (2016b). Application of dynamic headspace and gas chromatography coupled to mass spectrometry (DHS-GC-MS) for the determination of oxygenated volatile organic compounds in refinery effluents. Analytical Methods, 8(17), 3570–3577. doi: 10.1039/C5AY03043A.CrossRefGoogle Scholar
  13. Boczkaj, G., Przyjazny, A., & Kamiński, M. (2014). New procedures for control of industrial effluents treatment processes. Industrial & Engineering Chemistry Research, 53(4), 1503–1514. doi: 10.1021/ie402126d.CrossRefGoogle Scholar
  14. Boroujeni, N. A., Hassanshahian, M., Mohammad, S., & Khoshrou, R. (2014). Isolation and characterization of phenol degrading bacteria from Persian Gulf. International Journal of Advanced Biological and Biomedical Research, 2(2), 408–416.Google Scholar
  15. Broholm, M. M., & Arvin, E. (2000). Biodegradation of phenols in a sandstone aquifer under aerobic conditions and mixed nitrate and iron reducing conditions. Journal of Contaminant Hydrology, 44(3–4), 239–273. doi: 10.1016/S0169-7722(00)00103-0.Google Scholar
  16. Bux, F., Atkinson, B., & Kasan, H. C. (1999). Zinc biosorption by waste activated and digested sludges. Water Science and Technology, 39(10), 127–130. doi: 10.1016/S0273-1223(99)00262-0.CrossRefGoogle Scholar
  17. Chakraborty, S., Bhattacharya, T., Patel, T. N., & Tiwari, K. K. (2010). Biodegradation of phenol by native microorganisms isolated from coke processing wastewater. Journal of Environmental Biology, 31(3), 293–296.Google Scholar
  18. Chung, T.-P., Tseng, H.-Y., & Juang, R.-S. (2003). Mass transfer effect and intermediate detection for phenol degradation in immobilized Pseudomonas putida systems. Process Biochemistry, 38(10), 1497–1507. doi: 10.1016/S0032-9592(03)00038-4.CrossRefGoogle Scholar
  19. Cordova-Rosa, S. M., Dams, R. I., Cordova-Rosa, E. V., Radetski, M. R., Corrêa, A. X. R., & Radetski, C. M. (2009). Remediation of phenol-contaminated soil by a bacterial consortium and Acinetobacter calcoaceticus isolated from an industrial wastewater treatment plant. Journal of Hazardous Materials, 164(1), 61–66. doi: 10.1016/j.jhazmat.2008.07.120.CrossRefGoogle Scholar
  20. Davidson, R. S. (1996). The photodegradation of some naturally occurring polymers. Journal of Photochemistry and Photobiology B: Biology, 33(1), 3–25. doi: 10.1016/1011-1344(95)07262-4.CrossRefGoogle Scholar
  21. Dohet, L., Grégoire, J.-C., Berasategui, A., Kaltenpoth, M., & Biedermann, P. H. W. (2016). Bacterial and fungal symbionts of parasitic Dendroctonus bark beetles. FEMS Microbiology Ecology, 92(9). doi: 10.1093/femsec/fiw129.
  22. Federal State Statistics Service: Russia Federation. (2011), Accessed 20 Dec 2016.
  23. Feng, Z., Kunyan, C., Jiamo, F., Guoying, S., & Huifang, Y. (2002). Biodegradability of di(2-ethylhexyl) phthalate by Pseudomonas fluorescens FS1. Water, Air, and Soil Pollution, 140(1/4), 297–305. doi: 10.1023/A:1020108502776.CrossRefGoogle Scholar
  24. Fialová, A., Boschke, E., & Bley, T. (2004). Rapid monitoring of the biodegradation of phenol-like compounds by the yeast Candida maltosa using BOD measurements. International Biodeterioration & Biodegradation, 54(1), 69–76. doi: 10.1016/j.ibiod.2004.02.004.CrossRefGoogle Scholar
  25. Fricova, D., Valach, M., Farkas, Z., Pfeiffer, I., Kucsera, J., Tomaska, L., & Nosek, J. (2010). The mitochondrial genome of the pathogenic yeast Candida subhashii: GC-rich linear DNA with a protein covalently attached to the 5′ termini. Microbiology (Reading, England), 156(Pt 7), 2153–2163. doi: 10.1099/mic.0.038646-0.CrossRefGoogle Scholar
  26. Galíndez-Mayer, J., Ramón-Gallegos, J., Ruiz-Ordaz, N., Juárez-Ramírez, C., Salmerón-Alcocer, A., & Poggi-Varaldo, H. M. (2008). Phenol and 4-chlorophenol biodegradation by yeast Candida tropicalis in a fluidized bed reactor. Biochemical Engineering Journal, 38(2), 147–157. doi: 10.1016/j.bej.2007.06.011.CrossRefGoogle Scholar
  27. García-Peña, I., Ortiz, I., Hernández, S., & Revah, S. (2008). Biofiltration of BTEX by the fungus Paecilomyces variotii. International Biodeterioration & Biodegradation, 62(4), 442–447. doi: 10.1016/j.ibiod.2008.03.012.CrossRefGoogle Scholar
  28. Gimeno, O., Carbajo, M., Beltran, F., & Rivas, F. (2005). Phenol and substituted phenols AOPs remediation. Journal of Hazardous Materials, 119(1–3), 99–108. doi: 10.1016/j.jhazmat.2004.11.024.CrossRefGoogle Scholar
  29. Golubev, W. I., & Pfeiffer, I. (2014). A study of Schizoblastosporion starkeyi-henricii isolates from northern and southern hemispheres of the Earth. Microbiology, 83(5), 661–665. doi: 10.1134/S0026261714050117.CrossRefGoogle Scholar
  30. Gurujeyalakshmi, G., & Oriel, P. (1989). Isolation of phenol-degrading Bacillus stearothermophilus and partial characterization of the phenol hydroxylase. Applied and Environmental Microbiology, 55(2), 500–502.Google Scholar
  31. Hameed, B. H., & Rahman, A. A. (2008). Removal of phenol from aqueous solutions by adsorption onto activated carbon prepared from biomass material. Journal of Hazardous Materials, 160(2), 576–581. doi: 10.1016/j.jhazmat.2008.03.028.CrossRefGoogle Scholar
  32. Hesham, A. E.-L., Khan, S., Liu, X., Zhang, Y., Wang, Z., & Yang, M. (2006). Application of PCR–DGGE to analyse the yeast population dynamics in slurry reactors during degradation of polycyclic aromatic hydrocarbons in weathered oil. Yeast, 23(12), 879–887. doi: 10.1002/yea.1401.CrossRefGoogle Scholar
  33. Hill, G. A., Milne, B. J., & Nawrocki, P. A. (1996). Cometabolic degradation of 4-chlorophenol by Alcaligenes eutrophus. Applied Microbiology and Biotechnology, 46(2), 163–168. doi: 10.1007/s002530050799.CrossRefGoogle Scholar
  34. Holder, A. W., Bedient, P. B., & Hughes, J. B. (1999). Modeling the impact of oxygen reaeration on natural attenuation. Bioremediation Journal, 3(2), 137–149. doi: 10.1080/10889869991219451.CrossRefGoogle Scholar
  35. Hong, S. G., Lee, K. H., Kwak, J., & Bae, K. S. (2006). Diversity of yeasts associated with Panax ginseng. Journal of microbiology (Seoul, Korea), 44(6), 674–679.Google Scholar
  36. ICIS. (2016). Phenol and acetone—under pressure. Market outlook. Accessed 27 March 2016.Google Scholar
  37. Jarvinen, K. T., Melin, E. S., & Puhakka, J. A. (1994). High-rate bioremediation of chlorophenol-contaminated groundwater at low temperatures. Environmental Scence and Technology, 28, 2387–2392. doi: 10.1021/es00062a025.CrossRefGoogle Scholar
  38. Jiang, Y., Ren, N., Cai, X., Wu, D., Qiao, L., & Lin, S. (2008). Biodegradation of phenol and 4-chlorophenol by the mutant strain CTM 2. Chinese Journal of Chemical Engineering, 16(5), 796–800. doi: 10.1016/S1004-9541(08)60158-5.CrossRefGoogle Scholar
  39. Juárez-Ramírez, C., Ruiz-Ordaz, N., Cristiani-Urbina, E., & Galíndez-Mayer, J. (2001). Degradation kinetics of phenol by immobilized cells of Candida tropicalis in a fluidized bed reactor. World Journal of Microbiology and Biotechnology, 17(7), 697–705. doi: 10.1023/A:1012979100827.CrossRefGoogle Scholar
  40. Kafilzadeh, F., Farhangdoost, M.-S., & Tahery, Y. (2010). Isolation and identification of phenol degrading bacteria from Lake Parishan and their growth kinetic assay. African Journal of Biotechnology, 9(40), 6721–6726. doi: 10.5897/AJB10.665.Google Scholar
  41. Karimi, M., Hassanshahian, M., Karimi, M., & Hassanshahian, M. (2016). Isolation and characterization of phenol degrading yeasts from wastewater in the coking plant of Zarand, Kerman. Brazilian Journal of Microbiology, 47(1), 18–24. doi: 10.1016/j.bjm.2015.11.032.CrossRefGoogle Scholar
  42. Kibret, M., Somitsch, W., & Robra, K.-H. (2000). Characterization of phenol degrading mixed population by enzyme assay. Water Research, 34(4), 1127–1134. doi: 10.1016/S0043-1354(99)00248-1.CrossRefGoogle Scholar
  43. Kotturi, G., Robinson, C., & Inniss, W. (1991). Phenol degradation by a psychrotrophic strain of Pseudomonas putida. Applied Microbiology and Biotechnology, 34(4), 539–543. doi: 10.1007/BF00180585.CrossRefGoogle Scholar
  44. Kumar, A., Kumar, S., & Kumar, S. (2005). Biodegradation kinetics of phenol and catechol using Pseudomonas putida MTCC 1194. Biochemical Engineering Journal, 22(2), 151–159. doi: 10.1016/j.bej.2004.09.006.CrossRefGoogle Scholar
  45. Kurtz, A. M., & Crow, S. A. (1997). Transformation of chlororesorcinol by the hydrocarbonoclastic yeasts Candida maltosa, Candida tropicalis, and Trichosporon oivide. Current Microbiology, 35(3), 165–168. doi: 10.1007/s002849900232.CrossRefGoogle Scholar
  46. Kurtzman, C. P., & Robnett, C. J. (1997). Identification of clinically important ascomycetous yeast based on nucleotide divergence in the 5′ end of the large-subunit (26S) ribosomal DNA gene. Journal of Clinical Microbiology, 35(5), 1216–1223.Google Scholar
  47. Kurtzman, C. P., & Robnett, C. J. (1998). Identification and phylogeny of ascomycetous yeasts from analysis of nuclear large subunit (26S) ribosomal DNA partial sequences. Antonie Van Leeuwenhoek, 73(4), 331–371. doi: 10.1023/A:1001761008817.CrossRefGoogle Scholar
  48. Kurtzman, C. P., & Robnett, C. J. (2013). Alloascoidea hylecoeti gen. nov., comb. nov., Alloascoidea africana comb. nov., Ascoidea tarda sp. nov., and Nadsonia starkeyi-henricii comb. nov., new members of the Saccharomycotina (Ascomycota). FEMS Yeast Research, 13(5), 423–432. doi: 10.1111/1567-1364.12044.CrossRefGoogle Scholar
  49. Lin, S.-H., & Juang, R.-S. (2009). Adsorption of phenol and its derivatives from water using synthetic resins and low-cost natural adsorbents: a review. Journal of Environmental Management, 90(3), 1336–1349. doi: 10.1016/j.jenvman.2008.09.003.CrossRefGoogle Scholar
  50. Lou, Q.-Z., Lu, M., & Sun, J.-H. (2014). Yeast diversity associated with invasive Dendroctonus valens killing Pinus tabuliformis in China using culturing and molecular methods. Microbial Ecology, 68(2), 397–415. doi: 10.1007/s00248-014-0413-6.CrossRefGoogle Scholar
  51. Margesin, R., Bergauer, P., & Gander, S. (2004). Degradation of phenol and toxicity of phenolic compounds: a comparison of cold-tolerant Arthrobacter sp. and mesophilic Pseudomonas putida. Extremophiles, 8(3), 201–207. doi: 10.1007/s00792-004-0378-3.CrossRefGoogle Scholar
  52. Margesin, R., Fonteyne, P.-A., & Redl, B. (2005). Low-temperature biodegradation of high amounts of phenol by Rhodococcus spp. and basidiomycetous yeasts. Research in Microbiology, 156(1), 68–75. doi: 10.1016/j.resmic.2004.08.002.CrossRefGoogle Scholar
  53. Margesin, R., Gander, S., Zacke, G., Gounot, A. M., & Schinner, F. (2003). Hydrocarbon degradation and enzyme activities of cold-adapted bacteria and yeasts. Extremophiles, 7(6), 451–458. doi: 10.1007/s00792-003-0347-2.CrossRefGoogle Scholar
  54. Middelhoven, W. J., de Jong, I. M., & de Winter, M. (1991). Arxula adeninivorans, a yeast assimilating many nitrogenous and aromatic compounds. Antonie Van Leeuwenhoek, 59(2), 129–137. doi: 10.1007/BF00445657.CrossRefGoogle Scholar
  55. Middelhoven, W. J., Koorevaar, M., & Schuur, G. W. (1992). Degradation of benzene compounds by yeasts in acidic soils. Plant and Soil, 145(1), 37–43. doi: 10.1007/BF00009539.CrossRefGoogle Scholar
  56. Morasch, B., Richnow, H. H., Schink, B., Vieth, A., & Meckenstock, R. U. (2002). Carbon and hydrogen stable isotope fractionation during aerobic bacterial degradation of aromatic hydrocarbons. Applied and Environmental Microbiology, 68(10), 5191–5194. doi: 10.1128/aem.68.10.5191-5194.2002.CrossRefGoogle Scholar
  57. Pawłat, H. (1996). Ocena oddziaływania projektowanej eksploatacji torfu i rekultywacji potorfi obiektu “Rucianka” na środowisko przyrodnicze. Ocena Oddziaływania Inwestycji na Środowisko, (Environmental impact evaluation of the peat mining and reclamation of the post-mining pits at the Rucianka peat-bog. Evaluation of the investment environmental impact). Warszawa (in Polish).Google Scholar
  58. Phaff, H. J., & do Carmo-Sousa, L. (1962). Four new species of yeast isolated from insect frass in bark of Tsuga heterophylla (Raf.) Sargent. Antonie Van Leeuwenhoek, 28(1), 193–207. doi: 10.1007/BF02538734.CrossRefGoogle Scholar
  59. Polyakova, A. V., Chernov, I. Y., & Panikov, N. S. (2001). Yeast diversity in hydromorphic soils with reference to a grass–sphagnum wetland in western Siberia and a hummocky tundra region at Cape Barrow (Alaska). Microbiology, 70(5), 617–623. doi: 10.1023/A:1012328710111.CrossRefGoogle Scholar
  60. Polymenakou, P. N., & Stephanou, E. G. (2005). Effect of temperature and additional carbon sources on phenol degradation by an indigenous soil Pseudomonas. Biodegradation, 16(5), 403–413.CrossRefGoogle Scholar
  61. Puhakka, J. A., Melin, E. S., Järvinen, K. T., Koro, P. M., Rintala, J. A., Hartikainen, P., Shieh, W. K., & Ferguson, J. F. (1995). Fluidized-bed biofilms for chlorophenol mineralization. Water Science and Technology, 31(1), 227–235. doi: 10.1016/0273-1223(95)00170-R.CrossRefGoogle Scholar
  62. Qi, B., Moe, W. M., & Kinney, K. A. (2002). Biodegradation of volatile organic compounds by five fungal species. Applied Microbiology and Biotechnology, 58(5), 684–689. doi: 10.1007/s00253-002-0938-3.CrossRefGoogle Scholar
  63. Reardon, K. F., Mosteller, D. C., Rogers, J. B., DuTeau, N. M., & Kim, K.-H. (2002). Biodegradation kinetics of aromatic hydrocarbon mixtures by pure and mixed bacterial cultures. Environmental Health Perspectives, 110(Suppl 6), 1005–1011.CrossRefGoogle Scholar
  64. Rengaraj, S., Moon, S.-H., Sivabalan, R., Arabindoo, B., & Murugesan, V. (2002). Agricultural solid waste for the removal of organics: adsorption of phenol from water and wastewater by palm seed coat activated carbon. Waste Management, 22(5), 543–548. doi: 10.1016/S0956-053X(01)00016-2.CrossRefGoogle Scholar
  65. Rivera, F. N., Gómez, Z., González, E., López, N., Hernández Rodríguez, C. H., & Zúñiga, G. (2007). Yeasts associated with bark beetles of the genus Dendroctonus Erichson (Coleoptera: Curculionidae: Scolytinae): molecular identification and biochemical characterization. In: Proceedings from the Third Workshop on Genetics of Bark Beetles and Associated Microorganisms. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Research Station, 45–48.Google Scholar
  66. Rocha, L. L., de Aguiar Cordeiro, R., Cavalcante, R. M., do Nascimento, R. F., Martins, S. C. S., Santaella, S. T., & Melo, V. M. M. (2007). Isolation and characterization of phenol-degrading yeasts from an oil refinery wastewater in Brazil. Mycopathologia, 164(4), 183–188. doi: 10.1007/s11046-007-9043-6.CrossRefGoogle Scholar
  67. Roostaei, N., & Tezel, F. H. (2004). Removal of phenol from aqueous solutions by adsorption. Journal of Environmental Management, 70(2), 157–164. doi: 10.1016/j.jenvman.2003.11.004.CrossRefGoogle Scholar
  68. Sampaio, J. P. (1999). Utilization of low molecular weight aromatic compounds by heterobasidiomycetous yeasts: taxonomic implications. Canadian Journal of Microbiology, 45(6), 491–512. doi: 10.1139/w99-020.CrossRefGoogle Scholar
  69. Siva Kumar, N., Venkata Subbaiah, M., Subba Reddy, A., & Krishnaiah, A. (2009). Biosorption of phenolic compounds from aqueous solutions onto chitosan-abrus precatorius blended beads. Journal of Chemical Technology & Biotechnology, 84(7), 972–981. doi: 10.1002/jctb.2120.CrossRefGoogle Scholar
  70. Sivasubramanian, S., & Namasivayam, S. K. R. (2015). Phenol degradation studies using microbial consortium isolated from environmental sources. Journal of Environmental Chemical Engineering, 3(1), 243–252. doi: 10.1016/j.jece.2014.12.014.CrossRefGoogle Scholar
  71. Suh, S.-O., McHugh, J. V., & Blackwell, M. (2004). Expansion of the Candida tanzawaensis yeast clade: 16 novel Candida species from basidiocarp-feeding beetles. International Journal of Systematic and Evolutionary Microbiology, 54, 2409–2429. doi: 10.1099/ijs0.63246-0.CrossRefGoogle Scholar
  72. Tamura, K., Stecher, G., Peterson, D., Filipski, A., & Kumar, S. (2013). MEGA6: Molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution, 30, 2725–2729. doi: 10.1093/molbev/mst197.CrossRefGoogle Scholar
  73. Tanimura, A., Kikukawa, M., Yamaguchi, S., Kishino, S., Ogawa, J., & Shima, J. (2015). Direct ethanol production from starch using a natural isolate, Scheffersomyces shehatae: toward consolidated bioprocessing. Scientific Reports, 5, 9593. doi: 10.1038/srep09593.CrossRefGoogle Scholar
  74. Thormann, M. N., & Rice, A. V. (2007). Fungi from peatlands. Fungal Diversity, 24, 241–299.Google Scholar
  75. Thormann, M. N., Rice, A. V., & Beilman, D. W. (2007). Yeasts in peatlands: a review of richness and roles in peat decomposition. Wetlands, 27(3), 761–773. doi: 10.1672/0277-5212(2007)27[761:YIPARO]2.0.CO.CrossRefGoogle Scholar
  76. Tsuruta, Y., Kitai, S., Watanabe, S., & Inoue, H. (2000). 2-Methoxy-4-(2-phthalimidinyl)phenylsulfonyl chloride as a fluorescent labeling reagent for determination of phenols in high-performance liquid chromatography and application for determination of urinary phenol and p-cresol. Analytical Biochemistry, 280(1), 36–41. doi: 10.1006/abio.2000.4492.CrossRefGoogle Scholar
  77. Valach, M., Farkas, Z., Fricova, D., Kovac, J., Brejova, B., Vinar, T., Pfeiffer, I., Kucsera, J., Tomaska, L., Lang, B. F., & Nosek, J. (2011). Evolution of linear chromosomes and multipartite genomes in yeast mitochondria. Nucleic Acids Research, 39(10), 4202–4219. doi: 10.1093/nar/gkq1345.CrossRefGoogle Scholar
  78. Wang, C. C., Lee, C. M., Lu, C. J., Chuang, M. S., & Huang, C. Z. (2000). Biodegradation of 2,4,6-trichlorophenol in the presence of primary substrate by immobilized pure culture bacteria. Chemosphere, 41(12), 1873–1879. doi: 10.1016/S0045-6535(00)00090-4.CrossRefGoogle Scholar
  79. Wang, J., Ma, X., Liu, S., Sun, P., Fan, P., & Xia, C. (2012). Biodegradation of phenol and 4-chlorophenol by Candida tropicalis W1. Procedia Environmental Sciences, 16, 299–303. doi: 10.1016/j.proenv.2012.10.042.CrossRefGoogle Scholar
  80. Wang, L., Qiao, N., Sun, F., & Shao, Z. (2008). Isolation, gene detection and solvent tolerance of benzene, toluene and xylene degrading bacteria from nearshore surface water and Pacific Ocean sediment. Extremophiles, 12(3), 335–342. doi: 10.1007/s00792-007-0136-4.CrossRefGoogle Scholar
  81. Watanabe, K., Teramoto, M., Futamata, H., & Harayama, S. (1998). Molecular detection, isolation, and physiological characterization of functionally dominant phenol-degrading bacteria in activated sludge. Applied and Environmental Microbiology, 64(11), 4396–4402.Google Scholar
  82. Woertz, J. R., Kinney, K. A., McIntosh, N. D. P., & Szaniszlo, P. J. (2001). Removal of toluene in a vapor-phase bioreactor containing a strain of the dimorphic black yeast Exophiala lecanii-corni. Biotechnology and Bioengineering, 75(5), 550–558. doi: 10.1002/bit.10066.CrossRefGoogle Scholar
  83. Yan, J., Jianping, W., Hongmei, L., Suliang, Y., & Zongding, H. (2005). The biodegradation of phenol at high initial concentration by the yeast Candida tropicalis. Biochemical Engineering Journal, 24(3), 243–247. doi: 10.1016/j.bej.2005.02.016.CrossRefGoogle Scholar
  84. You, Y., Shim, J., Cho, C.-H., Ryu, M.-H., Shea, P. J., Kamala-Kannan, S., Chae, J. C., & Oh, B. T. (2013). Biodegradation of BTEX mixture by Pseudomonas putida YNS1 isolated from oil-contaminated soil. Journal of Basic Microbiology, 53(5), 469–475. doi: 10.1002/jobm.201200067.CrossRefGoogle Scholar
  85. Yu, Z., & Wen, X. (2005). Screening and identification of yeasts for decolorizing synthetic dyes in industrial wastewater. International Biodeterioration & Biodegradation, 56(2), 109–114. doi: 10.1016/j.ibiod.2005.05.006.CrossRefGoogle Scholar
  86. Zümriye, A., Derya, A., Elif, K., & Burcin, K. (1999). Simultaneous biosorption of phenol and nickel(II) from binary mixtures onto dried aerobic activated sludge. Process Biochemistry, 35, 301–308. doi: 10.1016/S0032-9592(99)00072-2.CrossRefGoogle Scholar

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Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors and Affiliations

  1. 1.Department of Molecular Biotechnology and MicrobiologyGdańsk University of TechnologyGdańskPoland
  2. 2.Department of Chemical and Process EngineeringGdańsk University of TechnologyGdańskPoland
  3. 3.Fermentum Mobile Sp. z o.o. [Ltd.]GdańskPoland

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