IMPACT OF NONYLPHENOL ON THE PHYSIOLOGICAL ACTIVITY OF FUNGI FROM THE COASTAL AREA OF THE GULF OF FINLAND
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IMPACT OF NONYLPHENOL ON THE PHYSIOLOGICAL ACTIVITY
OF FUNGI FROM THE COASTAL AREA OF THE GULF OF FINLAND
Irina Kuzikova, Saint-Petersburg Scientific-Research Centre for Ecological Safety RAS
(SRCES RAS), Russia
Vera Safronova, All-Russia Research Institute for Agricultural Microbiology (ARRIAM),
Russia
Nadezda Medvedeva, Saint-Petersburg Scientific-Research Centre for Ecological Safety RAS
(SRCES RAS), Russia
ngmedvedeva@gmail.com
Nonylphenol (NP) is the most abundant environmental estrogen listed as one of the
priority hazardous substances in the Water Framework Directive (EC 2000) and the
priority pollutant of Baltic Sea (HELCOM 2010). The present study aims to compare the
effects of technical nonylphenol (tNP) on the cellulase, amylase and protease activity of the
terrestrial fungal strains played a significant role in aquatic ecosystems due to their high
adaptive capacity and a large range of functional activity. The study also attempts to
understand the mechanisms behind the varying sensitivity of the terrestrial fungi to tNP.
The fungal strains were isolated from the bottom sediments of the coastal area of the
eastern part of the Gulf of Finland. The terrestrial fungi were identified based on their
morphological characteristics and nucleotide sequence analysis of internal transcribed
space region. One reason for significant differences in sensitivity to the toxicant studied
among the fungi is the change in the fungal cell permeability, in particular in cell
membrane permeability, induced by NP. Environmentally relevant concentrations of tNP
cause significant changes in activity of hydrolytic enzymes in the terrestrial fungi
Aspergillus tubingensis, Penicillium expansum, Penicillium glabrum, and Cadophora
fastigiata involved in organic matter degradation in bottom sediments. There can be
increasing or decreasing trend, depending on both the type of enzyme and the tNP
concentration. The revealed changes may disrupt the destructive processes in bottom
sediments, as well as succession and stability of microbial communities functioning in the
aquatic environment. It was found that tNP contributes to the activation of proteolytic
enzymes, considered as potential fungal virulence factors. This may lead to emergence
fungal strains with enhanced virulence in aquatic microbiocenoses. The investigations of
the physiological responses of terrestrial fungi under nonylphenol will be important for
biochemical processes dynamics and their environmental consequences evaluation.
Key words: coastal area, nonylphenol, fungi, bottom sediments, hydrolytic enzymes activity
I. INTRODUCTION
During many years the Baltic Sea is under an increasing anthropogenic pressure. The
main problems of the Baltic Sea pollution, particularly in the eastern part of the Gulf of Finland,
include the release of contaminants from wastewater discharges, development of navigation andconstruction of oil terminals on the shores and raised beaches of the Gulf. The Gulf of Finland is
affected by organic and metal pollution [1-3].
In recent decades, there has been increasing concern about environmental pollution with
endocrine-disrupting chemicals (EDCs). Due to their widespread presence in the environment
and toxic activity, EDCs have received increased attention in water quality management and
health care. Among EDCs, nonylphenol holds a prominent place.
NP is the most abundant environmental estrogen listed as one of the priority hazardous
substances in the Water Framework Directive (EC 2000) and the priority pollutant of Baltic Sea
(HELCOM 2010). It is used for the production of nonylphenol polyethoxylates (NPEOs) which
have been widely used as surfactants in industrial processes and households [4].
Nonylphenol enters the environment primarily through wastewater pathways. On entering
aqueous environment, NP with its high hydrophobicity (log K ow 4.8–5.3) and low water
solubility (5.43 mg/L at 20°C) is transferred to the near-bottom layers and is accumulated in
sediments and aquatic organisms [5]. As consequence, the transfer and accumulation of NP with
increasing trophic level leads to serious ecotoxicological risk [6]. Organisms that are preferred in
toxicity assessments of NP include algae, fish and invertebrates. Among them, algae have the
largest capacity for NP bioaccumulation, with NP concentrations ranging from 1.5 to 38 mg/kg
and bioconcentration factors, from 200 to 10,000. In fish, the concentrations vary from 0.03 to
1.59 mg/kg, with bioconcentration factors ranging from 13 to 408 [7, 8]. The NP levels in
bottom sediments vary from 0.01 to 1240 mg/kg sediments, reaching 3500 mg/kg in some cases
[7].
Bottom sediments are sites of intense biogeochemical cycling regulated by
microorganisms including terrestrial fungi. Fungi play a significant role in aquatic ecosystems
due to their high adaptive capacity and a large range of functional activity. They show high
efficiency in transforming organic substrates in aquatic ecosystems. Compared to other
organisms, fungi are considered to be fairly resistant to toxicants. This is the reason why
terrestrial fungi are often one of the dominant species in sediments contaminated with toxic
chemicals [9].
At present, only a few studies have been conducted on NPʼs toxicity to fungi.
Nonylphenol exerts toxic effects on the growth of filamentous fungi Neurospora crassa [10],
Fusarium oxysporum and Fusarium solani [11], and Metarhizium robertsii [12]. Under growth
suppression conditions, inhibition of fungal respiration and changes in fungal morphology were
observed. In Neurospora crassa, Fusarium solani and Metarhizium robertsii under NP treatment
cell shapes were abnormal and hyphal apical dominance was lost. These abnormalities were
presumably due to disruption of the hyphal free cytosolic Ca2+ gradient, the H+ gradient, and the
actin cytoskeleton of the apical cells [10]. In NP-treated Metarhizium robertsii samples, fungal
hyphae exhibited ultrastructural changes at the cytoplasmic level, with major differences
detected in vacuoles, mitochondria and cell walls [12].
Long-term exposure to low NP concentrations of 0.004 to 0.06 mg/L caused increased
biomass production in the fungi Fusarium oxysporum and Fusarium solani. Moreover, a strong
stimulation of spore production and germination was observed for Fusarium oxysporum [11].
However, until now, there have only been a few reports on the effects of NP on the
physiological activity of filamentous fungi. Our previous studies have noted the influence of
technical nonylphenol on cellulase and amylase activity of some terrestrial fungal strains of
genera Aspergillus, Cladosporium, Exophiala, and Penicillium [13].The present study aims to compare the effects of NP on the cellulase, amylase and
protease activity of the terrestrial fungal strains isolated from the bottom sediments of the coastal
area of the eastern part of the Gulf of Finland, which have different sensitivities to NP. The study
also attempts to understand the mechanisms behind the varying sensitivity of the terrestrial fungi
to NP.
II. MATERIALS AND METHODS
Chemicals
Technical nonylphenol (CAS: 84852-15-3) was purchased from Sigma-Aldrich, USA.
The other chemicals were obtained from Cryochrom, Russia.
Fungal strains and identification
The fungal strains Aspergillus tubingensis F11, Cadophora fastigiata F 17, Penicillium
expansum F 44, and Penicillium glabrum F 41 used in this work were isolated from the bottom
sediments of the coastal area of the eastern Gulf of Finland.
The fungal isolates were identified based on their morphological characteristics [14, 15]
and a nucleotide sequence analysis of the internal transcribed space (ITS) region.
Genomic DNA was isolated using a reagent kit, an AxyPrep Multisource Genomic DNA
Miniprep Kit (Corning, USA), in accordance with the manufacturer's instructions. The following
PCR primers were used for sequencing the ITS1-5.8S-ITS2 region: ITS1 5’-
TCCGTAGGTGAACCTGCGG-3’ and ITS4 5’-TCCTCCGCTTATTGATATGC-3’ [16]. PCR
was performed in 25-μL reaction mixtures containing 200 μM dNTPs (Helicon, Russia), 5 pmol
of each primer (Eurogen, Russia), 1 U of Taq polymerase (Helicon, Russia) and 20 - 50 ng of
purified template DNA. For amplification, a C1000TM Thermal Cycler was used (BioRad, USA).
The PCR conditions were as follows: initial denaturation at 95oC for 3 min and 30 sec; 35 cycles
of denaturation at 94oC for 1 min, annealing at 54oC for 1 min and extension at 72oC for 2 min;
and a final extension at 72oC for 6 min and 10 sec. Electrophoresis was carried out with 1%
agarose gel (Invitrogen, USA) in TAE. A 100-bp GeneRuler™ and Lambda DNA/HindIII
markers (Fermentas, USA) were used for the sizing and approximate quantification of the DNA
fragments. Purification of the PCR products was usually performed using a PureLink™ Quick
kit (Invitrogen, USA) according to the manufacturer’s instructions. Direct sequencing of the
PCR products was carried out using an ABI PRISM 3500xl genetic analyzer (Applied
Biosystems, USA). The sequences were compared with related sequences available in the
GenBank databases using BLAST analysis (http://www.ncbi.nlm.nih.gov).
Experimental set-up
The fungal cultures were grown in liquid media at 25°C on a rotary shaker Certomat BS-
1 (230 rpm). A spore suspension of the fungal cells with the titer of 1-2·106 CFU/mL was used
for inoculation of the culture media. The fungal biomass was determined by measuring its dry
cell weight.
tNP dissolved in ethanol (125 mg/mL stock solution) was aseptically added to the culture
media to reach the required concentration. The ethanol content, 0.04% v/v, was constant in all
the variants. The control culture media were supplemented with the same amount of ethanol.
The effect of ethanol (applied to dissolve tNP) on the growth of the fungi was found to be
negligible (data not shown).Permeability assays
For the analysis of the cell permeability, we used cultures of the fungi grown to stationary
phase in a liquid Czapek medium with 2 % glucose.
The changes of the cell permeability of the terrestrial fungi exposed to tNP were
monitored from the "loss" by the cells of metabolites exhibiting an absorption band in the
ultraviolet (220-350 nm) [17]. A 200-mg weighed portion of mycelium was resuspended in 20
mL of distilled water and incubated for 1 h on a rotary shaker (230 rpm) at 30°С. The
supernatant was analyzed with a Genesys 10 UV scanning spectrophotometer (Thermo
Spectronic, USA). The permeability was expressed as arbitrary units per gram of dry weight
biomass (d.w.b.).
Hydrolytic enzymes assays
For the analysis of сellulolytic enzyme activity we used 7-day cultures of the fungi grown
in a liquid Hutchinson medium with 1% sodium carboxymethylcellulose (Na-CMC). The
enzymatic activity of cellulase was determined with the use of Na-CMC according to the
procedures described by Li et al. [18]. The results were expressed in micrograms of glucose per
microgram of protein.
The biomass production in these experiments was estimated from the resultant protein
content determined by the method of Lowry et al. [19].
For the assay of proteolytic enzyme activity, the strains were grown in a liquid medium
containing (in g/L) MgSO 4 – 0.52; KCl – 0.52; KH 2 PO4 – 1.52; FeSO 4 ·7H 2 O – 0.01;
ZnSO 4 ·7H 2 O – 0.01; glucose – 20.0; and albumin – 10.0 for 5 days. The extracellular
proteolytic activity was determined according to the procedures described by Liu et al. [20]. The
results were expressed as units per gram of dry weight biomass.
For the analysis of the amylase activity, we used 5-day cultures of the fungi grown in a
liquid Czapek medium with 2% soluble starch at 28°C. The amylolytic activity was determined
using the colorimetric procedure based on starch hydrolysis by amylolytic complex enzymes to
dextrins of varying molecular weight according to Sandhu et al. [21]. The results were expressed
in grams of hydrolyzed starch per gram of dry weight biomass.
Statistical analyses
All statistical analyses were performed with Statistica software (version 6; Statsoft). All
of the data are presented as the mean ± SD of triplicates (n = 3). The data were tested with
standard variance ANOVA, followed by Student’s t-test to determine significant differences. The
differences were considered significant at P≤0.05
III. RESULTS AND DISCUSSION
In this study we used the terrestrial fungal strains isolated from the bottom sediments of the
coastal area of the eastern part of the Gulf of Finland.
The fungi were identified based both on the morphological characteristics according to the
most common criteria [14, 15] and on analysis of the sequences of the ITS region of DNA.
For the investigation we selected terrestrial fungal strains that exhibited different
sensitivities to NP (Table 1).The strains investigated can be arranged in increasing order of their sensitivity to NP in the
following sequence: Penicillium expansum F 44< Penicillium glabrum F 41100.0
Cadophora fastigiata F 17 1.0 7.0
Penicillium glabrum F 41 15.0 >100.0
Penicillium expansum F 44 20.0 >100.0
*ЕС 50 and ЕС 90 are the effective concentrations of 50 and 90% toxicant inhibition of fungal growth, respectively.
The values of the toxicity parameters were calculated per 48 hours.
One reason for such significant differences in sensitivity to the toxicant studied among
the fungi may be the change in the fungal cell permeability, in particular in cell membrane
permeability, induced by NP.
The cytoplasmatic membrane is the primary target of negative impact of many chemical
substances [22]. For all living cells, the ion transport through the cell membrane is essential for
maintaining the ionic and osmotic homeostasis of the cell, as well as for information transfer,
energy supply for cellular metabolism, substrate accumulation, and degradation products
removal [23].
Permeability describes the ease with which ions can pass through a cell membrane to
move substances into and out of the cell. Various toxicants have been reported to cause changes
in permeability of fungal cell membranes [24, 25] in consequence of adaptation to the toxicant
action. One factor that may be responsible for changes in cell membrane permeability is
oxidation of membrane lipids [26]. Enhancement of membrane lipid peroxidation under NP-
induced oxidative stress was observed in microalgae [27, 28].
Under tNP exposure, Cadophora fastigiata F 17 strain, the most sensitive to tNP,
exhibited a 1.6-fold increase in the cellular permeability relative to the control (without tNP)
(Table 2).
Table 2. Effect of nonylphenol on the cell permeability of the terrestrial fungi
Fungal culture tNP content, Permeability,
mg/L % to control
Aspergillus tubingensis F11 50.0 45±10
Cadophora fastigiata F 17 1.0 162±17
Penicillium expansum F 44 50.0 85±18
Penicillium glabrum F 41 50.0 91±14
Increased cellular permeability may facilitate the entry of toxicant into the cell, as well as
the loss of vital metabolites. In tNP-resistant strains of the filamentous fungi the permeability of
cell membranes either remained practically unchanged (Penicillium expansum F 44 and P.
glabrum F 41) or significantly, by up to 55%, decreased (Aspergillus tubingensis F11) relative tothe control variants (without tNP). These findings suggested that one possible mechanism behind
high resistance of terrestrial fungi of genera Penicillium and Aspergillus is a decrease in the cell
membrane permeability, which complicates the toxicant penetration into the cell.
Terrestrial fungi, which are an important component of aquatic ecosystems, including
bottom sediments, possess a wide range of extracellular hydrolytic enzymes which enable them
to actively degrade organic matter in the aquatic environment [9]. Therefore, the impact of tNP
on the hydrolytic enzymes involved in fungal degradation of organic matter in water and bottom
sediments is an issue that deserves special attention.
The cellulolytic enzymes performing biodegradation of cellulose, the most abundant
biopolymer on Earth, occupy the central position in the organic carbon cycle. Among the
terrestrial fungi isolated, Aspergillus tubingensis F11, Penicillium expansum F 44, and
Penicillium glabrum F 41 strains exhibited cellulase activities. As shown by our previous study
[29], the trend in the cellulase activity in the Penicillium expansum F 44 strain under the tNP
influence depends on the tNP concentration. At low tNP concentrations (up to 1.0 mg/L) that do
not significantly affect the Penicillium expansum F44 strain growth, the enzyme activity
increased by 125% relative to the control (without tNP). An increase in tNP concentration in the
medium to >1.0 mg/L resulted in both the culture growth inhibition and reduction in the cellulase
activity, to 71% of the control value at tNP concentration of 10.0 mg/L. The cellulase activity of
the Aspergillus tubingensis F11 strain was affected by tNP in a similar way (Fig. 1). By contrast
to Penicillium expansum F 44 and Aspergillus tubingensis F11 strains, Penicillium glabrum F 41
exhibited a significant reduction in the cellulase activity both at low tNP concentrations that left
the fungal growth practically unaffected (up to 5 mg/L) and at the growth inhibiting tNP
concentrations (10.0 mg/L) (Fig.1).
250
Extracellular cellulase activity,
200
% to control
150
100
50
0
1 5 10
tNP content, mg/L
Aspergillus tubingensis F 11 Penicillium expansum F 44 P. glabrum F 41
Fig.1. Effect of tNP on the cellulolytic enzyme activity of the terrestrial fungi. The samples
were taken in three independent trials.Starch-hydrolyzing amylolytic enzymes were detected in all the terrestrial fungi
investigated in this study. The tNP effect on the amylase activity of the terrestrial fungi was
found to be species-nonspecific. Under tNP exposure, a decrease in amylase activity by 35 to
60%, depending on the species to which the strain belongs, was observed for all the strains
investigated (Fig.2). It should be noted that the inhibitory effect of tNP on the amylase activity of
the filamentous fungi was observed both at tNP concentrations that have no effect on the
micromycete growth and at the growth inhibiting tNP concentrations.
Amylase activity, % to control 80
70
60
50
40
30
20
10
0
Aspergillus Cadophora Penicillium P. glabrum F 41
tubingensis F 11 fastigiata F 17 expansum F 44
Fig.2. Effect of tNP on the amylase activity of the terrestrial fungi. The samples were taken in
three independent trials.
Our previous study [13] has revealed similar effects from tNP treatments on the cellulase
and amylase activities of other fungal strains of the genera Aspergillus, Penicillium,
Сladosporium and Exophiala.
Along with cellulase and amylase activity, the activity of proteolytic enzymes as
influenced by tNP exposure of the terrestrial fungi seemed to be an important research subject.
This is due to the fact that not only these enzymes are known for their participation in protein
breakdown in bottom sediments but also the secreted proteases have been intensively
investigated as potential virulence factors of fungi [30].
Using the Aspergillus tubingensis F11 and Penicillium expansum F 44 strains as an
example, we demonstrated that, under tNP exposure, the protease activity of the strains isolated
increased 1.4-1.5 times (Fig.3).200
Extracellular protease activity, %
150
to control
100
50
0
Aspergillus Penicillium expansum
tubingensis F 11 F 44
Fig.3. Effect of tNP (20 mg/L) on the protease activity of the terrestrial fungi. The samples were
taken in three independent trials.
In our previous studies [13, 29] we have shown that tNP can also increase the synthesis of
other pathogenicity factors of fungi, pigments and polysaccharides.
Our data suggest that tNP has a potential to enhance fungal pathogenicity, which may lead
to adverse environmental impacts, namely, to emergence of strains with enhanced virulence.
IV. CONCLUSIONS
Thus, terrestrial fungal species having different resistances to tNP were isolated from the
bottom sediments of the coastal area of the eastern part of the Gulf of Finland. One reason for the
differences in sensitivity to tNP among the fungi is presumably the disturbance of the cellular
permeability. Environmentally relevant concentrations of tNP cause significant changes in
activity of hydrolytic enzymes (cellulases, proteases and amylases) in the terrestrial fungi
Aspergillus tubingensis, Penicillium expansum, Penicillium glabrum, and Cadophora fastigiata
involved in organic matter degradation in bottom sediments. There can be increasing or
decreasing trend, depending on both the type of enzyme and the tNP concentration. The revealed
changes may disrupt the destructive processes in bottom sediments, as well as succession and
stability of microbial communities functioning in the aquatic environment. It was also found that,
along with enhancement of the synthesis of such fungal pathogenicity factors as pigments and
polysaccharides, developed in fungi as adaptive mechanisms, tNP contributes to the activation of
proteolytic enzymes, also considered as potential virulence factors. This may lead to emergence
fungal strains with enhanced virulence in aquatic microbiocenoses. The investigations of the
physiological responses of terrestrial fungi under nonylphenol will be important for biochemical
processes dynamics and their environmental consequences evaluation.V. REFERENCES
[1] A.E. Rybalko, N.K. Fedorova, “Bottom sediments of the Neva estuary and its
contamination under influence of anthropogenic processes”, in: Ecosystem of the Neva Estuary:
Biological Diversity and Ecological Problems (A.F. Alimov, S.M. Golubkov), Eds. KMK, St.
Petersburg—Moscow, 2008, pp. 39–58 (in Russian).
[2] H. Vallius, “Arsenic and heavy metal distribution in the bottom sediments of the Gulf
of Finland through the last decade”, Baltica 25, 2012, pp. 23–32.
[3] A.A. Eglit, N.V. Orlova, K.V. Ostrikov, A.V. Vlasov, V.M. Skvortsov, I.I.
Murashko, et al., State of the Environment in Leningrad Region. Committee on Natural
Resources of Leningrad Region, St. Petersburg, 2012, 320 pp. (in Russian).
[4] A. Bergé, J. Gasperi, V. Rocher, L. Gras, A. Coursimault, R. Moilleron, “Phthalates
and alkylphenols in industrial and domestic effluents: Case of Paris conurbation (France)”,
Science of the Total Environment, 2014, pp. 26–35. doi:10.1016/j.scitotenv.2014.04.081.
[5] Y. Kim, G.V. Korshin, A.B. Velichenko, “Comparative study of electrochemical
degradation and ozonation of nonylphenol”, Water Res, 2005, vol. 39, pp. 2527–2534.
[6] D.Y. Shang, R.W. Macdonald, M.G. Ikonomou, “Persistence of nonylphenol
ethoxylate surfactants and their primary degradation products in sediments from near a
municipal outfall in the strait of Georgia, British Columbia, Canada”, Environ. Sci. Technol.,
1999, vol. 33, pp.1366–1372.
[7] A. Soares, B. Guieysse, B. Jefferson, E. Cartmell, J.N. Lester, “Nonylphenol in the
environment: a critical review on occurrence, fate, toxicity and treatment in wastewaters”,
Environ. Int., 2008, vol. 34, pp. 1033–1049.
[8] R. Vazquez-Duhalt, F. Marquez-Rocha, E. Ponce, A. F. Licea, M.T.Viana,
“Nonylphenol, an integrated vision of a pollutant. Scientific Review”, Applied Ecology and
Environmental Research, 2006, vol. 4 (1), pp. 1–25.
[9] V.A.Terekhova, The fungi in ecological assessment of the water and terrestrial
ecosystems, Мoscow, Nauka, 2007, 215p. (in Russian).
[10] A.J. Karley, S.I. Powell, J.M. Davie, “Effect of nonylphenol on growth of
Neurospora crassa and Candida albicans”, Applied and Environmental Microbiology, 1997, vol.
63 (4), pp. 1312–1317.
[11] A. Kollmann, A. Brault, I. Touton, J. Dubroca, V. Chaplain, C. Mougin, “Effect of
nonylphenol surfactants on fungi following the application of sewage sludge on agricultural
soils”, Journal of Environmental Quality, 2003, vol. 32 (4), pp. 1269–1276.
[12] S. Rozalska, S.Glinska, J. Dlugonsky, “Metarhizium robertsii morphological
flexibility during nonylphenol removal”, International Biodeterioration and Biodegradation,
2014, 95. pp. 285–293.
[13] I.L. Kuzikova, E.A. Tileva, T.B. Zaytseva, N.G. Medvedeva, “Effect of
nonylphenol on terrigenous fungi of the coastal zone of the eastern Gulf of Finland”, Mikologiya
i fitopatologiya, . 2015, vol. 49 (4), pp. 249– 256. (in Russian).
[14] K.H Domsch, W. Gams and T.H. Anderson, Compendium of soil fungi: Volume 1,
1980, Academic Press, London, p. 859.
[15] R.A. Samson, and E.S.Van Reenen-Hoekstra, Introduction to food-borne fungi, 3rd
ed. 1988, Baarn, p. 295.
[16] T.J. White, T. Bruns, S. Lee, J. Taylor, 1990. “Amplification and direct sequencing
of fungal ribosomal RNA genes for phylogenetics”, in: PCR Protocols: a guide to methods andapplications, (M.A. Innis, D.H. Gelfand, J.J. Sninsky, T.J. White, eds), Academic Press, New
York, USA, pp. 315–322.
[17] B.A. Fenderson, E.M. Eddy, S.I. Hakomori, “Glycoconjugate expression during
embryogenesis and its biological significance”, BioEssays, 1990, vol. 12 (4). pp. 173 –179.
[18] F. Li, X. Zhu, N. Li, P. Zhang, S. Zhang, X. Zhao, et al., “Screening of
Lignocellulose-Degrading Superior Mushroom Strains and Determination of Their CMCase and
Laccase Activity”, Scientific World Journal, 2014, Article ID 763108, 6. doi:
10.1155/2014/763108.
[19] O.H. Lowry, N.J. Rosebrough, A.L. Farr, R.J. Randall, “Protein measurement with
the Folin phenol reagent”, J. Biol. Chem., 1951, vol. 193 (1), pp. 265–275.
[20] F. Liu, W. Li, D. Ridgway, T. Gu, and M. Moo-Young, “Inhibition of extracellular
protease secretion by Aspergillus niger using cell immobilization”, Biotechnology Letters, 1998,
vol. 20, (6), pp. 539–542.
[21] D.K. Sandhu, K.S. Vilkhu, and S.K. Soni, “Production of α-amylase by
Saccharomyces fibuligera”, J. Ferm. Technol, 1987, vol. 65(4), pp. 387–394. doi:10.1016/0385-
6380(87)90134-8.
[22] G. McDonnell and A. D. Russell, “Antiseptics and Disinfectants: Activity, Action,
and Resistance”, Clin Microbiol Rev, 1999, vol. 12 (1), pp. 147–179.
[23] B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, P. Walter, Molecular Biology
of the Cell, Garland Science, New York, USA, 4th edition, 2002.
[24] N. Medvedeva, Yu. Polyak, I. Kuzikova, O. Orlova, G. Zharikov, “The effect of
mustard gas on the biological activity of soil”, Environmental Research, 2008, vol.106, pp. 289–
295.
[25] C. Yang, C. Hamel, V. Vujanovic, and Y. Gan, “Fungicide: Modes of Action and
Possible Impact on Nontarget Microorganisms”, International Scholarly Research Network,
2011, vol. 2011, Article ID 130289, 8 p. doi:10.5402/2011/130289.
[26] X.X. Tang, T.J. Jan, Y.Q. Li, “Damage effect of monocrotophos on Platymonas sp.
I. Active oxygen in Platymonas sp. cells”, Chinese Journal of Applied Ecology, 1998, vol. 9 (6),
pp. 627–630.
[27] N. Medvedeva, T. Zaytseva, I. Kuzikova, “Cellular responses and bioremoval of
nonylphenol by the cyanobacterium Planktothrix agardhii 1113”, Journal of Marine Systems,
2016, in press.
[28] H. Qian, X. Pan, S. Shi, S.Yu, H. Jiang, Z. Lin, Z. Fu, “Effect of nonylphenol on
response of physiology and photosynthesis-related gene transcription of Chlorella vulgaris”,
Environ. Monit. Assess, 2011, vol.182, pp. 61– 69.
[29] I. Kuzikova, V. Safronova, T. Zaytseva, N. Medvedeva, “Fate and effects of
nonylphenol in the filamentous fungus Penicillium expansum isolated from the bottom sediments
of the Gulf of Finland”, Journal of Marine Systems, 2016, in press.
[30] M. Monod, A. Fatih, L. Jaton-Ogey, S. Paris and J.P. Latge, “The secreted proteases
of pathogenic species of Aspergillus and their possible role in virulence”, Can. J. Bot., 1995,
vol. 73, pp. 1081–1086.You can also read
