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Oxidative stress in bacteria (Pseudomonas putida) exposed to nanostructures of silicon carbide
Chemosphere 135 (2015) 233–239
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Chemosphere journal homepage: www.elsevier.com/locate/chemosphere
Oxidative stress in bacteria (Pseudomonas putida) exposed to nanostructures of silicon carbide _ e, Andrzej Borkowski a,⇑, Mateusz Szala b, Paweł Kowalczyk c, Tomasz Cłapa d, Dorota Narozna Marek Selwet d _ i Wigury 93, 02-089 Warsaw, Poland Faculty of Geology, University of Warsaw, Zwirki Faculty of Advanced Technologies and Chemistry, Military University of Technology, Kaliskiego 2, 00-908 Warsaw, Poland c Bionicum Ltd., Chełmska 21, Warsaw, Poland d Department of General and Environmental Microbiology, Poznan University of Life Sciences, Szydłowska 50, 60-656 Poznan, Poland e Department of Biochemistry and Biotechnology, Poznan University of Life Sciences, Dojazd 11, 60-632 Poznan, Poland a
b
h i g h l i g h t s
g r a p h i c a l a b s t r a c t
Silicon carbide nanostructures were
produced by combustion synthesis. Oxidative stress in Pseudomonas
putida was investigated. Peroxidase activity, expression of zwf-
1 gene and cell viability were analyzed. SiC nanofibers and nanorods can induce oxidative stress in P. putida.
a r t i c l e
i n f o
Article history: Received 12 December 2014 Received in revised form 10 April 2015 Accepted 17 April 2015 Available online 15 May 2015 Keywords: Silicon carbide Nanofibers Nanorods Oxidative stress Pseudomonas putida Toxicity
a b s t r a c t Silicon carbide (SiC) nanostructures produced by combustion synthesis can cause oxidative stress in the bacterium Pseudomonas putida. The results of this study showed that SiC nanostructures damaged the cell membrane, which can lead to oxidative stress in living cells and to the loss of cell viability. As a reference, micrometric SiC was also used, which did not exhibit toxicity toward cells. Oxidative stress was studied by analyzing the activity of peroxidases, and the expression of the glucose-6-phosphate dehydrogenase gene (zwf1) using real-time PCR and northern blot techniques. Damage to nucleic acid was studied by isolating and hydrolyzing plasmids with the formamidopyrimidine [fapy]-DNA glycosylase (also known as 8-oxoguanine DNA glycosylase) (Fpg), which is able to detect damaged DNA. The level of viable microbial cells was investigated by propidium iodide and acridine orange staining. Ó 2015 Elsevier Ltd. All rights reserved.
1. Introduction
⇑ Corresponding author. E-mail address: [email protected] (A. Borkowski). http://dx.doi.org/10.1016/j.chemosphere.2015.04.066 0045-6535/Ó 2015 Elsevier Ltd. All rights reserved.
A review of the literature shows that nanomaterials can be cytoand genotoxic to prokaryotic and eukaryotic cells. Most of the tests on nanostructures were run with one- and two dimension
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nanomaterials: carbon nanotubes, metal nanoparticles, graphene nanoplates, and graphene. Genotoxicity of graphene nanoplates and graphene nanoribbons toward human cells was reported by Akhavan et al. (2012, 2013). There are few studies on silicon carbide (SiC) nanostructures, which can be considered biocompatible and chemically non-active. Szala and Borkowski (2014) reported that SiC nanofibers and nanorods can be cytotoxic to bacteria by mechanically damaging their cell membrane. The toxicological data on mammalian cells in the literature were mainly obtained from experiments conducted on microcrystalline SiC particles or fibers. It has been shown that SiC powder does not induce harmful effects on tissues, which led to the view that microcrystalline SiC material is biologically inert (Bruch et al., 1993a,b). A few studies have described possible toxic effects of exposure of mammalian cells to SiC nanostructures in vitro (Barillet et al., 2010; Pourchez et al., 2012). There are two basic mechanisms of nanostructure toxicity toward living organisms: (i) mechanical damage of the cell membrane and (ii) oxidative stress, which can be caused by mechanical damage of the cell membrane or the presence of reactive forms of oxygen. Reactive forms of oxygen can be present on the surface of nanomaterials because of the high catalytic activity of the materials and their large specific surface areas. In a previous study, SiC nanostructures were covered with a thin layer (thickness of approximately 2 nm) of partially amorphous SiO2 (Huczko et al., 2005). Silica nanofilms can promote the formation of highly reactive hydroxyl radicals and singlet oxygen, which are responsible for hydroxyl radical-mediated DNA damage (Cadet et al., 1999). The toxic effect of silica nanomaterials was reported in relation to both bacteria and mammalian cells (Jiang et al., 2009; Yu et al., 2011). The aim of this study was to determine the dependence between the textures of SiC nanostructures and their ability to cause oxidative stress in Pseudomonas putida. Therefore, bacteria were treated with SiC nanostructures (nanofibers and nanorods, (NFSiC and NRSiC respectively)) and micrometric SiC (lmSiC) as a control. The results show that lmSiC was not toxic to the cells, and that it can be used as a reference material.
(TEM) (Huczko et al., 2005). The results clearly show that NFSiC and NRSiC were covered with a thin layer of silica. SEM images of the investigated materials are presented in Fig. 1. 2.2. Microorganisms and media The strain of P. putida was obtained from our own collection of isolated microorganisms from samples of organic soils (Geomicrobiology Laboratory, Faculty of Geology, University of Warsaw). Taxonomic affiliation was confirmed by sequencing
2. Materials and methods 2.1. Nanomaterials and lmSiC NFSiC were prepared by a combustion route. The combustion mixture was prepared by dry mixing powders of calcium disilicide (CaSi2) and poly(tetrafluoroethene) (PTFE) in a ceramic mortar. After pressing the powders into a cylindrical pellet, 5 g of the sample were placed in a graphite crucible and placed in a stainless steel autoclave (350 cm3 in volume), which was subsequently filled with helium at an initial pressure of 1.0 MPa. The combustion process was initiated with an electrically-heated resistance wire (diameter 0.1 mm). The spongy combustion products were removed from the autoclave with water. The suspension was filtered, and the gray deposit obtained was purified in a three-step process: heating in 98% H2SO4, calcination in air (700 °C), and, finally, heating in 25% NaOH and washing with plenty of water. The NFSiC prepared using this method was used in all experiments. NRSiC were also prepared by the combustion route, but the starting mixture comprised fluorinated graphite (CF) and aluminum–silicon (AlSi). Purification was conducted following the same procedure, but the step of heating in 98% H2SO4 was omitted. lmSiC was purchased from Sigma– Aldrich (St. Louis, MO, USA) and used without purification. Nanomaterials were characterized using X-ray powder diffraction (XRD), Raman spectroscopy, elemental analysis, scanning electron microscopy (SEM) and transmission electron microscopy
Fig. 1. Scanning electron microscopy (SEM) images of silicon carbide nanofibers (NFSiC) (A), nanorods NRSiC (B), and micrometric SiC (lmSiC) used as a reference material (C).
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analysis of the 16S rDNA gene. The bacteria were cultivated in liquid and solid nutrient media (pH 7.5) comprising the following (g/ L): glucose, 10; peptone, 5; yeast extract, 2; NaCl, 4; and agar (in the case of solid medium), 20. The media were autoclaved at 121 °C for 15 min.
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was kept at 1.8–2.0. The DNA solution was stored at 20 °C until further use. The quality of the reaction products were analyzed electrophoretically in 1% agarose gels as the ratio of the covalently closed circular to the open circular form of the plasmid. 2.6. Expression of the zwf1 gene (RNA analysis)
2.3. Cultures of P. putida with investigated SiC materials Forty milligrams of lmSiC, NFSiC, or NRSiC were added to 10 mL of sterile liquid medium in a sterile glass bottle (Simax, 100 mL), and 0.5 mL of P. putida inoculum was added to the mixture. The obtained suspension was incubated at 25 °C under constant stirring (200 rpm) for 24 h. The inoculum was prepared as follows: P. putida was cultured on nutrient agar plates for 24 h at 30 °C. Then, the cultured bacteria were added to 5 mL of NaCl solution (0.9%) to a concentration of approximately 108 colony-forming units (cfu)/mL. After incubation with investigated materials, the peroxidase activity of the obtained bacterial cultures was analyzed, and bacterial DNA and RNA were isolated. 2.4. Assay of peroxidase activity in bacterial cells Peroxidase activity was determined in extracts from bacterial cells grown in liquid medium cultures. At defined times during the bacterial growth culture, cells were harvested by centrifugation and resuspended in 50 mM phosphate buffer (pH 7.8) and sonicated. Sonicates were centrifuged at 13,000 rpm (26,000g) for 30 min, with 10 s intervals to obtain the supernatant, which was used in the assay. The peroxidase activity was measured spectrophotometrically at 420 nm using 50 mM ABTS [2,2-azinobis(3-ethylbenzothiazoline-6-sulfonic acid)] solution in 0.5 M phosphate-citrate buffer (pH 4.0). Measurement of peroxidase activity was performed after incubating for 60 min at 37 °C in a cell counter (Thermo Scientific, Waltham, MA, USA). The colorless ABTS molecule is converted into the blue–green radical, ABTS+, via the oxidation of one electron in the presence of antioxidants in the sample. Additionally, the activity of antioxidants in the samples with lmSiC, NRSiC, or NFSiC without bacteria was measured in the same way as described above. This test was conducted to assess whether the investigated SiC materials can interfere with the measurements of peroxidase activity. Negative control contained buffer without SiC. 2.5. Estimation of oxidative stress in plasmid DNA after digestion with the Fpg protein Plasmids were extracted from cultures: without SiC – control (K), and with lmSiC, NFSiC, or NRSiC using the plasmid mini prep isolation kit (A&A Biotechnology, Gdynia, Poland) according to the manufacturer’s instructions. The purity and concentration of the DNA preparation were determined spectrophotometrically at 260 nm. Next, the obtained samples of DNA were digested with the Fpg protein. The standard reaction mixture (final volume of 20 lL) for the Fpg protein contained 10 lg of plasmid DNA, 0.09 lg of Fpg, 100 mM HEPES-KOH, pH 7.6, 150 mM KCl, 10 mM EDTA, 5 mM MgCl2, 1 mM b-mercaptoethanol, and 100 lg/mL of bovine serum albumin (BSA), and it was incubated for 30 min at 37 °C. After DNA cleavage, the enzyme was removed by chloroform extraction and DNA was precipitated with 4 volumes of cold 96% ethanol and 0.1 volume of 3 M sodium acetate (pH 5.2) at 20 °C overnight, or at 80 °C for 2 h, to sediment all small DNA fragments, and subsequently centrifuged at 12,000 rpm (22,000g) for 15 min. The DNA pellet was resuspended in water, and the DNA concentration was measured using a Varian Cary 3E spectrophotometer with ADL News software (Varian, Sunnyvale, CA, USA). To ensure the complete removal of protein, the A260/A280 ratio
2.6.1. Sample preparation for real-time PCR For this analysis, total RNA was isolated from bacterial cells using Trizol (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. The concentration of total RNA was calculated from its absorbance at 260 nm. Total RNA was used to synthesize cDNA using Advantage kit RTfor-PCR (Clontech Laboratories, Inc., Mountain View, CA, USA) and oligo (dT) primers according to the manufacturer’s instructions. One microgram of total RNA was used as the template, to which was added 1 lL of oligo (dT) primers (2 ng), 1 lL of dNTPs (10 mM final concentration), 4 lL of 5 reaction buffer, and 0.5 lL of RNase. RNA and oligo primers were gently mixed together and heated for 2 min at 70 °C. Then, they were placed on ice and mixed with the other reaction components. Subsequently, 1 lL of reverse transcriptase MMLV (20 U/lL) was added, and the samples were incubated for 60 min at 37 °C. MMLV was inactivated by heating at 95 °C for 5 min. 2.6.2. Real-time PCR with SYBR green An ABI Prism 7500 (Applied Biosystems, Foster City, CA, USA) was used for the analysis. Twenty-five microliters of reaction mixture contained 1 Taq polymerase buffer, 40,000 diluted SYBR green, 3 mM MgCl2, 80 nM of each primer, 400 nM dATP, dGTP, dCTP, and dTTP, and 0.025 U/lL of Taq polymerase. Six hundred forty or 160 ng of cDNA was added to each reaction. To examine the constitutive level of gene expression in the tested bacterial samples, the Ct values of 16S rRNA and zwf1 were investigated. The Ct value is the number of cycles at which the generated fluorescence in the reaction reaches a threshold value, which occurs in the linear stage of the amplification profile. All reactions were performed in two replicates with 640 ng of template and in two replications with 160 ng of template, as well as with a negative control without template. The PCR conditions were as follows: an initial denaturation at 95 °C for 3 min, followed by 45 cycles of 95 °C for 15 s (denaturation), 60 °C for 15 s (annealing), 72 °C for 15 s (extension), and 72 °C for 1 min (final extension). The fluorescence was detected in each cycle after every extension phase. To differentiate specific products from non-specific products, the dissociation curve was created. The Ct values were calculated by SDS v.1.7 software (Applied Biosystems) with the automatic basic curve, which was the median fluorescence value from 3 to 15 cycles. The data were exported to the Microsoft Excel program for further analyses. The control curve was created for the 16S rRNA and zwf-1 genes using a series of dilutions of reference RNAs. The level of transcription of mRNA was normalized to that of the 16S rRNA in each dilution. The standard curve was created based on the normalized transcription level of the initial logarithmic concentration of cDNA in each dilution. As the reference, the median Ct values from three samples were used. The gene ratio was calculated using the equation: ratio = 2 ddCt, where ddCt is the difference between the Ct of the investigated gene samples and the Ct of the reference samples. Additionally, RNA was measured by northern blotting. RNA was isolated from 3 mL of exponentially growing cells using the RNeasy kit (Qiagen) according to the manufacturer’s instructions. RNA was quantified by measuring the optical absorbance at 260 nm (A260) of 4–5 mg of total RNA per sample. Fractionated RNA was transferred to a nylon membrane (GE Healthcare, Little Chalfont,
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United Kingdom) using a Turboblotter (Schleicher & Schuell), and the amounts of zwf1 mRNA were determined by hybridizing the membrane with an a-32P-labelled probe specific for each gene (TaKaRa Bio). To produce probes with high specific activities, the following primers were used: zwf1 pro1 CGCGAATTCCGAGTCGCAGCGTTTTGTCT and zwf1 pro2 CGCGGATCCCCAGCAGGCGGGTGTCC (Kim et al., 2008). The results were compared with 16S rRNA gene sequences, which were a control for the zwf1 gene. 2.7. Viability test To analyze the loss of viability, solutions of propidium iodide (PI) (2 mg/0.1 L, pH = 7.4) and acridine orange (AO) (5 mg/0.1 L, pH = 7.4) were prepared in phosphate buffer. Tests were conducted as follows: in a 100-mL sterile glass bottle (Simax), 10 mL of sterile saline solution (0.9% NaCl) and 40 mg of lmSiC, NFSiC, or NRSiC were added. Subsequently, 1 mL of P. putida inoculum (approximately 108 cfu/mL in 0.9% NaCl) was added to the mixture and mixed (with shaking at 200 rpm) for 120 min at 25 °C. After mixing, 1 mL of suspension was placed into an Eppendorf tube, and 0.3 mL of sucrose (60%) was added. The mixture was centrifuged for 2 min at 4000 rpm (2600g). Sucrose was added to separate unadsorbed bacteria from the investigated nanostructures that had adsorbed to cells. Then, the supernatant (containing free-living cells) and residuum (NFSiC or NRSiC with adsorbed cells) were stained as follows. Initially, 0.2 mL of the supernatant or residuum, and 30 lL of PI were added to a test tube and left for 10 min in the dark. Later, 15 lL of AO was added (staining for 2 min) to the test tube. After staining, 10 lL of suspension was placed on a glass slide and covered with a coverslip. Ten representative fluorescence images of free-living cells and cells adsorbed onto the surface of aggregates of SiC nanostructures were acquired using an epifluorescence microscope with a B-filter. Dead cells appeared red–orange, while viable cells were green. The result of the microscopic analysis was expressed as the ratio of the number of cells stained with PI (red–orange) divided by the number of cells stained with PI plus cells stained with AO (green). 2.8. Scanning and transmission electron microscopy SEM images were obtained with a JEOL JSM-6380LA and Zeiss Leo 1530 microscope operating at 5 kV. Following incubation with bacteria, samples of NFSiC and NRSiC were placed on an aluminum plate and coated with gold. Then, samples were viewed by SEM. Pure SiC nanostructures were observed without coating.
to identify groups of bacteria. In the investigated microbial cultures, only the bacterium P. putida was found. 3.1. Peroxidase activity Peroxidase activity was measured using a colorimetric method, in which the product of the oxidation of ABTS is green–blue, as presented in Fig. 2. The color intensity was measured at 420 nm using a spectrophotometer. As a control, a culture without SiC (abiotic in the diagram) was used. The results showed that only nanostructural SiC had an influence on the P. putida culture (lmSiC did not affect the bacteria). In the case of the abiotic system, statistically significant differences were not observed. Only a slight increase in the absorption level was found, probably due to the presence of reactive forms of oxygen on the surface of the nanostructures. Nanostructures with a very thin and amorphous layer of silica dioxide had slightly higher levels of absorbance. Silica nanofilms on nanofibers and nanorods can promote the formation of highly reactive hydroxyl radicals and singlet oxygen. These highly reactive forms are reported to be responsible for hydroxyl radical-mediated DNA damage in cells. The toxic effect of silica nanomaterials was reported in relation to both bacterial and mammalian cells (Jiang et al., 2009; Yu et al., 2011). It is very likely that the SiC nanostructures can, as a result of mechanical cell damage or the presence of reactive forms of oxygen on the surface of the NRSiC and NFSiC, cause P. putida to increase its production of enzymes that protect the cell against reactive forms of oxygen. 3.2. Estimation of oxidative stress in plasmid DNA after digestion with the Fpg protein The presence of reactive forms of oxygen can damage nucleic acids, such as the oxidization of base pairs in plasmids. This kind of damage can be repaired by the Fpg protein. Fpg has monofunctional glycosylase and AP endonuclease activity. The damaged nucleotide base is recognized and cleaved. As a result, changes in the topology of the plasmid DNA can be observed, from CCC (covalently closed circular) to OC (open circular), thereby enabling the measurement of DNA damage as a consequence of oxidation stress. Nanostructures may induce oxidative stress in bacteria and stop DNA synthesis (they were observed as the conversion of topological DNA forms by the repair enzyme). 8-oxoG present in the DNA, a common bacterial marker of oxidative stress, was converted into DNA strand breaks by digestion of the modified plasmid with Fpg.
biotic 2.9. Statistical analysis The obtained data (activity of peroxidase test and viability test) were analyzed for the significance of mean differences using oneway analyses of variance (ANOVA) at p < 0.05. Post-hoc tests for pair-wise differences and the identification of homogeneous subgroups were conducted using Tukey’s HSD procedure. Homogenous subgroups are indicated by diagrams with the same small letters. The ANOVA was computed with Statistica 10 software (StatSoft Inc., Tulsa, OK, USA).
Absorbance
abiotic
d
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0.40 0.30
c
bc
0.20 0.10
ab
ab
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a
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0.00
3. Results and discussion We first verified whether the cultures, after incubation with the investigated materials, were contaminated with other bacteria. Therefore, DNA was isolated and the 16S rRNA gene was amplified by PCR. Furthermore, the product of amplification was sequenced
Control
NRSiC
Fig. 2. Peroxidase activity in extracts from bacterial cells after incubation of cultures with the investigated materials (biotic). Peroxidase activity test in sterile media containing the investigated materials (abiotic). The same letters indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
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3.3. Expression of the zwf1 gene – real-time PCR and northern blot analysis
NFSiC F
NFSiC
NRSiC F
NRSiC
µmSiC F
µmSiC
control F
control
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Northern blot analysis revealed that the expression of the zwf1 gene was the highest in the NFSiC and NRSiC samples. The lowest gene expression was found in the control treated with lmSiC (Fig. 4). Gene expression was investigated relative to that of the 16S rDNA genes, which revealed that the expression of the zwf1 gene was extremely high. The zwf1 gene is one of the genes that are crucial for oxidative stress protection in yeast and bacteria cells (Sung and Lee, 2007). zwf1 encodes glucose-6-phosphate dehydrogenase (G6PD). G6PD is a major source of electrons, and it combats oxidative stress in cells. G6PD catalyzes a step in the pentose phosphate pathway, which ultimately provides the cell with NADPH. NADPH is essential in protecting the cell from oxygen (Giro‘ et al., 2006). Deletion of the zwf1 gene increases bacterial and yeast sensitivity to oxidative stresses (Giro‘ et al., 2006). The zwf1 gene is activated by the SoxRS transcriptional activator in Escherichia coli (Demple, 1996). The zwf1 gene of P. putida KT2440 is strongly induced by different sources of oxidative stress reagents, such as nitric oxide, menadione, and paraquat (Park et al., 2006).
Fig. 3. Isolation of undigested and Fpg-digested (marked by an ‘‘F’’) plasmid DNA isolated from cultures treated with the investigated materials. In the control and lmSiC samples, two forms of plasmid DNA are visible – the lower band is a covalently closed circular (CCC) form, and the upper band is an open circular (OC) form. In the case of digested plasmid DNA the smears resulting from silicon carbide nanorods (NRSiC F), nanofibers (NFSiC F), and, partly, for micrometric SiC (lmSiC F), are a visible effect of DNA damage after oxidative stress.
Relative expression of zwf1 gene normalized to 16S rRNA gene
As a qualitative research method for detecting DNA damage, hydrolysis with Fpg can be used. Fig. 3 shows the results of the electrophoresis of plasmid DNA isolated from tested cultures (control, lmSiC, NRSiC, NFSiC) and plasmid DNA after hydrolysis with Fpg (control F, lmSiC F, NRSiC F, NFSiC F). The results of Fpg hydrolysis show a large number of DNA fragments; larger DNA fragments indicate greater damage to the DNA, which can be observed as smears of DNA in an agarose gel. In the control, DNA hydrolysis with Fpg did not show such smears, as only a distinct band of DNA was found. In other samples, Fpg hydrolysis resulted in smears – very faint smears were observed in the sample after bacterial incubation with lmSiC, and the most intense smears were found in the sample incubated with NFSiC, which is evidence of a large amount of DNA damage.
3.4. Loss of viability The previous results showed that SiC nanostructures damage DNA in living cells. Additionally, the expression of the zwf1 gene confirmed that the DNA damage likely resulted from oxidative stress. Therefore, we investigated how NFSiC and NRSiC can affect P. putida cell viability. The attachment of living cells to SiC material could disrupt membrane integrity, thus leading to cell death. In Fig. 5, the results of the loss of cell viability in response to SiC nanostructure treatment are shown. Bacteria stained with PI may reflect cell membrane damage; such cells appear red or orange– red, depending on the wavelength of excitation used in the microscope. Furthermore, staining with AO, or another dye proposed by the authors (e.g., DAPI) (Kang et al., 2007), allows the differentiation of dead cells from viable cells. The results showed that presence of nanostructural SiC material can cause a significant loss of viability of living cells. In response to the NFSiC and NRSiC treatments, approximately 90% of cells that adsorbed to the surface of the tested materials were damaged or dead. The population of living cells not adsorbed on the tested materials also showed a loss of viability, 45% for NFSiC and 56%
e
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B
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- expression of zwf1 gene - expression of 16S rRNA gene Fig. 4. A – Real-time PCR analysis of the expression of the zwf1 gene. B – Northern blot analysis showing the intensity of zwf1 mRNA. The same letters indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
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loss of viability [%]
e
e
100
free living 80
on surface
c
d
60 40 20
b
b
a
0
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µmSiC
NFSiC
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Fig. 5. The loss of viability (%) of free-living cells, and cells adsorbed onto the surface of aggregates of silicon carbide (SiC) nanostructures. The same letters in the diagram indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
for NRSiC. In the control and lmSiC-treated samples, the loss of viability was less than 20% of all cells. In the SEM images, bacteria associated with the NFSiC and NRSiC are visible (Fig. 6). Many recent studies have focused on the antimicrobial activity of carbon nanostructures, such as single- and multi-walled nanotubes (Kang et al., 2007, 2008a,b; Akhavan et al., 2011; Su et al., 2013), or other carbon materials, such as graphite, graphene, and fullerene (Akhavan and Ghaderi, 2010; Liu et al., 2011; Lyon et al., 2006). The mechanism proposed to explain the antibacterial
properties of these materials is primarily based on oxidative stress and physical interactions with the cell membrane (Liu et al., 2011). Physical interactions with the cell membrane leading to a loss of integrity have been proposed as the key antibacterial mechanism of carbon nanotubes, and it is possible that the same mechanism could play a crucial role in NFSiC and NRSiC. Inherent chemical toxicity to bacteria is unlikely because bulk SiC is, in general, chemically stable and biocompatible. In this study, lmSiC also seemed to be non-toxic to bacteria. However, silica nanofilms on nanofibers and nanorods SiC can promote the formation of highly reactive hydroxyl radicals and singlet oxygen. These highly reactive forms are reported to be responsible for hydroxyl radical-mediated DNA damages in cells. Catalytic properties of b-SiC nanofibers during formation of reactive oxygen species was studied by Svensson et al. (1997). They demonstrated that silicon carbide nanofibers with average length about 12 lm six-times increase the concentration of hydroxyl radicals and hydrogen peroxide in relation to control without nanostructured SiC. Pourchez et al. (2012) showed that concentration of hydrogen peroxide generated in solutions in presence of nanometric b-SiC is 20–100 times higher than in the culture without nanomaterial. The Authors also showed that the concentration of hydroxyl radicals and carboxylic radical ion depends on the shape of b-SiC nanostructures, the degree of crystallinity and surface oxidation of fibers. In our paper we examined the b-SiC (confirmed by XRD method) which are coated with thin layer of amorphous silica (confirmed by TEM technique). In the context of Svensson et al. (1997) and Pourchez et al. (2012) results, one of the reasons for toxicity of SiC nanofibers tested in this study was the generation of reactive oxygen radicals and radical ions catalyzed by silicon carbide nanofibers.
4. Conclusion Our work has provided the first evidence that NFSiC and NRSiC, produced by combustion synthesis, can induce oxidative stress in P. putida. Based on the results presented in the paper, it can be stated that nanostructured SiC may affect the activity of microorganisms by increasing the formation of highly reactive radicals and disrupting cell membrane integrity. These mechanisms of cytotoxicity can lead to DNA damage in bacterial cells, and they can affect the expression of genes involved in the suppression of oxidative stress. Therefore, we analyzed the expression of the zwf-1 gene. SiC can affect bacterial activity, but this process is controlled by the texture of the material. The lmSiC used in this study as a reference material did not significantly influence the activity of P. putida. On the contrary, the nanostructured SiC seemed to affect the expression of oxidative stress genes and cell viability. Thus, the bacteria were likely undergoing increased cytotoxicity. However, it cannot be clearly stated whether this process led to cell death or only to cell damage. In addition to the physical interaction of SiC with the cell membrane, it is possible that the toxicity of nanostructured SiC is connected with the presence of silica nanofilms on the surface of the investigated materials.
Acknowledgments
Fig. 6. P. putida bacteria associated with silicon carbide nanofibers (NFSiC) (A) and nanorods (NRSiC) (B) after incubation with the investigated materials.
This research was partially supported by the European Union, within the European Regional Development Fund, through the Innovative Economy grant (POIG.02.02.00-00-025/09) and by the Faculty of Geology, University of Warsaw, BST 166901/2013. The authors thank reviewers for their critical remarks and comments which have improved this article.
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Kang, S., Herzberg, M., Rodrigues, D.F., Elimelech, M., 2008a. Antibacterial effects of carbon nanotubes: size does matter! Langmuir 24, 6409–6413. Kang, S., Mauter, M.S., Elimelech, M., 2008b. Physicochemical determinants of multiwalled carbon nanotube bacterial cytotoxicity. Environ. Sci. Technol. 42, 7528–7534. Kim, J., Jeon, C.O., Park, W., 2008. Dual regulation of zwf-1 by both 2-keto-3-deoxy6-phosphogluconate and oxidative stress in Pseudomonas putida. Microbiology 154, 3905–3916. Liu, S., Zeng, T.H., Hofmann, M., Burcombe, E., Wei, J., Jiang, R., Kong, J., Chen, Y., 2011. Antibacterial activity of graphite, graphite oxide, graphene oxide, and reduced graphene oxide: membrane and oxidative stress. ACS Nano 5, 6971– 6980. Lyon, D.Y., Adams, L.K., Falkner, J.C., Alvarez, P.J.J., 2006. Antibacterial activity of fullerene water suspensions: effects of preparation method and particle size. Environ. Sci. Technol. 40, 4360–4366. Park, W., Peña-Llopis, S., Lee, Y., Demple, B., 2006. Regulation of superoxide stress in Pseudomonas putida KT2440 is different from SoxR paradigm in Escherichia coli. Biochem. Biophys. Res. Commun. 341, 51–56. Pourchez, J., Forest, V., Boumahdi, N., Boudard, D., Tomatis, M., Fubini, B., HerlinBoime, N., Leconte, Y., Guilhot, B., Cottier, M., Grosseau, P., 2012. In vitro cellular responses to silicon carbide nanoparticles: impact of physic-chemical features on pro-inflammatory and pro-oxidative effects. J. Nanopart. Res. 14 (10), 1143– 1155. Su, R., Jin, Y., Tong, M., Kim, H., 2013. Bactericidal activity of Ag-doped multi-walled carbon nanotubes and the effects of extracellular polymeric substances and natural organic matter. Colloids Surf. B 104, 133–139. Sung, J.Y., Lee, Y.N., 2007. Isoforms of glucose 6-phosphate dehydrogenase in Deinococcus radiophilus. J. Microbiol. 45, 318–325. Svensson, I., Artursson, E., Leanderson, P., Berglind, R., Lindgren, F., 1997. Toxicity in vitro of some silicon carbides and silicon nitrides: whiskers and powders. Am. J. Ind. Med. 31 (3), 335–343. Szala, M., Borkowski, A., 2014. Toxicity assessment of SiC nanofibers and nanorods against bacteria. Ecotoxicol. Environ. Saf. 100, 287–293. Yu, T., Malugin, A., Ghandehari, H., 2011. Impact of silica nanoparticle design on cellular toxicity and hemolytic activity. ACS Nano 5, 5717–5728.
Contents lists available at ScienceDirect
Chemosphere journal homepage: www.elsevier.com/locate/chemosphere
Oxidative stress in bacteria (Pseudomonas putida) exposed to nanostructures of silicon carbide _ e, Andrzej Borkowski a,⇑, Mateusz Szala b, Paweł Kowalczyk c, Tomasz Cłapa d, Dorota Narozna Marek Selwet d _ i Wigury 93, 02-089 Warsaw, Poland Faculty of Geology, University of Warsaw, Zwirki Faculty of Advanced Technologies and Chemistry, Military University of Technology, Kaliskiego 2, 00-908 Warsaw, Poland c Bionicum Ltd., Chełmska 21, Warsaw, Poland d Department of General and Environmental Microbiology, Poznan University of Life Sciences, Szydłowska 50, 60-656 Poznan, Poland e Department of Biochemistry and Biotechnology, Poznan University of Life Sciences, Dojazd 11, 60-632 Poznan, Poland a
b
h i g h l i g h t s
g r a p h i c a l a b s t r a c t
Silicon carbide nanostructures were
produced by combustion synthesis. Oxidative stress in Pseudomonas
putida was investigated. Peroxidase activity, expression of zwf-
1 gene and cell viability were analyzed. SiC nanofibers and nanorods can induce oxidative stress in P. putida.
a r t i c l e
i n f o
Article history: Received 12 December 2014 Received in revised form 10 April 2015 Accepted 17 April 2015 Available online 15 May 2015 Keywords: Silicon carbide Nanofibers Nanorods Oxidative stress Pseudomonas putida Toxicity
a b s t r a c t Silicon carbide (SiC) nanostructures produced by combustion synthesis can cause oxidative stress in the bacterium Pseudomonas putida. The results of this study showed that SiC nanostructures damaged the cell membrane, which can lead to oxidative stress in living cells and to the loss of cell viability. As a reference, micrometric SiC was also used, which did not exhibit toxicity toward cells. Oxidative stress was studied by analyzing the activity of peroxidases, and the expression of the glucose-6-phosphate dehydrogenase gene (zwf1) using real-time PCR and northern blot techniques. Damage to nucleic acid was studied by isolating and hydrolyzing plasmids with the formamidopyrimidine [fapy]-DNA glycosylase (also known as 8-oxoguanine DNA glycosylase) (Fpg), which is able to detect damaged DNA. The level of viable microbial cells was investigated by propidium iodide and acridine orange staining. Ó 2015 Elsevier Ltd. All rights reserved.
1. Introduction
⇑ Corresponding author. E-mail address: [email protected] (A. Borkowski). http://dx.doi.org/10.1016/j.chemosphere.2015.04.066 0045-6535/Ó 2015 Elsevier Ltd. All rights reserved.
A review of the literature shows that nanomaterials can be cytoand genotoxic to prokaryotic and eukaryotic cells. Most of the tests on nanostructures were run with one- and two dimension
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nanomaterials: carbon nanotubes, metal nanoparticles, graphene nanoplates, and graphene. Genotoxicity of graphene nanoplates and graphene nanoribbons toward human cells was reported by Akhavan et al. (2012, 2013). There are few studies on silicon carbide (SiC) nanostructures, which can be considered biocompatible and chemically non-active. Szala and Borkowski (2014) reported that SiC nanofibers and nanorods can be cytotoxic to bacteria by mechanically damaging their cell membrane. The toxicological data on mammalian cells in the literature were mainly obtained from experiments conducted on microcrystalline SiC particles or fibers. It has been shown that SiC powder does not induce harmful effects on tissues, which led to the view that microcrystalline SiC material is biologically inert (Bruch et al., 1993a,b). A few studies have described possible toxic effects of exposure of mammalian cells to SiC nanostructures in vitro (Barillet et al., 2010; Pourchez et al., 2012). There are two basic mechanisms of nanostructure toxicity toward living organisms: (i) mechanical damage of the cell membrane and (ii) oxidative stress, which can be caused by mechanical damage of the cell membrane or the presence of reactive forms of oxygen. Reactive forms of oxygen can be present on the surface of nanomaterials because of the high catalytic activity of the materials and their large specific surface areas. In a previous study, SiC nanostructures were covered with a thin layer (thickness of approximately 2 nm) of partially amorphous SiO2 (Huczko et al., 2005). Silica nanofilms can promote the formation of highly reactive hydroxyl radicals and singlet oxygen, which are responsible for hydroxyl radical-mediated DNA damage (Cadet et al., 1999). The toxic effect of silica nanomaterials was reported in relation to both bacteria and mammalian cells (Jiang et al., 2009; Yu et al., 2011). The aim of this study was to determine the dependence between the textures of SiC nanostructures and their ability to cause oxidative stress in Pseudomonas putida. Therefore, bacteria were treated with SiC nanostructures (nanofibers and nanorods, (NFSiC and NRSiC respectively)) and micrometric SiC (lmSiC) as a control. The results show that lmSiC was not toxic to the cells, and that it can be used as a reference material.
(TEM) (Huczko et al., 2005). The results clearly show that NFSiC and NRSiC were covered with a thin layer of silica. SEM images of the investigated materials are presented in Fig. 1. 2.2. Microorganisms and media The strain of P. putida was obtained from our own collection of isolated microorganisms from samples of organic soils (Geomicrobiology Laboratory, Faculty of Geology, University of Warsaw). Taxonomic affiliation was confirmed by sequencing
2. Materials and methods 2.1. Nanomaterials and lmSiC NFSiC were prepared by a combustion route. The combustion mixture was prepared by dry mixing powders of calcium disilicide (CaSi2) and poly(tetrafluoroethene) (PTFE) in a ceramic mortar. After pressing the powders into a cylindrical pellet, 5 g of the sample were placed in a graphite crucible and placed in a stainless steel autoclave (350 cm3 in volume), which was subsequently filled with helium at an initial pressure of 1.0 MPa. The combustion process was initiated with an electrically-heated resistance wire (diameter 0.1 mm). The spongy combustion products were removed from the autoclave with water. The suspension was filtered, and the gray deposit obtained was purified in a three-step process: heating in 98% H2SO4, calcination in air (700 °C), and, finally, heating in 25% NaOH and washing with plenty of water. The NFSiC prepared using this method was used in all experiments. NRSiC were also prepared by the combustion route, but the starting mixture comprised fluorinated graphite (CF) and aluminum–silicon (AlSi). Purification was conducted following the same procedure, but the step of heating in 98% H2SO4 was omitted. lmSiC was purchased from Sigma– Aldrich (St. Louis, MO, USA) and used without purification. Nanomaterials were characterized using X-ray powder diffraction (XRD), Raman spectroscopy, elemental analysis, scanning electron microscopy (SEM) and transmission electron microscopy
Fig. 1. Scanning electron microscopy (SEM) images of silicon carbide nanofibers (NFSiC) (A), nanorods NRSiC (B), and micrometric SiC (lmSiC) used as a reference material (C).
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analysis of the 16S rDNA gene. The bacteria were cultivated in liquid and solid nutrient media (pH 7.5) comprising the following (g/ L): glucose, 10; peptone, 5; yeast extract, 2; NaCl, 4; and agar (in the case of solid medium), 20. The media were autoclaved at 121 °C for 15 min.
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was kept at 1.8–2.0. The DNA solution was stored at 20 °C until further use. The quality of the reaction products were analyzed electrophoretically in 1% agarose gels as the ratio of the covalently closed circular to the open circular form of the plasmid. 2.6. Expression of the zwf1 gene (RNA analysis)
2.3. Cultures of P. putida with investigated SiC materials Forty milligrams of lmSiC, NFSiC, or NRSiC were added to 10 mL of sterile liquid medium in a sterile glass bottle (Simax, 100 mL), and 0.5 mL of P. putida inoculum was added to the mixture. The obtained suspension was incubated at 25 °C under constant stirring (200 rpm) for 24 h. The inoculum was prepared as follows: P. putida was cultured on nutrient agar plates for 24 h at 30 °C. Then, the cultured bacteria were added to 5 mL of NaCl solution (0.9%) to a concentration of approximately 108 colony-forming units (cfu)/mL. After incubation with investigated materials, the peroxidase activity of the obtained bacterial cultures was analyzed, and bacterial DNA and RNA were isolated. 2.4. Assay of peroxidase activity in bacterial cells Peroxidase activity was determined in extracts from bacterial cells grown in liquid medium cultures. At defined times during the bacterial growth culture, cells were harvested by centrifugation and resuspended in 50 mM phosphate buffer (pH 7.8) and sonicated. Sonicates were centrifuged at 13,000 rpm (26,000g) for 30 min, with 10 s intervals to obtain the supernatant, which was used in the assay. The peroxidase activity was measured spectrophotometrically at 420 nm using 50 mM ABTS [2,2-azinobis(3-ethylbenzothiazoline-6-sulfonic acid)] solution in 0.5 M phosphate-citrate buffer (pH 4.0). Measurement of peroxidase activity was performed after incubating for 60 min at 37 °C in a cell counter (Thermo Scientific, Waltham, MA, USA). The colorless ABTS molecule is converted into the blue–green radical, ABTS+, via the oxidation of one electron in the presence of antioxidants in the sample. Additionally, the activity of antioxidants in the samples with lmSiC, NRSiC, or NFSiC without bacteria was measured in the same way as described above. This test was conducted to assess whether the investigated SiC materials can interfere with the measurements of peroxidase activity. Negative control contained buffer without SiC. 2.5. Estimation of oxidative stress in plasmid DNA after digestion with the Fpg protein Plasmids were extracted from cultures: without SiC – control (K), and with lmSiC, NFSiC, or NRSiC using the plasmid mini prep isolation kit (A&A Biotechnology, Gdynia, Poland) according to the manufacturer’s instructions. The purity and concentration of the DNA preparation were determined spectrophotometrically at 260 nm. Next, the obtained samples of DNA were digested with the Fpg protein. The standard reaction mixture (final volume of 20 lL) for the Fpg protein contained 10 lg of plasmid DNA, 0.09 lg of Fpg, 100 mM HEPES-KOH, pH 7.6, 150 mM KCl, 10 mM EDTA, 5 mM MgCl2, 1 mM b-mercaptoethanol, and 100 lg/mL of bovine serum albumin (BSA), and it was incubated for 30 min at 37 °C. After DNA cleavage, the enzyme was removed by chloroform extraction and DNA was precipitated with 4 volumes of cold 96% ethanol and 0.1 volume of 3 M sodium acetate (pH 5.2) at 20 °C overnight, or at 80 °C for 2 h, to sediment all small DNA fragments, and subsequently centrifuged at 12,000 rpm (22,000g) for 15 min. The DNA pellet was resuspended in water, and the DNA concentration was measured using a Varian Cary 3E spectrophotometer with ADL News software (Varian, Sunnyvale, CA, USA). To ensure the complete removal of protein, the A260/A280 ratio
2.6.1. Sample preparation for real-time PCR For this analysis, total RNA was isolated from bacterial cells using Trizol (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. The concentration of total RNA was calculated from its absorbance at 260 nm. Total RNA was used to synthesize cDNA using Advantage kit RTfor-PCR (Clontech Laboratories, Inc., Mountain View, CA, USA) and oligo (dT) primers according to the manufacturer’s instructions. One microgram of total RNA was used as the template, to which was added 1 lL of oligo (dT) primers (2 ng), 1 lL of dNTPs (10 mM final concentration), 4 lL of 5 reaction buffer, and 0.5 lL of RNase. RNA and oligo primers were gently mixed together and heated for 2 min at 70 °C. Then, they were placed on ice and mixed with the other reaction components. Subsequently, 1 lL of reverse transcriptase MMLV (20 U/lL) was added, and the samples were incubated for 60 min at 37 °C. MMLV was inactivated by heating at 95 °C for 5 min. 2.6.2. Real-time PCR with SYBR green An ABI Prism 7500 (Applied Biosystems, Foster City, CA, USA) was used for the analysis. Twenty-five microliters of reaction mixture contained 1 Taq polymerase buffer, 40,000 diluted SYBR green, 3 mM MgCl2, 80 nM of each primer, 400 nM dATP, dGTP, dCTP, and dTTP, and 0.025 U/lL of Taq polymerase. Six hundred forty or 160 ng of cDNA was added to each reaction. To examine the constitutive level of gene expression in the tested bacterial samples, the Ct values of 16S rRNA and zwf1 were investigated. The Ct value is the number of cycles at which the generated fluorescence in the reaction reaches a threshold value, which occurs in the linear stage of the amplification profile. All reactions were performed in two replicates with 640 ng of template and in two replications with 160 ng of template, as well as with a negative control without template. The PCR conditions were as follows: an initial denaturation at 95 °C for 3 min, followed by 45 cycles of 95 °C for 15 s (denaturation), 60 °C for 15 s (annealing), 72 °C for 15 s (extension), and 72 °C for 1 min (final extension). The fluorescence was detected in each cycle after every extension phase. To differentiate specific products from non-specific products, the dissociation curve was created. The Ct values were calculated by SDS v.1.7 software (Applied Biosystems) with the automatic basic curve, which was the median fluorescence value from 3 to 15 cycles. The data were exported to the Microsoft Excel program for further analyses. The control curve was created for the 16S rRNA and zwf-1 genes using a series of dilutions of reference RNAs. The level of transcription of mRNA was normalized to that of the 16S rRNA in each dilution. The standard curve was created based on the normalized transcription level of the initial logarithmic concentration of cDNA in each dilution. As the reference, the median Ct values from three samples were used. The gene ratio was calculated using the equation: ratio = 2 ddCt, where ddCt is the difference between the Ct of the investigated gene samples and the Ct of the reference samples. Additionally, RNA was measured by northern blotting. RNA was isolated from 3 mL of exponentially growing cells using the RNeasy kit (Qiagen) according to the manufacturer’s instructions. RNA was quantified by measuring the optical absorbance at 260 nm (A260) of 4–5 mg of total RNA per sample. Fractionated RNA was transferred to a nylon membrane (GE Healthcare, Little Chalfont,
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United Kingdom) using a Turboblotter (Schleicher & Schuell), and the amounts of zwf1 mRNA were determined by hybridizing the membrane with an a-32P-labelled probe specific for each gene (TaKaRa Bio). To produce probes with high specific activities, the following primers were used: zwf1 pro1 CGCGAATTCCGAGTCGCAGCGTTTTGTCT and zwf1 pro2 CGCGGATCCCCAGCAGGCGGGTGTCC (Kim et al., 2008). The results were compared with 16S rRNA gene sequences, which were a control for the zwf1 gene. 2.7. Viability test To analyze the loss of viability, solutions of propidium iodide (PI) (2 mg/0.1 L, pH = 7.4) and acridine orange (AO) (5 mg/0.1 L, pH = 7.4) were prepared in phosphate buffer. Tests were conducted as follows: in a 100-mL sterile glass bottle (Simax), 10 mL of sterile saline solution (0.9% NaCl) and 40 mg of lmSiC, NFSiC, or NRSiC were added. Subsequently, 1 mL of P. putida inoculum (approximately 108 cfu/mL in 0.9% NaCl) was added to the mixture and mixed (with shaking at 200 rpm) for 120 min at 25 °C. After mixing, 1 mL of suspension was placed into an Eppendorf tube, and 0.3 mL of sucrose (60%) was added. The mixture was centrifuged for 2 min at 4000 rpm (2600g). Sucrose was added to separate unadsorbed bacteria from the investigated nanostructures that had adsorbed to cells. Then, the supernatant (containing free-living cells) and residuum (NFSiC or NRSiC with adsorbed cells) were stained as follows. Initially, 0.2 mL of the supernatant or residuum, and 30 lL of PI were added to a test tube and left for 10 min in the dark. Later, 15 lL of AO was added (staining for 2 min) to the test tube. After staining, 10 lL of suspension was placed on a glass slide and covered with a coverslip. Ten representative fluorescence images of free-living cells and cells adsorbed onto the surface of aggregates of SiC nanostructures were acquired using an epifluorescence microscope with a B-filter. Dead cells appeared red–orange, while viable cells were green. The result of the microscopic analysis was expressed as the ratio of the number of cells stained with PI (red–orange) divided by the number of cells stained with PI plus cells stained with AO (green). 2.8. Scanning and transmission electron microscopy SEM images were obtained with a JEOL JSM-6380LA and Zeiss Leo 1530 microscope operating at 5 kV. Following incubation with bacteria, samples of NFSiC and NRSiC were placed on an aluminum plate and coated with gold. Then, samples were viewed by SEM. Pure SiC nanostructures were observed without coating.
to identify groups of bacteria. In the investigated microbial cultures, only the bacterium P. putida was found. 3.1. Peroxidase activity Peroxidase activity was measured using a colorimetric method, in which the product of the oxidation of ABTS is green–blue, as presented in Fig. 2. The color intensity was measured at 420 nm using a spectrophotometer. As a control, a culture without SiC (abiotic in the diagram) was used. The results showed that only nanostructural SiC had an influence on the P. putida culture (lmSiC did not affect the bacteria). In the case of the abiotic system, statistically significant differences were not observed. Only a slight increase in the absorption level was found, probably due to the presence of reactive forms of oxygen on the surface of the nanostructures. Nanostructures with a very thin and amorphous layer of silica dioxide had slightly higher levels of absorbance. Silica nanofilms on nanofibers and nanorods can promote the formation of highly reactive hydroxyl radicals and singlet oxygen. These highly reactive forms are reported to be responsible for hydroxyl radical-mediated DNA damage in cells. The toxic effect of silica nanomaterials was reported in relation to both bacterial and mammalian cells (Jiang et al., 2009; Yu et al., 2011). It is very likely that the SiC nanostructures can, as a result of mechanical cell damage or the presence of reactive forms of oxygen on the surface of the NRSiC and NFSiC, cause P. putida to increase its production of enzymes that protect the cell against reactive forms of oxygen. 3.2. Estimation of oxidative stress in plasmid DNA after digestion with the Fpg protein The presence of reactive forms of oxygen can damage nucleic acids, such as the oxidization of base pairs in plasmids. This kind of damage can be repaired by the Fpg protein. Fpg has monofunctional glycosylase and AP endonuclease activity. The damaged nucleotide base is recognized and cleaved. As a result, changes in the topology of the plasmid DNA can be observed, from CCC (covalently closed circular) to OC (open circular), thereby enabling the measurement of DNA damage as a consequence of oxidation stress. Nanostructures may induce oxidative stress in bacteria and stop DNA synthesis (they were observed as the conversion of topological DNA forms by the repair enzyme). 8-oxoG present in the DNA, a common bacterial marker of oxidative stress, was converted into DNA strand breaks by digestion of the modified plasmid with Fpg.
biotic 2.9. Statistical analysis The obtained data (activity of peroxidase test and viability test) were analyzed for the significance of mean differences using oneway analyses of variance (ANOVA) at p < 0.05. Post-hoc tests for pair-wise differences and the identification of homogeneous subgroups were conducted using Tukey’s HSD procedure. Homogenous subgroups are indicated by diagrams with the same small letters. The ANOVA was computed with Statistica 10 software (StatSoft Inc., Tulsa, OK, USA).
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3. Results and discussion We first verified whether the cultures, after incubation with the investigated materials, were contaminated with other bacteria. Therefore, DNA was isolated and the 16S rRNA gene was amplified by PCR. Furthermore, the product of amplification was sequenced
Control
NRSiC
Fig. 2. Peroxidase activity in extracts from bacterial cells after incubation of cultures with the investigated materials (biotic). Peroxidase activity test in sterile media containing the investigated materials (abiotic). The same letters indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
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3.3. Expression of the zwf1 gene – real-time PCR and northern blot analysis
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Northern blot analysis revealed that the expression of the zwf1 gene was the highest in the NFSiC and NRSiC samples. The lowest gene expression was found in the control treated with lmSiC (Fig. 4). Gene expression was investigated relative to that of the 16S rDNA genes, which revealed that the expression of the zwf1 gene was extremely high. The zwf1 gene is one of the genes that are crucial for oxidative stress protection in yeast and bacteria cells (Sung and Lee, 2007). zwf1 encodes glucose-6-phosphate dehydrogenase (G6PD). G6PD is a major source of electrons, and it combats oxidative stress in cells. G6PD catalyzes a step in the pentose phosphate pathway, which ultimately provides the cell with NADPH. NADPH is essential in protecting the cell from oxygen (Giro‘ et al., 2006). Deletion of the zwf1 gene increases bacterial and yeast sensitivity to oxidative stresses (Giro‘ et al., 2006). The zwf1 gene is activated by the SoxRS transcriptional activator in Escherichia coli (Demple, 1996). The zwf1 gene of P. putida KT2440 is strongly induced by different sources of oxidative stress reagents, such as nitric oxide, menadione, and paraquat (Park et al., 2006).
Fig. 3. Isolation of undigested and Fpg-digested (marked by an ‘‘F’’) plasmid DNA isolated from cultures treated with the investigated materials. In the control and lmSiC samples, two forms of plasmid DNA are visible – the lower band is a covalently closed circular (CCC) form, and the upper band is an open circular (OC) form. In the case of digested plasmid DNA the smears resulting from silicon carbide nanorods (NRSiC F), nanofibers (NFSiC F), and, partly, for micrometric SiC (lmSiC F), are a visible effect of DNA damage after oxidative stress.
Relative expression of zwf1 gene normalized to 16S rRNA gene
As a qualitative research method for detecting DNA damage, hydrolysis with Fpg can be used. Fig. 3 shows the results of the electrophoresis of plasmid DNA isolated from tested cultures (control, lmSiC, NRSiC, NFSiC) and plasmid DNA after hydrolysis with Fpg (control F, lmSiC F, NRSiC F, NFSiC F). The results of Fpg hydrolysis show a large number of DNA fragments; larger DNA fragments indicate greater damage to the DNA, which can be observed as smears of DNA in an agarose gel. In the control, DNA hydrolysis with Fpg did not show such smears, as only a distinct band of DNA was found. In other samples, Fpg hydrolysis resulted in smears – very faint smears were observed in the sample after bacterial incubation with lmSiC, and the most intense smears were found in the sample incubated with NFSiC, which is evidence of a large amount of DNA damage.
3.4. Loss of viability The previous results showed that SiC nanostructures damage DNA in living cells. Additionally, the expression of the zwf1 gene confirmed that the DNA damage likely resulted from oxidative stress. Therefore, we investigated how NFSiC and NRSiC can affect P. putida cell viability. The attachment of living cells to SiC material could disrupt membrane integrity, thus leading to cell death. In Fig. 5, the results of the loss of cell viability in response to SiC nanostructure treatment are shown. Bacteria stained with PI may reflect cell membrane damage; such cells appear red or orange– red, depending on the wavelength of excitation used in the microscope. Furthermore, staining with AO, or another dye proposed by the authors (e.g., DAPI) (Kang et al., 2007), allows the differentiation of dead cells from viable cells. The results showed that presence of nanostructural SiC material can cause a significant loss of viability of living cells. In response to the NFSiC and NRSiC treatments, approximately 90% of cells that adsorbed to the surface of the tested materials were damaged or dead. The population of living cells not adsorbed on the tested materials also showed a loss of viability, 45% for NFSiC and 56%
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- expression of zwf1 gene - expression of 16S rRNA gene Fig. 4. A – Real-time PCR analysis of the expression of the zwf1 gene. B – Northern blot analysis showing the intensity of zwf1 mRNA. The same letters indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
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loss of viability [%]
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Fig. 5. The loss of viability (%) of free-living cells, and cells adsorbed onto the surface of aggregates of silicon carbide (SiC) nanostructures. The same letters in the diagram indicate values that do not differ significantly at p < 0.05. Error bars indicate standard deviations.
for NRSiC. In the control and lmSiC-treated samples, the loss of viability was less than 20% of all cells. In the SEM images, bacteria associated with the NFSiC and NRSiC are visible (Fig. 6). Many recent studies have focused on the antimicrobial activity of carbon nanostructures, such as single- and multi-walled nanotubes (Kang et al., 2007, 2008a,b; Akhavan et al., 2011; Su et al., 2013), or other carbon materials, such as graphite, graphene, and fullerene (Akhavan and Ghaderi, 2010; Liu et al., 2011; Lyon et al., 2006). The mechanism proposed to explain the antibacterial
properties of these materials is primarily based on oxidative stress and physical interactions with the cell membrane (Liu et al., 2011). Physical interactions with the cell membrane leading to a loss of integrity have been proposed as the key antibacterial mechanism of carbon nanotubes, and it is possible that the same mechanism could play a crucial role in NFSiC and NRSiC. Inherent chemical toxicity to bacteria is unlikely because bulk SiC is, in general, chemically stable and biocompatible. In this study, lmSiC also seemed to be non-toxic to bacteria. However, silica nanofilms on nanofibers and nanorods SiC can promote the formation of highly reactive hydroxyl radicals and singlet oxygen. These highly reactive forms are reported to be responsible for hydroxyl radical-mediated DNA damages in cells. Catalytic properties of b-SiC nanofibers during formation of reactive oxygen species was studied by Svensson et al. (1997). They demonstrated that silicon carbide nanofibers with average length about 12 lm six-times increase the concentration of hydroxyl radicals and hydrogen peroxide in relation to control without nanostructured SiC. Pourchez et al. (2012) showed that concentration of hydrogen peroxide generated in solutions in presence of nanometric b-SiC is 20–100 times higher than in the culture without nanomaterial. The Authors also showed that the concentration of hydroxyl radicals and carboxylic radical ion depends on the shape of b-SiC nanostructures, the degree of crystallinity and surface oxidation of fibers. In our paper we examined the b-SiC (confirmed by XRD method) which are coated with thin layer of amorphous silica (confirmed by TEM technique). In the context of Svensson et al. (1997) and Pourchez et al. (2012) results, one of the reasons for toxicity of SiC nanofibers tested in this study was the generation of reactive oxygen radicals and radical ions catalyzed by silicon carbide nanofibers.
4. Conclusion Our work has provided the first evidence that NFSiC and NRSiC, produced by combustion synthesis, can induce oxidative stress in P. putida. Based on the results presented in the paper, it can be stated that nanostructured SiC may affect the activity of microorganisms by increasing the formation of highly reactive radicals and disrupting cell membrane integrity. These mechanisms of cytotoxicity can lead to DNA damage in bacterial cells, and they can affect the expression of genes involved in the suppression of oxidative stress. Therefore, we analyzed the expression of the zwf-1 gene. SiC can affect bacterial activity, but this process is controlled by the texture of the material. The lmSiC used in this study as a reference material did not significantly influence the activity of P. putida. On the contrary, the nanostructured SiC seemed to affect the expression of oxidative stress genes and cell viability. Thus, the bacteria were likely undergoing increased cytotoxicity. However, it cannot be clearly stated whether this process led to cell death or only to cell damage. In addition to the physical interaction of SiC with the cell membrane, it is possible that the toxicity of nanostructured SiC is connected with the presence of silica nanofilms on the surface of the investigated materials.
Acknowledgments
Fig. 6. P. putida bacteria associated with silicon carbide nanofibers (NFSiC) (A) and nanorods (NRSiC) (B) after incubation with the investigated materials.
This research was partially supported by the European Union, within the European Regional Development Fund, through the Innovative Economy grant (POIG.02.02.00-00-025/09) and by the Faculty of Geology, University of Warsaw, BST 166901/2013. The authors thank reviewers for their critical remarks and comments which have improved this article.
A. Borkowski et al. / Chemosphere 135 (2015) 233–239
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