Growth and phenol consumption in phenol-supplemented medium
To investigate the tolerance of C. tropicalis SHC-03 to phenol, we transferred the pre-cultured cells into yeast extract peptone dextrose (YPD) medium containing 0.0 g/l, 0.5 g/l, 1.0 g/l, 2.0 g/l, and 3.0 g/l phenol, and observed the pattern of cell growth (change in cell density). The initial concentration of cells in the culture was adjusted to approximately 1.0 ´ absorbance value (optical density at 600 nm wavelength, OD600). Compared with the non-phenol-treated culture, the cell growth of phenol-treated C. tropicalis SHC-03 was slightly inhibited when the media contained 0.5 g/l and 1.0 g/l phenol (Fig. 1A). However, in the presence of 2.0 g/l and 3.0 g/l phenol, cell growth was hindered by approximately 40% and 100%, respectively (Fig. 1A). As illustrated in Fig. 1A, the growth rate of cells treated with 2.0 g/l phenol remained at a low level from 3 h to 24 h, implying that the phenol caused cell damage and influenced cell growth. However, after treatment for 24 h in the presence of 2.0 g/l phenol, the cells returned to normal levels of growth.
We speculated that phenol might be degraded to less toxic products via the β-ketoadipate pathway in C. tropicalis SHC-03 [33]. We therefore used the 4-aminoantipyrine spectrophotometric method to test for biodegradation of phenol in the different treatments (Fig. 1B). The results revealed no significant change in the concentration of phenol in any of the treatments at 0-72 h (Fig. 1B), which indicated that the phenol degradation mechanism of C. tropicalis SHC-03 was not activated in these conditions. To summarize, a range of resistance and/or detoxification mechanisms for protecting cells from the toxic damage of phenol were activated during the lag phase. Therefore, we employed RNA-seq technology and cytological techniques to reveal the potential tolerance mechanisms of C. tropicalis to phenol.
RNA-Seq, transcriptomic analysis, and qRT-PCR assays
After incubation with or without phenol for 3 h, the cells were harvested for RNA-seq (NCBI Accession Number: PRJNA591802). Based on quality-control test results, it was found that the quality score (Q30), clean data, and sequencing depth of each sample was, respectively, more than 94%, 4 Gb, and 675× (Additional File 1), which indicated a high level of accuracy for the RNA-seq results. To mitigate errors induced by biological variability between the samples, three biological replicates were used for the RNA-seq. The classification of the groups receiving different concentrations of phenol were as follows: 0.0 g/l: T01, T02, T03; 0.5 g/l: T04, T05, T06; 1.0 g/l: T07, T08, T09; 2.0 g/l: T10, T11, T12. As illustrated in Fig. 2A, the correlations between T01, T06, T09 and their corresponding biological replicates were relatively low. To guarantee reliability and accuracy of the results from the differential expression analysis, T01, T06, and T09 were eliminated (Fig. 2B).
The expression pattern of the 0.5 g/l phenol group was highly consistent with that of the control group; only 39 genes and 40 genes showed up- and down-regulated expression, respectively, with twofold changes in the 0.5 g/l phenol group compared with the control (Fig. 2D and 2E). In contrast, for the 1.0 and 2.0 g/l phenol groups, 353 and 1985 genes showed altered expression levels, respectively (Fig. 2C). Among these genes, 215 and 1050 genes were identified as up-regulated genes in the 1.0 and 2.0 g/l phenol groups, respectively (Fig. 2D), and 138 and 935 genes, respectively, were repressed by phenol (Fig. 2E). To sum up, the number of differentially expressed genes rose along with increasing phenol concentration.
In this study, 21 differentially expressed genes in response to 1.0 g/l phenol were selected to validate the accuracy of the results from RNA-seq through a qRT-PCR assay. The criterion for gene selection was a combination of high-low gene expression level (FPKM) and absolute value of log2(fold change). Through comparison and analysis, we discovered that the expression levels of 18 of 21 genes (approximately 85%) were consistent in the trend of up- and down-regulation (Fig. 2F). Of the other three genes, inconsistent with the data from RNA-seq, two displayed lower abosolute values of log2(fold change) (Fig. 2F). In summary, the results from RNA-Seq showed high accuracy, which contributed to the exploration of the phenol tolerance mechanism utilized by C. tropicalis.
Accumulation and scavenging of reactive oxygen species (ROS) in cells
Previous studies have found that phenol can pass through the cellular membrane and lead to increased membrane permeability and decreased membrane lipid-to-protein ratios, indicating the potential for phenol to cause dysfunction of organelles containing membranes, such as mitochondria, the endoplasmic reticulum, and the vacuole [9-11]. To detect damage to the mitochondrial membrane, Mito TrackerTM Green FM was used to stain the untreated and treated cells. Under phenol stress, the mitochondrial membrane displayed four different types of morphologies. In order of increasing damage, these were: tubular, fragmented, aggregated shapes, and necrotic (Fig. 3A). At 3 h, cells grown in media containing 0.0 g/l and 0.5 g/l phenol displayed tubular (95% and 50%, respectively) and fragmented (5% and 50%, respectively) mitochondria, while cells grown in media containing 2.0 g/l and 3.0 g/l phenol displayed aggregated (84% and 70%, respectively) and necrotic mitochondria (16% and 30%, respectively) (Fig. 3C). In contrast, at 9 h, only 3% of the non-phenol-treated cells appeared to be necrotic. At the same time point, among cells grown in 0.5 g/l phenol, the percentage of cells with aggregated and necrotic mitochondria remained at a low level (1% and 2%, respectively) (Fig. 3C). At 18 h, cells with tubular mitochondria were no longer seen in cells grown in media with or without phenol. In addition, compared with the results at 3 h and 9 h, the distribution of cells containing aggregated mitochondria increased tremendously, with 95% of the yeast cells cultured in 1.0 g/l phenol showing aggregated mitochondria at 18 h. Furthermore, in cells cultured in 2.0 g/l and 3.0 g/l phenol, the proportion of necrotic cells increased to 33% and 58% at 18 h, respectively (Fig. 3C). In summary, the ratio of mitochondrial deformation increased with the elevation of phenol concentration and treatment duration.
As most of the exogenous ROS were produced by damaged mitochondria [35], we hypothesized that cells treated with phenol should display excessive ROS. Therefore, 2’ 7’-dichlorofluorescein diacetates (DCFH-DA) was employed to stain the treated and untreated cells in order to detect intracellular ROS. The percentage of cells staining positive for ROS was considered to be representative of severity of oxidative stress (Fig. 3B). Cultures in media containing 2.5 mM, 5.0 mM, and 7.5 mM hydrogen peroxide served as the positive controls for ROS (Fig. 3D). In the medium without phenol, at 3 h and 9 h after treatment at 30°C, 5.6% and 6.5% of the cells exhibited a positive ROS signal, respectively (Fig. 3D). At 3 h, 9.1%, 10.2%, 30.3%, and 42.1% of cells stained positive for ROS when 0.5 g/l, 1.0 g/l, 2.0 g/l, and 3.0 g/l phenol were present, respectively (Fig. 3D), showing that the proportion of cells with ROS increased with increasing phenol concentration. At 9 h, the percent of these same cells staining positive for accumulated ROS had become 5.7%, 12.8%, 11.6%, and 100.0%, respectively (Fig. 3D). In addition, in the presence of 2.5 mM, 5.0 mM, and 7.5 mM hydrogen peroxide, 13.2%, 30.0%, and 53.1% of cells which had shown a positive ROS signal at 3 h decreased to 4.2%, 6.8%, and 5.6% at 9 h, respectively (Fig. 3D). The above results implied that the accumulation of ROS reached its peak at 3 h after treatment, and that, while the accumulation of excessive ROS at this time might cause damage to DNA, proteins, and lipids [37-39], the intracellular ROS were eliminated by several molecular mechanisms between 3 h and 9 h after treatment.
Efficient enzymatic and non-enzymatic antioxidant defense systems were found to be responsible for the scavenging of excessive ROS and protection of cells from oxidative damage [36]. Superoxide dismutases (SODs), glutathione peroxidases (GPX), catalase (CTT), glutathione reductase (GLR) and other scavenging enzymes functioned as enzymatic scavengers in enzyme-based antioxidant defense systems (Additional File 2). Thus, we investigated the transcription levels of these genes in the treated samples (Additional File 2). Based on the transcriptome data, we found that the expression levels of most of these genes were not significantly up-regulated against 0.5, 1.0, and 2.0 g/l phenol (Additional File 2). Among these genes, CTRG_04448, CTRG_04203, CTRG_01769, CTRG_00610, and CTRG_05111 exhibited 4.3-, 2.3-, 2.1-, 4.0-, and 3.2-fold downregulation, and CTRG_03986, CTRG_00152, CTRG_06042, and CTRG_02189 displayed 3.0-, 2.5-, 2.1-, and 2.6-fold upregulation when exposed to 2.0 g/l phenol (Additional File 2). In addition, one of these genes, CTRG_00142, displayed 2.1-fold upregulation when exposed to 1.0 g/l phenol. Given the transcriptome data, we had hypothesized that the enzymatic antioxidant defense systems would not be significantly activated for scavenging excessive ROS at 3 h. In contrast, the above results from the determination of ROS showed that the excessive ROS in the phenol-treated cells were scavenged between 3 h and 9 h (Fig. 3D).
The major ROS-scavenging pathways, the reduction of superoxide anion radicals (O2-) and hydrogen peroxide (H2O2), is catalyzed by SOD, GPX, and CTT (Fig. 4A) [36]. Therefore, we carried out enzyme activity assays for SOD, CTT, and GPX in the phenol-treated and non-phenol-treated cells (Fig. 4B, C). The results from the enzyme activity assays for SOD showed that, at 3 h, the enzyme activity of intracellular SOD increased dramatically along with increasing phenol concentration (Fig. 4B). At 6 h, the SOD activity of cells treated with 2.0 g/l phenol was significantly higher than that of cells treated with 0.0, 0.5, and 1.0 g/l phenol, but the SOD activity in cells treated with 3.0 g/l phenol was considerably lower (Fig. 4B). The SOD activities of all the treatments were lower at 6 h than those of the corresponding treatments at 3 h. However, few samples exhibited any SOD activity at 9 h (Fig. 4B). Since GPX and CTT are the key enzymes for the reduction of H2O2 (Fig. 4A), the activity of both of these enzymes was assayed. We found no significant differences in GPX activity in any of the treatments at 3 h. However, cells treated with 1.0 g/l phenol exhibited an increase in GPX activity at 9 h, and cells treated with 2.0 g/l phenol exhibited an increase in GPX activity at 6 h and 9 h. At both 6 h and 9 h, cells treated with 3.0 g/l phenol had lost their GPX activity almost completely (Fig. 4C). These results indicated that GPX activity was significantly upregulated in cells treated with 1.0 g/l and 2.0 g/l phenol after 6 h and 9 h, but that the high concentration of phenol (3.0 g/l phenol) induced the loss of cellular GPX activity at 6 h. With regard to CTT, the enzyme activity assays demonstrated no CTT activity in either phenol-treated or non-phenol-treated cells at the different processing times (data not shown). In addition, it has been found that high GLR activity and high GSH content can support high catalytic efficiency of GPX, which could protect the cells against ROS (Fig. 4A) [40]. In the present study, the enzyme activity of intracellular GLR was found to increase with phenol concentration at different processing times, except in those cells treated with 3.0 g/l phenol, at 6 h and 9 h (Fig. 4D). From the viewpoint of processing time, the cellular GLR activity after treatment for 9 h was higher than that after 3 h and 6 h in the presence of 0.0-2.0 g/l phenol. However, no GLR activity could be detected in cells treated with 3.0 g/l phenol, at either 6 h or 9 h (Fig. 4D). The GSH content of cells treated with 2.0 g/l and 3.0 g/l phenol was lower than that in cells treated with less than 2.0 g/l phenol at 3 h. However, after 3 h, the GSH content of cells treated with 0.0-2.0 g/l phenol rose rapidly after 6 h and 9 h, and remained at a high level (Fig. 4E). In addition, we found that the GSH content remained at the same level in the presence of different concentrations of phenol after treatment for 6 h and 9 h, except in the 3.0 g/l phenol group (Fig. 4E).
Damage to chromatin and protection of chromosomal DNA
In addition to DNA damage, ROS can cause nuclei to become less compacted and appear larger and more diffuse [41]. To monitor changes in the morphological structure of nuclei induced by ROS and/or phenol, the harvested cells were fixed and stained by a mixture of ethanol, ultrapure water, and DAPI (DNA specific dye). The structurally abnormal nuclear chromatin appeared larger and more diffuse, while the normal chromatin remained small and compacted (Fig. 5A). The percentage of cells with abnormal diffuse nuclear chromatin was recorded in order to investigate the severity of nuclear chromatin damage. At various time points, the non-phenol-treated and phenol-treated cells were harvested and stained with DAPI. As illustrated in Fig. 5B, the percentage of cells with nuclear chromatin disorganization remained at a low level (3.86%-7.30%) when phenol was present or absent, which implied that 0.0-3.0 g/l phenol did not cause obvious damage to DNA and chromatin.
The transcriptome data suggested that most of the genes participating in DNA replication and DNA repair showed significant upregulation in the presence of 2.0 g/l phenol (Fig. 5C and 5D) (Additional File 3). In addition, genes encoding ribonucleoside-diphosphate reductase subunit M2 (CTRG_01327 and CTRG_01698), as well as genes encoding ribonucleoside-diphosphate reductase subunit M1 (CTRG_01309), were highly expressed in cells exposed to 1.0 and 2.0 g/l phenol (Additional File 3). The encoded proteins of the above genes are small and large subunits of ribonucleotide reductase (RNR), which plays a crucial role in dNTP production and DNA synthesis [42]. Meanwhile, chromosome transmission fidelity protein 18 (CTRG_00975), which is essential for fidelity of chromosome transmission, was up-regulated by more than 4-fold in the presence of both 1.0 g/l and 2.0 g/l phenol [43] (Additional File 3).
Accumulation and degradation of unfolded and misfolded proteins
External or internal ROS can cause the accumulation of unfolded or misfolded proteins in the endoplasmic reticulum (ER) lumen; a phenomenon known as ER stress (ERS) [37]. To monitor ROS-induced ER membrane damage, the phenol-treated cells were stained with the endoplasmic reticulum dye, ER-TrackerTM Red. For the observation, the ER structures of the viable cells were divided into three groups: normal (unfolded), abnormal shapes (folded and fragmented), and necrotic (Fig. 6A). When cells were exposed to 0.0 g/l, 0.5 g/l, and 1.0 g/l phenol, normal and abnormal ER structures were distributed in roughly 35% and 65% of cells at 3 h, respectively (Fig. 6C). Under the same conditions, there was no significant change in the ratio of cells containing normal and abnormal ER structures from 3 h to 9 h, but there was a slight increase in the proportion of cells containing abnormal ER at 18 h (Fig. 6C). The necrotic cells and the cells containing abnormal ER accounted for a large share of the observed cells treated with high concentrations of phenol (2.0 g/l and 3.0 g/l) at various time points (Fig. 6C). These results demonstrated that ER was not significantly injured by low concentrations of phenol (0.5 g/l and 1.0 g/l), but was markedly damaged by high concentrations of phenol (2.0 g/l and 3.0 g/l). In contrast, the recovery of cell growth in cells exposed to 2.0 g/l phenol after 24 h suggested that certain mechanisms provided enough protein to retain homeostasis in the cells. Previous studies have demonstrated that, after the unfolded protein response (UPR) and autophagy, cells could recover homeostasis and normal ER function [44]. HSPs functioning as chaperones were implicated in the reversal of amino acid oxidation and refolding of denatured proteins resulting from the UPR [45-47]. The expression of one gene, annotated as ‘small heat shock protein 21’ (Hsp21) (CTRG_01443), was up-regulated by 11.3-, 64-, and 13.9-fold change in the presence of 0.5 g/l, 1.0 g/l, and 2.0 g/l phenol, respectively (Additional File 4). Besides that, the gene CTRG_04372 annotated as co-chaperones in the Hsp70/Hsp90 family was significantly upregulated in the presence of 2.0 g/l phenol (Additional File 4). Autophagy, as a conserved trafficking pathway, delivered unfolded or misfolded proteins, components, and organelles from the cytoplasm to the vacuole for degradation and recycling [44]. Four pivotal steps, including the activation of autophagy, the formation of the autophagosome, the cytoplasm-to-vacuole targeting (CVT) pathway, and vacuole fusion, ensured the effective degradation of the contents by vacuole hydrolases [44, 48, 49]. In the presence of 2.0 g/l phenol, it was found that the expression of genes IRE1, ATG11, ATG23, and ATG25, related to the activation of autophagy and the formation of the autophagosome, were up-regulated by more than 2-fold (Additional File 4). However, APE1 and AMS1, which are related to autophagy, showed down-regulation by 2-fold (Additional File 4).
The vacuole plays a key role in autophagy for the degradation and recycling of unfolded or misfolded proteins [48]. Therefore, Yeast Vacuole Membrane Marker MDY-64 was adopted to stain the vacuole membrane for visualizing changes in vacuole morphology in the non-phenol-treated and phenol-treated cells. The observation revealed vacuoles in different configurations that could be classified as follows: a single large vacuole, two to four medium-sized vacuoles, and massively fragmented vacuoles (Fig. 6B). After treatment for 3 h, compared with the other treatments, 75% of cells treated with 0.5 g/l phenol contained a single large vacuole. A single large vacuole was also predominant in cells treated with 1.0 and 2.0 g/l phenol (71% and 68%, respectively) (Fig. 6D). In the same processing time, cells treated with non-lethal doses of phenol (0.5 g/l, 1.0 g/l, and 2.0 g/l) exhibited a lower proportion of fragmented vacuoles than untreated cells (Fig. 6D). At 9 h after treatment, cells treated with 1.0 g/l phenol (49%) showed the highest proportion of single large vacuoles, followed by cells treated with 0.5 g/l phenol (40%) (Fig. 6D). From these observations, we speculated that low concentrations of phenol suppressed the fragmentation of the cellular vacuole. However, the cells treated with 2.0 g/l and 3.0 g/l phenol exhibited a higher proportion of fragmented vacuoles than the untreated cells, after treatment for both 9 h and 18 h (Fig. 6D). Since mutation of VAC8 has been correlated with vacuole fragmentation [49], we speculated that the above results might be related to downregulation of VAC8 (Additional File 4).
Accumulation of fatty acid
The KEGG pathway enrichment analysis revealed downregulation of 13 genes involved in fatty acid degradation (Fig. 7). In this pathway, long-chain acyl-CoA synthetase (EC: 6.2.1.3) is responsible for the degradation of hexadecanoate (fatty acid), and the corresponding genes, CTRG_02563 and CTRG_05500, were downregulated by 11.3- and 7-fold in response to 2.0 g/l phenol, respectively (Fig. 7). When hexa-decanoyl-CoA was converted into trans-hexadec-2-enoyl-CoA, the expression of the corresponding genes (CTRG_02374, CTRG_02377, CTRG_02721, and CTRG_05958), annotated as acyl-CoA oxidase (EC: 1.3.3.6) and acyl-CoA dehydrogenase (EC: 1.3.8.7), showed 147-, 12.1-, 2.8- and 5.7-fold decreases in cells treated with 2.0 g/l phenol, respectively (Fig. 7). The genes CTRG_01068 and CTRG_02168, which encode acetyl-CoA acyltransferase1 (2.3.1.16), were 7- and 3.7-fold downregulated in cells treated with 2.0 g/l phenol (Fig. 7). In addition, the differential expression analysis implicated genes encoding aldehyde dehydrogenase (1.2.1.3) and alcohol dehydrogenase (1.1.1.1) in the conversion of fatty acid to alcohol (Fig. 7). Specifically, CTRG_05836, CTRG_00882, CTRG_05482, CTRG_01329, and CTRG_05010 were significantly downregulated in response to 2.0 g/l phenol exposure (Fig. 7). These results suggest that decreased fatty acid degradation efficiency, caused by the downregulation of expression of related genes, may serve to elevate the intracellular fatty acid content or change the fatty acid component in cells [13, 50].
Other tolerance mechanisms
Previous studies have found that cell-wall remodeling can lead to increased ethanol resistance in yeast [51]. Our comparative transcriptome analysis revealed >twofold upregulation of six genes related to cell-wall biogenesis and integrity in response to 1.0 and 2.0 g/l phenol, including CTRG_05721, CTRG_05949, CTRG_00608, CTRG_00036, CTRG_01855, and CTRG_03473 (Additional File 5). We therefore reasoned that genes involved in cell-wall biogenesis may contribute to the increased resistance of C. tropicalis SHC-03 to phenol. A series of differential expression analyses of genes upregulated in response to 2.0 g/l phenol revealed enrichment in the chitin synthesis pathway (Fig. 8A; Additional File 5). As illustrated in Fig. 8A, during the conversion of glucose to chitin, the expression levels of CTRG_00414, CTRG_00601, CTRG_01436, CTRG_03651, CTRG_03585, CTRG_05721, and CTRG_05949 were markedly increased. In addition, only two proteins are known to play a reverse role in the chitin synthesis pathway, and their encoding genes (CTRG_03727 and CTRG_03728) were downregulated by 23.4- and 13.6-fold, respectively (Fig. 8A; Additional File 5). In addition, most of the genes in the chitin degradation pathway (chitin to chitobiose or N-Acetyl-D-glucosamine), including CTRG_05456 and CTRG_05827 (encoding chitinase) and CTRG_01063 (encoding beta-N-acetylhexosaminidase), were significantly downregulated in expression (Fig. 8A; Additional File 5). In the process of converting chitin to chitosan, CTRG_01049, which encodes chitin deacetylase, was dramatically upregulated; by 26.5-fold (Fig. 8A; Additional File 5). Therefore, we speculated that the up- and down-regulated expression of these genes probably served to increase the accumulation of chitin and chitosan in the cell wall. To confirm the increased phenol tolerance of cells that had undergone cell wall remodeling, we conducted cell-wall susceptibility analyses of the phenol-treated and non-phenol-treated cells using lytic enzyme, a β-1,3-glucanase from Arthrobacter luteus [51]. After incubation in media supplemented with different concentrations of phenol for 3 h, 9 h, and 18 h at 30 °C, cells were isolated for cell-wall susceptibility analysis. After 3 h, the cell density (OD600) of samples treated with 0.0 and 0.5 g/l phenol decreased significantly upon addition of lyticase to the medium. This decrease was even more pronounced in cells treated with 1.0 g/l phenol and 3.0 g/l phenol, decreasing to less than 10% of the initial cell density at 4 h (Fig. 8B). The cell density of samples treated with 2.0 g/l phenol dropped slowly after lyticase was added into the medium for 4 h, decreasing to about 65% at 4 h (Fig. 8B). After treatment with phenol for 9 h, the samples in 0.0 g/l and 0.5 g/l phenol still displayed the most rapid cell-density decline in the lyticase-supplemented medium; the decrease in cell density in these samples was greater than the cell-density decrease seen in the samples in 1.0-3.0 g/l phenol (Fig. 8C). However, the cell density of the sample in 3.0 g/l phenol dropped more quickly than all of the others, at 3 h (Fig. 8C). The resistance of cells to lyticase after treatment with 1.0 g/l and 2.0 g/l phenol for 9 h was similar to that of cells treated with the corresponding concentration of phenol for 3 h (Fig. 8C). In cells exposed to phenol for 18 h, the cell density of the sample treated with 3.0 g/l phenol descended sharply, decreasing to about 12% after addition of lyticase to the medium and incubation for 4 h (Fig. 8D). The cell density of samples treated with 1.0 g/l and 2.0 g/l phenol for 18 hours dropped slowly after the 4-h lyticase treatment. In contrast, the cell density in the samples with 0.0 g/l and 0.5 g/l phenol showed no significant decline (Fig. 8D). The above results demonstrated that treatment for 3 h and 9 h with 1.0-3.0 g/l phenol induced increased resistance of cells to lyticase, with a maximal effect seen at 2.0 g/l phenol. While 2.0 g/l phenol appeared to be the optimal concentration for lyticase resistance, cells treated with 0.0 g/l and 0.5 g/l phenol also exhibited significantly increased resistance to lyticase after 18 h. However, cells treated with 3.0 g/l phenol for 18 h showed decreased resistance to lyticase compared with this treatment after 3 h and 9 h.
One important detoxification mechanism is the active efflux mechanism, which has been shown to reduce the level of intracellular toxic compounds, resulting in retention of the physiological activities of the cells [52]. The major facilitator superfamily (MFS) and ATP-binding cassette (ABC) subfamily are the two most important groups of multidrug/multixenobiotic resistance (MDR/MXR) transporters responsible for the efflux of toxic compounds [53, 54]. In response to at least two concentrations of phenol, the MFS genes CTRG_03938, CTRG_00385, and CTRG_03729 showed a >2-fold increase in expression (Additional File 6). In particular, CTRG_00385 was up-regulated by 7-fold, 18-fold, and 169-fold in response to 0.5 g/l, 1.0 g/l, and 2.0 g/l phenol. Additionally, a statistical analysis of differential gene expression showed that 14 genes belonging to the MFS and ATP-binding cassette (ABC) subfamily exhibited greatly up-regulated expression in the presence of phenol (Additional File 6). As illustrated in Additional File 6, two, six, and ten transporter genes were significantly upregulated in response to 0.5 g/l, 1.0 g/l, and 2.0 g/l phenol, which indicated that the number of upregulated transporter genes increased with the rise in phenol concentration. Compared with tolerance mechanisms, detoxification mechanisms can contribute to the detoxification of the toxic compounds to less toxic or nontoxic compounds, improving the growing environment of the strains [19, 20]. Previous studies have illustrated that phenol degradation is the vital detoxification mechanism of C. tropicalis in response to phenol [34, 55]. From the point of view of the molecular mechanism, the biodegradation of phenol mainly relies on subsequent enzymatic steps via the β-ketoadipate pathway in C. tropicalis [33]. The crucial enzyme in this pathway, phenol 2-monooxygenase (EC 1.14.13.7), is responsible for the hydroxylation of phenol to catechol, which is a rate-limiting step [33](Fig. 9). To date, two genes encoding phenol 2-monooxygenase, CTRG_00423 and CTRG_03102, have been discovered in C. tropicalis strains JH8 and MYA-3404, respectively [32, 56]. In the present study, the comparative transcriptomics data did not indicate that CTRG_00423 and CTRG_03102 were significantly up-regulated under phenol stress after treatment for 3 h (Fig. 9). In the second step of the β-ketoadipate pathway, catechol 1,2-dioxygenase (EC 1.13.11.1) is responsible for the conversion of catechol to cis,cis-muconate (Fig. 9). CTRG_01732 and CTRG_00171, encoding catechol 1,2-dioxygenase, were not significantly up-regulated in response to different concentrations of phenol (Fig. 9). The above results suggest that the phenol degradation mechanism of C. tropicalis SHC-03 was probably not activated.