Influence of Growth Substrate and Contaminant Mixtures on the Degradation of BTEX and MTBE by Rhodococcus rhodochrous ATCC Strain 21198

Abstract

The degradation of the prevalent environmental contaminants benzene, toluene, ethylbenzene, and xylenes (BTEX) along with a common co-contaminant methyl tert-butyl ether (MTBE) by Rhodococcus rhodochrous ATCC Strain 21198 was investigated. The ability of 21198 to degrade these contaminants individually and in mixtures was evaluated with resting cells grown on isobutane, 1-butanol, and 2-butanol. Growth of 21198 in the presence of BTEX and MTBE was also studied to determine the growth substrate that best supports simultaneous microbial growth and contaminants degradation. Cells grown on isobutane, 1-butanol, and 2-butanol were all capable of degrading the contaminants, with isobutane grown cells exhibiting the most rapid degradation rates and 1-butanol grown cells exhibiting the slowest. However, in conditions where BTEX and MTBE were present during microbial growth, 1-butanol was determined to be an effective substrate for supporting concurrent growth and contaminant degradation. Contaminant degradation was found to be a combination of metabolic and cometabolic processes. Evidence for growth of 21198 on benzene and toluene is presented along with a possible transformation pathway. MTBE was cometabolically transformed to tertiary butyl alcohol, which was also observed to be transformed by 21198. This work demonstrates the possible utility of primary and secondary alcohols to support biodegradation of monoaromatic hydrocarbons and MTBE. Furthermore, the utility of 21198 for bioremediation applications has been expanded to include BTEX and MTBE.

1.0. Introduction

Benzene, toluene, ethylbenzene, and xylenes (BTEX) are ubiquitous environmental contaminants introduced to the environment from both natural and anthropogenic sources such as forest fires, the combustion and spills of petroleum products, and manufacturing industries including solvent, paint, and rubber industries (Latif et al. 2019; Shim et al. 2002; Barboni and Chiaramonti 2010). The US Environmental Protection Agency (EPA) recognizes BTEX as priority pollutants (US EPA 2014), as they are associated with a multitude of adverse health effects including carcinogenicity, neurotoxicity, and reproductive toxicity (Masih et al. 2016; Hazrati et al. 2016; Sairat et al. 2015). Despite their known toxicity, BTEX continue to have a significant presence in atmospheric and aquatic environments leading to regular human exposures (Y. Li et al. 2020; Dobaradaran et al. 2021; Alfoldy et al. 2019). Of particular concern is drinking water BTEX contamination, as even trace levels of BTEX in drinking water are considered hazardous. Maximum contaminant levels (MCLs) for benzene, toluene, ethylbenzene, and total xylenes are 0.005, 1, 0.7, and 10 mg/L, respectively (US EPA 2009).

Methyl tert-butyl ether (MTBE) was a common gasoline additive in the late 20^th century used to increase the octane level of gasoline and thus promote more complete fuel combustion in automobiles (Steffan et al. 1997; Hernandez-Perez et al. 2001). MTBE quickly became dispersed in the environment due to its extensive use, and despite its replacement in gasoline by ethanol in the 2000s, it continues to be a common surface water and groundwater contaminant in the US (Fiorenza and Rifai 2003.; Carter et al. 2006). Furthermore, MTBE is still used as a gasoline additive in many countries, such as China and Mexico (S. Li et al. 2019a; González et al. 2018). MTBE is a suspected human carcinogen, and although no MCL has been established for it, an action level range of 20 μg/L to 40 μg/L has been proposed by the EPA (US EPA 1997).

Although BTEX is suitable for physical removal strategies such as air stripping or adsorption for treatment of contaminated water, a less invasive and inexpensive method of BTEX remediation is bioremediation (Farhadian et al. 2008; Juwarkar et al. 2010). Many microorganisms are capable of degrading BTEX as carbon and energy sources via metabolism, or in conjunction with an alternative carbon and energy source via cometabolism. Despite initially being considered recalcitrant to microorganisms, MTBE has also been demonstrated to be biodegradable by many microorganisms (Hristova et al. 2003; Chen et al. 2011; Mahmoodsaleh and Roayaei Ardakani 2021). However, MTBE is most often found in the environment with hydrocarbon co-contaminants such as BTEX, and BTEX contamination itself typically occurs as a mixture. Thus, bioremediation efforts for BTEX and MTBE (BTEXM) must account for mixture interactions that occur within BTEX and between BTEX and MTBE. Contaminant mixtures often introduce complexities such as competition and inhibition that can reduce the success of bioremediation efforts (Deeb and Alvarez-Cohen 1999; Bielefeldt and Stensel 1999). To overcome the negative impacts of mixtures on biodegradation, microorganisms have been isolated using BTEX mixtures as enrichment substrates in pursuit of microbes that can degrade, or at minimum tolerate, all components of BTEX (Benedek et al. 2021; Hocinat et al. 2020; Wongbunmak et al. 2020; Kasi et al. 2013). Similar methods have also been used for enriching BTEXM degrading microorganisms using MTBE as the enrichment substrate (Lin et al. 2007; Raynal and Pruden 2008; Pruden and Suidan 2004). In doing so, isolates usually require BTEX or MTBE to sustain growth and activity, limiting their ability to degrade contaminants to remediation goal concentrations. Cometabolic processes avoid this issue, as microorganisms that cometabolize BTEXM are not dependent on the contaminants for their survival. Additionally, enzymes involved in MTBE oxidation typically have a low affinity for MTBE (high K_s), thus increasing the difficulty of achieving low concentrations in the environment (House and Hyman 2010; Smith and Hyman 2004).

Rhodococcus rhodochrous ATCC 21198 (21198) is a gram-positive bacterium with many qualities attractive for bioremediation applications such as planktonic growth, metabolic diversity, and expression of a short chain alkane monooxygenase (SCAM) that can transform chlorinated aliphatic hydrocarbons (CAHs) as well the cyclic ether 1,4-dioxane (1,4-D) (Rasmussen et al. 2020; Rolston et al., 2022). High expression of SCAM has been observed when 21198 is grown on alkanes such as propane and isobutane, however SCAM expression has also been shown to be supported by alcohols such as 2-butanol (Chen 2016). The Rhodococcus genera has an established relevance in bioremediation for n-alkanes and aromatic hydrocarbons (Brzeszcz and Kaszycki 2018). Although other studies of Rhodococcus strains with BTEXM have been conducted (Deeb et al. 2001; Jung and Park 2004; Kim et al. 2002; Deeb and Alvarez-Cohen 1999), including studies using aliphatic hydrocarbons as growth substrates (Auffret et al. 2009; Lee and Cho 2009), alcohols have not been investigated as potential growth substrates to support degradation of BTEXM. The objectives of this research were to establish the ability of 21198 to degrade BTEX and MTBE both individually and in mixtures, distinguish between metabolic and cometabolic processes involved in BTEXM degradation, and demonstrate the utility of alcohol growth substrates to support the degradation of BTEXM in settings of resting cell biomass and proliferating cell biomass.

2.0. Materials and Methods

2.1. Chemicals

All chemicals used were analytical grade purity: benzene (99%, Beantown Chemical, New Hampshire), toluene (99.8%, Sigma Aldrich, Missouri), ethylbenzene (99.8%, Acros Organics, Massachusetts), o-xylene, m-xylene, p-xylene (99%, Alfa Aesar, Massachusetts), MTBE, TBA (99%, TCI America, Oregon), isobutane (99.99%, Gas Innovations, Texas), 1-butanol (99.4%, Acros Organics, Massachusetts), 2-butanol (99%, Sigma Aldrich, Missouri).

2.2. Rhodococcus rhodochrous ATCC 21198 Culture Methods

The pure culture Rhodococcus rhodochrous ATCC 21198 (21198) was provided by Dr. Michael Hyman from North Carolina State University. 21198 was maintained as a pure culture on minimal salt media (MSM) agar plates in a sterile airtight jar with isobutane supplied as the sole carbon and energy source. Growth reactors were prepared as described in Rolston et al. (2019). Briefly, an inoculum scraped from a minimal plate was added to a 500 mL Wheaton glass media bottle containing 300 mL of sterile MSM (Kottegoda et al. 2015) amended with either isobutane, 1-butanol, or 2-butanol in excess based on the oxygen content of a sealed bottle. Growth reactors were incubated in the dark at 30^oC on a 150 RPM shaker table. Microbial growth was monitored via optical density absorption at 600 nm (OD₆₀₀) using a Thermo Scientific Orion Aquamate 8000 UV–Vis Spectrophotometer. When an OD₆₀₀ of approximately 0.3 was reached, growth reactors were refreshed to enhance biomass growth. During refreshment, reactors were opened in a laminar flow hood to equilibrate with atmospheric oxygen and streaked on a tryptic soy glucose agar (TSGA) plate to confirm culture purity. A second addition of growth substrate was added to growth reactors before reclosing them. Cells in late exponential growth phase were concentrated into a 50 mM sodium phosphate buffer via centrifugation the day after refreshing as described in Murnane et al. (2021). TSS analysis on the cell concentrate was performed according to AWWA standard protocol (Baird et al. 2017) to determine the biomass concentration. Concentrated cell solutions were stored at 4^oC and used within 2 days of harvesting.

2.3. Analytical Methods

2.3.1. Headspace Analysis

Volatile compounds were analyzed using gas chromatography (GC). For all headspace GC analyses, a Hamilton 1710 Series gas-tight syringe was used to inject 100 μL of headspace sample from the batch reactors into the gas chromatograph. BTEX and isobutane were measured using a Series 6890 Hewlett Packard Gas Chromatograph equipped with a flame ionization detector (FID). Helium flowed through the capillary column (Agilent DB-624 UI 30m x 0.53mm) at 15 mL/minute. The detector temperature and oven temperature were held constant at 250^oC and 220^oC, respectively. Chromatographic separation of all volatile hydrocarbons was achieved with the exception of m-X and p-X, which coeluted. Oxygen was measured using a Hewlet Packard 5890 Series Gas Chromatograph equipped with thermal conductivity detector (GC-TCD) and capillary column (Supelco 60/80 Caroboxen 1000). Helium flowing at 30 mL/minute was used as the carrier gas, with the oven temperature held constant at 40^oC. All headspace methods were calibrated using external standards, with good linear fit (R²> 0.99) achieved for all calibration curves. Henry’s coefficients for each compound were used to calculate the total mass of contaminants in bottles based on the headspace measurements. A list of Henry’s coefficients used is reported in Table S4.

2.3.2. Liquid Sample Analysis

Non-volatile compounds, namely MTBE, tert-butyl alcohol (TBA), 1-butanol, and 2-butanol, were analyzed in liquid samples from batch reactors. 1 mL liquid samples were taken throughout the course of batch experiments through the cap septa using a sterile syringe. These samples were filtered through 0.2 μm polyvinylidene difluoride (PVDF) filters to remove cells and stored at 4^oC until analysis. MTBE, TBA, 1-butanol, and 2-butanol were analyzed using a Hewlet Packard 6890 Series Gas Chromatograph equipped with a Hewlet Packard 5873 mass selective detector (GC-MS) and capillary column (Restk Rtx-VMS). Filtered liquid samples were diluted in 40 mL VOA vials filled with 25 mL deionized water to concentrations below 20 μg/L. 5 mL of diluted sample was taken from each VOA via a Teledyne Tekmar AQUATek100 autosampler and run through a Teledyne Tekmar Lumin PTC purge and trap. Details on the heated purge and trap method are provided in Rolston et al. (2019). The mass detector was operated in selective ion monitoring mode to quantify ion fragments associated with MTBE (m/z 73), TBA (m/z 59), 1-butanol (m/z 56), and 2-butanol (m/z 45). This method was calibrated with external standards and standards were included in sample analyses runs to monitor instrument sensitivity fluctuations.

2.4. Batch Reactor Experiment Design

2.4.1. Resting Cell Experiments

Resting cell batch reactor tests with single contaminants and contaminant mixtures were conducted in 125 mL Wheaton serum bottles with butyl septa caps. Each bottle contained 100 mL of MSM and approximately 1 mg of the contaminant(s) added as neat liquids. Actual masses of contaminants added in each experiment are reported in Table S5. Bottles were allowed to reach liquid-gas phase equilibrium at 30^oC on a 150 RPM shaker table, and these conditions were maintained for the duration of the experiments. Initial headspace measurements were taken before 5 mg of cells were added to bottles through the septa using a sterile syringe. Headspace and liquid samples were taken periodically to monitor contaminant concentrations. Control bottles were prepared without cells to monitor abiotic losses of analytes. Active bottles were prepared in triplicates, while control bottles were prepared in duplicates. Zero-order rates were derived from the calculated total contaminant mass using the linear regression model “fitlm” in MATLAB. The linear regions used to calculate rates and goodness of fit are reported in Figure S2. Initial periods of minimal transformation, referred to hereafter as lag periods, were estimated by the length of time between the first measurement and the beginning of the defined linear region. On occasion, the initial timepoint was omitted from the linear portion to improve goodness of fit. Such situations were not considered to be part of the lag period if this was the only data point omitted.

2.4.2. Growth Batch Reactor Experiments

Four sets of batch reactors were prepared to investigate 21198 growth in the presence of BTEXM. Each set was prepared in 125 mL sterilized Wheaton serum bottles with butyl septa caps with 100 mL MSM, ~1 mg of each contaminant, and a low initial biomass, 0.05 mg of isobutane grown cells, compared to the resting cell tests. Batch reactors were amended with approximately 1 mg of growth substrate, with the exception of set 1, which did not receive any additional growth substrate. Isobutane, 1-butanol, and 2-butanol were added to sets 2, 3, and 4, respectively as growth substrates. As oxygen was depleted from the headspace air, pure gaseous oxygen was added to the headspace through the cap septa. Control bottles were prepared without cells to monitor abiotic losses of analytes. Batch reactors were prepared in triplicates, while control bottles were prepared in duplicates. All bottles were incubated in the dark on a 100 RPM shaker table at 20^oC for the duration of the experiments.

2.4.3. Growth Substrate Experiments

To explore the range of growth substrates for 21198, growth substrate experiments were conducted in 125 mL Wheaton serum bottles (nominal volume 155 mL) with butyl septa caps and 25 mL MSM. Potential growth substrates were added to from neat stocks in excess based on the amount of available oxygen in the air headspace. Specific volumes of each substrate added are reported in Table S1. Growth bottles were inoculated with ~0.1 mg isobutane grown cells from a cell concentrate, and cell growth was monitored periodically using OD₆₀₀ measurements. Bottles were incubated in the dark at 30^oC on a 150 RPM shaker table for the duration of the experiments.

3.0. Results and Discussion

3.1. Individual Contaminant Batch Experiments

3.1.1. BTEX Compounds

The influence of growth substrate on 21198’s BTEXM degradative capability was investigated using resting 21198 cells grown on either isobutane, 1-butanol, or 2-butanol. Degradation was first studied in individual contaminant tests shown in Figure 1.

Fig. 1.

Degradation of benzene (A), toluene (B), ethylbenzene(C), o-Xylene (D), m-Xylene (E), and p-Xylene (F) in individual contaminant tests by 21198 grown on isobutane (circles), 1-butanol (diamonds), or 2-butanol (squares). Error bars represent standard deviation between triplicates

Isobutane grown cells consistently exhibited the most rapid transformation rates for all contaminants tested, followed by 2-butanol grown cells and 1-butanol grown cells. Lag periods were only observed with alcohol grown cells, as reported in Table 1.

Table 1.

Zero-order transformation rates and lag periods associated with batch experiments. Error associated with transformation rates is the standard deviation among triplicates.

Contaminant	Batch Experiment	Isobutane Grown Cells		1-Butanol Grown Cells		2-Butanol Grown Cells
		Rate (μmol/hour/mg biomass)	Lag Time (hours)	Rate (μmol/hour/mg biomass)	Lag Time (hours)	Rate (μmol/hour/mg biomass)	Lag Time (hours)
Benzene	Individual BTEX BTEXM	0.43 ± 0.04 0.48 ± 0.06 0.50 ± 0.05	0 1.3 0	0.16 ± 0.02 0.10 ± 0.01 0.11 ± 0.01	11 10 23	0.25 ± 0.01 0.24 ± 0.06 0.25 ± 0.04	3.9 4.6 6.5
Toluene	Individual BTEX BTEXM	5.83 ± 0.04 1.35 ± 0.06 2.17 ± 0.14	0 0 0	0.31 ± 0.001 0.25 ± 0.02 0.13 ± 0.01	7 5.9 6.4	0.84 ± 0.04 0.30 ± 0.01 0.29 ± 0.01	0 0 0
Ethylbenzene	Individual BTEX BTEXM	0.86 ± 0.02 0.73 ± 0.04 1.11 ± 0.05	0 0 0	0.10 ± 0.01 0.10 ± 0.02 0.08 ± 0.01	0 5.9 6.4	0.34 ± 0.004 0.18 ± 0.01 0.20 ± 0.01	0 0 0
o-Xylene	Individual BTEX BTEXM	0.33 ± 0.02 0.29 ± 0.01 0.47 ± 0.04	0 0 0	0.04 ± 0.01 0.12 ± 0.01 0.08 ± 0.01	0 10 10	0.34 ± 0.004 0.18 ± 0.01 0.20 ± 0.01	0 3.0 2.0
m-Xylene	Individual	7.17 ± 0.16	0	0.17 ± 0.01	0	0.58 ± 0.04	0
p-Xylene	Individual	1.56 ± 0.11	0	0.07 ± 0.003	0	0.18 ± 0.01	0
MTBE	Individual BTEXM	0.16 ± 0.02 0.25 ± 0.04	0 0	0.03 ± 0.01 0.03 ± 0.008	0 26	0.16 ± 0.03 0.08 ± 0.01	0 0

A lag period for benzene transformation was observed with 1- and 2-butanol grown 21198, while a lag period for toluene was only observed with 1-butanol grown cells. The incidence of lag periods with alcohol grown cells indicated a potential induction period, during which the cells produced the enzyme required to transform the contaminants. Because transformation of these contaminants commenced following this period, enzyme(s) required for these transformations are potentially inducible by the presence of BTEX. Among the aromatic compounds, T and m-X were transformed most rapidly, while o-X was transformed slowly. Based on the significant differences in transformation rates reported in Table 1 among the BTEX components, it appears that a single methyl or ethyl substitution on the ring makes the compound more susceptible to enzymatic attack by 21198. However, as demonstrated by the range of degradation rates between xylene isomers, a second methyl substitution on the ring can have negative impacts on degradability. Of the three xylene isomers, o-X is commonly observed to be the most difficult to degrade microbially (Auffret et al. 2009; Taki et al. 2006; Jeong et al. 2008. Because of this, o-X has been used as an enrichment substrate to isolate o-X degrading Rhodococcus strains. Rhodococcus isolates that preferentially degrade o-X over the other isomers have been characterized (Jeong et al. 2008; You et al. 2018). The same was achieved by Auffret et al. (2009) using a BTEX and aliphatic hydrocarbon mixture to isolate a Rhodococcus wratislaviensis strain. However, Taki et al. (2006) reported the same xylene isomer preference as 21198 (m-X > p-X > o-X) with a Rhodococcus consortia despite using o-X as the consortia’s enrichment substrate. 21198’s ability to cometabolically degrade all xylene isomers, while not requiring their presence during its growth phase, indicates the relaxed substrate specificity of the enzyme(s) involved in their degradation. Abiotic losses in these experiments were minimal compared to transformation of contaminants observed in active bottles, and are reported in Table S2.

Along with the rate of contaminant degradation, the extent of degradation was impacted by growth substrate. While complete contaminant degradation was achieved by isobutane grown cells for all BTEX components, 1-butanol and 2-butanol grown cells were unable to fully degrade o-X in the duration of the experiment. 1-butanol grown cells were also unable to fully degrade p-X. A lower transformation capacity, or mass of contaminant able to be transformed by a certain resting cell mass, for o-X and p-X may be the result of stress to the microbes such as transformation product toxicity or depletion of the cells’ energy reserves.

3.1.2. MTBE and TBA

MTBE transformation progressed slower than the BTEX compounds across all growth substrates, as shown in Figure 2A. However, the relative rate and extent of degradation followed the same pattern amongst the growth substrates as observed with BTEX (isobutane > 2-butanol > 1-butanol).

Fig. 2.

Transformation of MTBE by 21198 grown on isobutane (circles), 1-butanol (diamonds), or 2-butanol (squares) shown in A. Simultaneous MTBE (filled circle) transformation and TBA (open circle) production by isobutane grown 21198 shown in B. Error bars represent standard deviation between triplicates in A and duplicates in B.

No lag period was observed for MTBE transformation by cells grown on either isobutane or butanol. Isobutane grown cells achieved MTBE concentrations below the EPA action level concentration (20 μg/L) after 76 hours of incubation, while 1-butanol and 2-butanol grown cells reached concentrations of 1.2 mg/L and 0.12 mg/L, respectively, after 100 hours of incubation. TBA was identified as the major transformation product of MTBE produced by 21198. TBA is the most common intermediate reported in MTBE aerobic degradation pathways (Nava et al. 2007; Fiorenza and Rifai, 2003). Some microorganisms such as Arthrobacter ATCC 27778 (Liu, Speitel, and Georgiou 2001), Acinetobacter Sp. Strain SL3 (Li et al. 2019b) and ENV425 ATCC 55798 (Steffan et al. 1997) are able to utilize the same enzyme for MTBE and TBA oxidation, producing precursors for intermediates, such as 2-hydroxyisobutyric acid (HIBA). Other microorganisms utilize different enzymes for MTBE and TBA oxidation, such as Mycobacterium austroafricanum IFP 2012 (François et al. 2002) and Methylibium petroleiphilum strain PM1 (Deeb and Alvarez-Cohen 2000). However, many MTBE oxidizing organisms are unable to further oxidize TBA, including Pseudomonas mendocina KR-1 (Smith et al. 2003), Pseudomonas putida GPo1 (Smith and Hyman 2004), and Gordonia terrae IFP 2001 (Hernandez-Perez et al. 2001). Accumulation of TBA is not desirable, as TBA has similar properties, including toxicity, to MTBE (Nava et al.2007).

The transformation of TBA was observed in MTBE batch tests with resting isobutane grown 21198 cells as shown in Figure 2B, indicating that the transformation of MTBE does not end at TBA, however the transformation of TBA occurs more slowly than that of MTBE. House and Hyman (2010) observed slower TBA oxidation rates compared to MTBE as well with Mycobacterium austroafricanum JOB5. It was concluded that TBA and MTBE compete for the same monooxygenase, and that the lower TBA transformation rate is due to a higher K_s value for TBA compared to MTBE. Catalytic similarities between 21198 and JOB5 with regards to both BTEX and MTBE degradation suggest that this phenomenon documented in JOB5 may also be occurring in 21198.

3.1.3. Connections to Additional Contaminants of Concern

The relative activity of 21198 grown on different substrates towards BTEXM follows trends observed with 21198 transforming CAHs and 1,4-D reported previously (Bealessio 2021). This trend is reflective of the extent of expression of enzyme(s) capable of cometabolizing these contaminants. Murnane et al. (2021) used activity-based profiling to demonstrate that the alkane monooxygenase responsible for oxidizing CAHs and 1,4-D is expressed highly in isobutane grown 21198, moderately in 2-butanol grown cells, and minimally in 1-butanol grown cells. This is presumably due to the need for the alkane monooxygenase to initiate isobutane metabolism, but not butanol metabolism. Rather, butanol metabolism is initiated by an alcohol dehydrogenase, which cannot initiate cometabolism of BTEXM. A similar explanation for lag periods has been suggested with a Rhodococcus aetherivorans strain grown on 1-butanol (Inoue et al. 2018). Murnane et al. also observed an acceleration in rates of 1,4-D cometabolism with time for 1-butanol grown cells, which is consistent with observations in B and T individual tests shown in Figure 1A and 1B and indicative of increased monooxygenase production with time. The similarity in trends observed across growth substrates for BTEXM, CAHs, and 1,4-D indicates that the alkane monooxygenase involved in CAH and 1,4-D oxidation may also be involved in the oxidation of BTEXM.

3.2. Contaminant Mixture Batch Experiments

3.2.1. BTEX Mixture

Contaminant mixtures were tested with 21198 after confirming its ability to degrade all components of BTEXM individually. The order of contaminant degradation and emergence of lag periods shown in Figure 3 illustrate the impact of contaminant mixtures on 21198, and how that impact differs based on the substrate used to grow the cells.

Fig. 3.

Degradation of BTEX in resting cell mixture batch tests by isobutane (A), 1-butanol (B), or 2-butanol (C) grown 21198. Error bars represent standard deviation between triplicates. Data normalized to initial masses for each contaminant. Initial mass of each contaminant is reported in Table S4

The order of contaminant transformation followed the relative rates of transformation established in the individual contaminant tests, with contaminant degradation commencing as T > E > B > o-X for all growth substrates. As observed in the individual contaminant tests, there was no lag period in TEX transformation with isobutane grown cells, however a short lag period of 1.3 hours was observed for B. A lag period was observed for B, T, and o-X in tests conducted with 1-butanol grown cells, and B and o-X in tests conducted with 2-butanol grown cells (Table 1). For 1-butanol grown cells, this lag period may be representative of combined effects of inhibition and induction. Lag periods were observed for B and T in individual contaminant tests and were therefore expected to be observed in mixture tests as well. The lag periods observed in BTEX mixture tests were similar in length to those observed in individual tests, however the presence of lag periods for contaminants in mixture tests that were not observed in individual tests indicate that other factors, such as inhibition, may be contributing to the lag in transformation. Inhibition by the presence of T and E is a commonly reported phenomenon among BTEX mixture transformation (Jung and Park 2004; Deeb and Alvarez-Cohen 1999). This hypothesis is further supported by the fact that significant B and o-X transformation progressed only after T and E concentrations were significantly reduced. These interactions were also observed with 2-butanol grown cells. For 1-butanol and 2-butanol grown cells, the presence of other aromatic compounds had a positive effect on the extent of o-X degradation, as they were able to fully transform o-X in the mixture tests. Enhanced degradation when BTEX compounds are present in a mixture has been reported for other microorganisms as well (Wongbunmak et al. 2020; Benedek et al. 2021; Khodaei et al. 2017), with authors attributing this observation to microbial growth supported by components of BTEX and/or enhanced expression of enzymes required for BTEX transformation.

3.2.2. BTEXM Mixture

The order of BTEX degradation followed the same pattern as described in the previous section, with MTBE transformation occurring slower than all BTEX components as shown in Figure 4. Abiotic losses in these experiments were minimal and are reported in Table S3. The presence of MTBE in the BTEX mixture impacted transformation rates differently depending on the substrate the cells were grown on, although the rate of B transformation for all growth substrates was not significantly impacted by the presence of MTBE in the BTEX mixture.

Fig. 4.

Degradation of BTEXM in resting cell mixture batch tests by isobutane (A), 1-butanol (B), or 2-butanol (C) grown 21198. Error bars represent standard deviation between triplicates. Data normalized to initial masses for each contaminant. Initial mass of each contaminant is reported in Table S4

For isobutane grown cells, an increase in zero-order transformation rate was observed for T, E, and o-X in the presence of MTBE. The opposite was observed for 1-butanol grown cells, with T, E, and o-X degradation rates decreasing in the presence of MTBE. Interestingly, for 2-butanol grown cells, no significant impact on BTEX transformation rate was observed with the presence of MTBE in the BTEX mixture (Table 1). No lag periods were observed for isobutane grown cells, while B and o-X lag periods observed for 2-butanol grown cells were consistent with observations from the BTEX mixture test. An extended lag period for B and MTBE were observed for 1-butanol grown cells, which was twice as long as that observed in the BTEX mixture test. MTBE degradation commenced once significant transformation of BTEX was achieved by 1-butanol grown cells, suggesting that BTEX inhibits the transformation of MTBE. MTBE inhibition by BTEX is a commonly reported phenomenon (Raynal and Pruden 2008; Deeb and Alvarez-Cohen 2000; Lee and Cho 2009). Similar lag periods were also observed by Zhou et al. (2016) with BTEX and 1,4-dioxane mixtures, with lag being attributed to enzyme competition, intermediate toxicity, and/or energy deficits. Despite the initial inhibition of MTBE transformation, as BTEX concentrations decreased, rapid transformation of MTBE was observed. What’s more, lower concentrations of MTBE were achieved in BTEXM mixture tests than in tests conducted with MTBE alone as shown in Figure 5, despite there being a greater mass of contaminants for 21198 to transform.

Fig. 5.

Comparison of MTBE transformation by isobutane (A), 1-butanol (B), and 2-butanol (C) grown 21198 in individual contaminant tests (diamonds/solid) and BTEXM mixture tests (circles/dashed). Note the y-axis is in log scale. The red line represents the 20 μg/L EPA advisory concentration of MTBE. Error bars represent standard deviation among triplicates

For microorganisms that are able to transform MTBE in the presence of BTEX, several studies have reported no detrimental effect to MTBE transformation ( Pruden et al. 2003; Pruden and Suidan 2004; Sedran et al. 2002). However, it is uncommon for MTBE transformation to be enhanced by the presence of BTEX. Aufrett et al. (2009) reported that MTBE could only be degraded by a novel Rhodococcus wratislaviensis isolate when present in a mixture of aromatic and aliphatic hydrocarbons (Auffret et al. 2009). It was suggested that this is due to enhanced cometabolic activity towards MTBE when present in a mixture with other hydrocarbons. This enhanced activity observed with 21198 could be due to BTEX acting as stronger inducers of relevant enzymes than MTBE alone, or due to the additional energy provided from hydrocarbon metabolism.

3.3. Growth Batch Reactors

A low initial biomass loading, one hundredth of what was added to the resting cell batch reactors, was studied in the growth batch reactors. Startup time, defined as the time required to transform the initial mass of BTEX in the batch reactors, was influenced by the absence or presence of an added growth substrate. Progression of BTEXM degradation followed the same order as observed in BTEXM mixture tests with resting cells. Oxygen consumption and microbial growth, reported in Table S6, support the contaminant transformation and growth substrate utilization trends shown in Figure 6.

Fig. 6.

Degradation of BTEXM and consumption of growth substrate in no substrate (A), isobutane (B), 1-butanol (C), or 2-butanol (D) microcosms in first 7 days post-inoculation. Error bars represent standard deviation among triplicates. Data normalized to initial masses for each contaminant. Initial mass of each contaminant is reported in Table S4

1-butanol was most rapidly utilized, followed by 2-butanol, and then isobutane. O₂ consumption in the no substrate added batch bottles is representative of the oxygen demand of the resting cells and their oxidation of the added contaminants, while the increases in oxygen consumption for isobutane, 1-butanol, and 2-butanol amended batch bottles demonstrate the oxygen demand associated with the resting cells, BTEXM transformation and growth substrate utilization. As expected, optical density increased slightly more in batch bottles with added growth substrate compared to those without substrate. Increases in OD₆₀₀ for all batch bottles indicate that 21198’s growth is not inhibited by the presence of BTEXM. Additionally, an increase in the OD₆₀₀ in bottles that did not receive any additional substrate alluded to the possibility that contaminant metabolism, rather than exclusively cometabolism, was occurring. This was further investigated in individual substrate growth tests described in section 3.4.

Isobutane and 2-butanol amended batch bottles transformed BTEXM similarly to the batch bottles with no added growth substrate. A lag period in isobutane consumption was observed for the first 4 days, during which time T and E were transformed. This preferential transformation of T and E over isobutane utilization may be a result of the lower aqueous concentration of isobutane compared to T and E due to its higher volatility, and/or competition between the compounds for the same enzyme. Despite 1-butanol grown cells consistently exhibiting the lowest transformation rates and capacities towards BTEXM in resting cell tests, the 1-butanol amended batch bottles required the shortest start up time for growth substrate utilization and contaminant transformation. The 1-butanol amended batch bottles indicate the combined advantage of rapid substrate utilization, cell growth, the high enzyme expression associated with the initial addition of isobutane grown cells and maintained enzyme expression, possibly due to the potential inductive effects of BTEX, to promote cometabolic transformation of MTBE. The use of alcohols to support growth and BTEXM transformation also avoids introducing competition between the growth substrate and contaminants, as 1-and 2-butanol do not require initial oxidation by a monooxygenase. The high performance of the 1-butanol amended batch bottles and the increase in OD600 in batch bottles without growth substrate prompted further investigation into the potential for utilization of BTEXM as carbon and energy sources by 21198.

3.4. 21198 Growth Experiments

ATCC Strain 21198’s ability to utilize a wide range of aliphatic growth substrates, including isobutane, 1-butanol, and 2-butanol, has been established previously (Chen 2016). Growth tests conducted with BTEX and MTBE revealed that B and T can be used as the sole carbon and energy source by 21198, but E, X, and MTBE cannot. A summary of these results is reported in Table 2. Therefore, transformations of E, X, and MTBE observed with 21198 cells are indicative of cometabolic processes, whereas those observed with B and T are representative of utilization through metabolic processes. Results from the growth experiments along with KEGG pathways for B and T and the annotated genome of 21198 sequenced previously (Shields-Menard et al., 2014) were used to construct possible metabolic pathways for 21198’s metabolism of B and T, shown in Figure S1. Initial oxidation with a monooxygenase is proposed for B and T, however T metabolism may also be initiated with a toluene hydroxylase. It is hypothesized that T oxidation must be initiated at the methyl group, as growth was observed on benzyl alcohol but not p- or m- cresol. B and T metabolic pathways merge at a catechol intermediate, however it requires several more enzymatic steps for T to be transformed into catechol than B, which is supported by the faster growth observed on B compared to T. 21198 has genes for both a catechol-1,2-dioxygenase and catechol-2,3-dioxygenase, therefore catechol ring cleavage may occur in either the 1,2 or 2,3 position. Enzymes required to breakdown products of either catechol dioxygenase to substrates for glycolysis or the TCA cycle were found to be present in 21198’s genome. Tools such as proteomics, differential gene expression analysis, or gene knockout studies are needed to confirm the proposed metabolic pathways. Growth on B was delayed compared to substrates such as isobutane, 1-butanol, phenol, and benzyl alcohol, which all supported observable growth (OD₆₀₀ > 0.1) within 3 days of incubation. Generally, faster growth was observed with the more oxidized substrates. For example, growth on 1-butanol commenced more rapidly than growth on isobutane. Similarly, growth on phenol or benzyl alcohol was more rapid than growth on B or T. Exceptions are 2-butanol and benzyl aldehyde, which took longer to grow cells than isobutane and T. Growth on B and T by Rhodococcus rhodochrous strains has been reported previously (Deeb and Alvarez-Cohen 1999; Vanderberg et al. 2000; Warhurst et al. 1994), but to the authors’ knowledge this is the first demonstration of B and T metabolism by 21198. Cometabolism of E, X, and MTBE likely involves the same enzymes used in the beginning steps of the B and T metabolic pathways shown in Figure S1. However, because growth was not observed on these substrates, the cometabolic transformations likely lead to dead end products. Identifying dead end products that accumulate during BTEXM degradation would inform the extent of transformation and enzymes involved in the cometabolism of E, X, and MTBE.

Table 2.

OD₆₀₀ measurements from 21198 growth experiments reported as OD₆₀₀ < 0.1 (−), OD₆₀₀ > 0.1 (+), OD₆₀₀ > 0.3 (++), OD₆₀₀ > 0.6 (+++) among triplicates or duplicates (denoted with an asterisk). Substrates that supported 21198 growth are bolded.

Growth Substrate	Day 3	Day 7	Day 14
Isobutane	+	+++	+++
1-Butanol	++	++	++
2-Butanol	−	+	+++
Benzene	−	++	++
Toluene	−	+	++
Ethylbenzene	−	−	−
o-Xylene	−	−	−
m-Xylene*	−	−	−
p-Xylene*	−	−	−
Phenol	+	++	++
m-Cresol	−	−	−
p-Cresol*	−	−	−
Benzyl Alcohol	++	++	++
Benzyl Aldehyde	−	−	+
Benzoic Acid*	+	++	++
MTBE*	−	−	−
Control	−	−	−

The identification of these novel growth substrates corroborates the findings of enhanced contaminant transformation in BTEX and BTEXM mixture tests. Energy gained from B and T metabolism likely supported the cometabolism of cometabolic substrates such as o-X and MTBE. This is reflected in the greater transformation capacity of o-X and MTBE observed in mixture tests compared to the individual contaminant tests. These findings also explain the increase in optical density in the growth batch reactors that were not supplied additional growth substrates (Table S6). The increase in OD₆₀₀ measured in these bottles is the result of growth from B and T metabolism, while the greater OD₆₀₀ in the growth substrate amended bottles is the result of the combined growth on B, T, and either isobutane, 1-butanol, or 2-butanol.

4.0. Conclusions

The results demonstrate that 21198 is capable of degrading all components of BTEXM individually and in mixtures using a combination of metabolic and cometabolic processes. While isobutane, 1-butanol, and 2-butanol all supported 21198’s growth and degradation of BTEXM, resting 21198 cells’ performance was optimized by growth on isobutane and minimized by growth on 1-butanol. However, 1-butanol was found to be an effective substrate for supporting the growth of 21198 in the presence of BTEXM as it is most rapidly utilized by 21198 compared to isobutane and 2-butanol, and likely does not compete for enzymes involved in BTEXM oxidation. Positive outcomes such as increased rate and extent of degradation followed the initial lag periods that were observed in BTEX and BTEXM mixture tests. 21198’s ability to degrade all components of BTEXM, particularly B and T, allows for more rapid and complete MTBE transformation. This work demonstrates the possible utility of primary and secondary alcohols to support biodegradation of BTEXM in batch studies that can be expanded to continuous flow treatment studies in the future. The results also illustrate complex interactions of B and T metabolism and cometabolic transformation of MTBE that require further investigation. Finally, the utility of 21198 for bioremediation applications has been expanded to include MTBE, TBA, and monoaromatic hydrocarbons.