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Critical factors for microbial contamination of domestic heating oil Bernd Leuchtle, Wei Xie, Thiemo Zambanini, Simon Eiden, Winfried Koch, Klaus Lucka, Martin Zimmermann, and Lars M. Blank Energy Fuels, Just Accepted Manuscript • DOI: 10.1021/acs.energyfuels.5b01023 • Publication Date (Web): 11 Aug 2015 Downloaded from http://pubs.acs.org on September 3, 2015
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Critical factors for microbial contamination of domestic heating oil
Bernd Leuchtle†, Wei Xie†, Thiemo Zambanini†, Simon Eiden††, Winfried Koch††, Klaus Lucka††, Martin Zimmermann*†, Lars M. Blank †
†Institute of Applied Microbiology - iAMB, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, 52074 Aachen, Germany ††Oel-Waerme-Institut - OWI - Affiliated institute RWTH Aachen, 52134 Herzogenrath, Germany
Corresponding author: Martin Zimmermann E-mail:
[email protected] Address: Institute of Applied Microbiology - iAMB, ABBt - Aachen Biology and Biotechnology, RWTH Aachen University, 52074 Aachen, Germany Tel.: +49-2418026617
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Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.
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ABSTRACT
Microbial contamination can occur during long-term storage of domestic heating oil (DHO), thus biofouling the fuel and equipment. In this study, microorganisms in domestic heating oil storage tanks were identified, and the factors that promote growth in DHO storage tanks were examined. Notably, obligate aerobic but not strict anaerobic microbes were detected, suggesting the existence of an oxic environment. The measured and calculated diffusion rates of oxygen in the oil support the finding that oxygen limitation in a storage tank is unlikely. Fatty acid methyl ester (FAME) fostered microbial growth in DHO tanks, and fewer microorganisms were required to initiate growth in FAME-blended fuels. The results are discussed in the context of minimizing microbial contamination during long-term storage of FAME blends in domestic heating oil applications.
Keywords: domestic heating oil, biodiesel, microorganisms, contamination, long-term storage stability
INTRODUCTION Microbial contamination of fuels is a problem for producers and consumers. For example, microbiological contamination creates issues for diesel-powered military engine systems.1 Domestic heating oil (DHO) is a middle distillate fuel. Microbial growth in fuel storage tanks depends on the accumulation of water from condensation, phase separation of dissolved and dispersed water, the ability to degrade a broad range of molecules in middle distillates, and storage time.1, 2 The latter has garnered particular interest in domestic heating oil applications because certain components have low degradation rates.3
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Worldwide, approximately 40 billion liters of domestic heating oil are used annually, 67% of which is consumed by private households.4, 5 Most domestic heating oil is used in decentralized domestic heating oil installations (e.g., USA: 6.5 million, Germany: 6 million),6,
7
which are
often kept inside for months or years at temperatures that facilitate microbial growth. In addition, the longer storage time increases the likelihood of developing a free water phase. In contrast to company-owned service stations, households do not normally perform regular maintenance on their tanks. Biomass growth can lead to operating failure or the destruction of storage tanks and parts because microbial biofilms can block filters, and organic acids produced by microbes accelerate corrosion.8, 9 These living organisms consist primarily of C (48%), O (24%), N (14%), H (7%), P (3%), and S (0.5%) (in w/w), uptake of which is required by microbes for survival.10 In a storage tank, these and all other required nutrients originate from the fuel (Table 1).
By using DHO as a growth substrate, microbes can negatively affect oil properties.11 Microbial contamination can also affect the formation and stabilization of a water-in-oil emulsion resulting from the secretion of emulsifiers.12 The use of renewable energy is promoted by many governments worldwide to reduce CO2 emissions,
for example by blending biofuels—such as FAME—with regular fuels.13 For
domestic heating oil, this substitution is possible using biodiesel. However, the addition of FAME imposes technical challenges, including changes in fuel stability and heating values.14 Further, microorganisms degrade biodiesel more easily than fossil fuels.15, 16 Several studies have characterized the microorganisms that degrade fuel components.17,
18
Because an individual organism can often degrade only a single component, more thorough
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utilization of fuels as a carbon source requires a mixture of microorganisms.19, 20 Such mixed cultures can produce biofilms and emulsifiers that increase the water-oil interphase and fuel degradation rates due to the greater surface-to-volume ratio.21, 22 In this study, we examined the factors required for microbial contamination of heating oil tanks and the organisms involved to develop strategies for minimizing undesired biofouling. Microbial development in contaminated DHO tanks was simulated in long-term storage experiments. The effect of free water on microbial growth in various domestic heating oil-FAME blends was measured, and the influence of storage temperature and microbial cell number was determined. The results are discussed in the context of possible technical solutions to minimize biofouling of storage tanks and equipment damage.
MATERIAL AND METHODS Microorganisms The organisms used in this study were provided by Schülke & Mayr GmbH (Norderstedt, Germany) as an uncharacterized, mixed liquid culture of eukaryotes and prokaryotes isolated from a diesel fuel service station and enriched during growth in mineral medium. To identify the microbiome of DHO storage tanks, fuel samples were taken from 10 different tanks located in Aachen and Hamburg, Germany. Three samples were taken per tank (directly beneath the fuel-surface, at half of the filling height and directly above the bottom of the storage tank) by drawing 1 L of fuel through a sterile tube in a sterile bottle. After filling, the bottles were closed directly and stored in the dark at 4°C until further usage. To enrich biomass and simplify DNA extraction, the 31 gathered samples (1 provided mixed culture, 10 tanks with 3 samples each) were used to inoculate 5 different growth media
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(standard I [15 g/L peptone, 3 g/L yeast extract, 6 g/L NaCl, 1 g/L glucose], yeast extract peptone dextrose medium (YPD) [20 g/L peptone, 10 g/L yeast extract, 20 g/L glucose], mineral medium [1 g/l (NH4)2SO4, 0.1 g/l MgSO4 * 7H2O, 0.5 g/l KH2PO4, and 0.76 g/l K2HPO4, supplemented with trace elements (100 µg/l ZnSO4, 300 µg/l H3BO3, 134 µg/l CaCl2 * 2H2O, 2000 µg/l FeSO4 * 7H2O, 10 µg/l CuCl2 * 2H2O, 30 µg/l Na2MoO4 * 2H2O, 20 µg/l NiCl2 * 6H2O, 30 µg/l MnCl2 * 4H2O)]+ DHO, mineral medium + B5, and mineral medium + B20). After cultivation, total DNA was extracted, and 16S or 18S rDNA was amplified. The
primer
pair
U789F
(TAGATACCCSSGTAGTCC)
and
E1541R
(AAGGAGGTGATCCANCCRCA) was used to amplify the 16S rDNA variable regions V5-V9.23-25
The
primer
pair
NS17
(CATGTCTAAGTTTAAGCAA)
and
NS20
(CCCTATTAATCATTACG) was used to amplify the 18S rDNA variable regions V1–V3.26, 27 The concentrations of PCR products were adjusted, and the products were combined and sequenced on the Roche GS Flx 454 platform (GATC Biotech, Konstanz, Germany) and characterized by genus.
Oxygen Transfer Oxygen transfer was analyzed with an oxygen probe (Oxyferm FDA 225, Hamilton, Reno, USA) that was placed in a measuring cylinder filled with DHO, DHO-FAME blends, and, if indicated, water. After outgassing all available oxygen with nitrogen, the amount of oxygen that diffused through the DHO/water was measured. Measurements were made at the bottom of the cylinder in the DHO phase or, if added, in the water phase (beneath the DHO, if both were present) 10 cm from the surface.
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Storage Experiments To mimic real-life storage conditions, lab equipment was not sterilized prior to the experiments, except for preculture materials and storage bottles. The DHO [low-sulfur heating oil (sulfur < 50 ppm)] fulfilled DIN 51603-1 criteria.28 The DHO from crude oil was designated B0 (zero biodiesel). The FAME (rapeseed methyl ester - RME) was produced per DIN EN 14214.29 The blends were generated per DIN SPEC 51603-6 by mixing pure DHO and FAME.30 DHO-FAME blends were designated B5 (95% (v/v) DHO + 5% (v/v) RME), B7 (93% (v/v) DHO + 7% (v/v) RME), B10 (90% (v/v) DHO + 10% (v/v) RME), and B20 (80 % (v/v) DHO + 20% (v/v) RME). These blends were chosen because a 5% biofuel substitution can be achieved in nearly all heaters without technical changes, whereas a 7% blend is used as a substitution for diesel. B20 fuel is the highest biodiesel substitution (depending on the DHO) that does not undercut current limits.30 Further, B20 can be used in many existing heaters without the need for technical changes.31 RME was used exclusively throughout this study because it is the chief FAME in Europe.32 The fuels and their usage are listed in Table 2.
The biofuel blends for all experiments (including precultures) were prepared by adding 1 L water to 10 L fuel and stirring the mixture for 1 h at 150 rpm to ensure water saturation (approximately 50 ppm in pure DHO, 90 ppm in B5, and 200 ppm in B20). For all experiments, only the water-saturated fuel was used, while the remaining water phase was discarded. The microbial inoculum was precultivated in mineral medium for 5 days at 28°C, with three DHO-FAME blends (B0, B5, and B20) serving as the sole carbon source. The 3 cultures were
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pooled, and the cells were washed and lyophilized, if required; 500 µl of the pooled liquid preculture was used for lyophilization. Prior to inoculation of the storage bottles, the concentration of microorganism was determined by optical density using a spectrophotometer (Ultrospec III, Pharmacia Biotech, Uppsala, Sweden) at a wavelength of 600 nm (OD600). Also, different biomass amounts from OD600 0.05 to 0.7 were investigated (Figure 3) by diluting the preculture to the desired OD600. Alternatively, to determine colony-forming units (Figure 2), cells were plated on rich media (YPD). For the experiments, sterile 1-L glass bottles were filled with 500 mL fuel containing 0% to 20% (v/v) FAME. When indicated, 500 µL, 5 mL, or 50 mL of a free water phase (0.1% (w/v) NaCl) were added to the oil. No nutrient-promoting growth media was used in the storage experiments. The fuel was inoculated directly from a washed preculture or with lyophilized cells as indicated. Lyophilized cells were resuspended in 500 µL distillated water or, in samples without a free water phase (Figure 2A), transferred as a water-free powder. In experiments to determine whether the amount of dissolved water was sufficient for microbial growth in DHO and DHO-FAME blends, 4 replicates per condition (i.e., FAMEcontent, free water phase/water-saturated DHO) were made and inoculated with lyophilized cells at the beginning of the experiment. After 3 and 35 days, the cell dry weight was determined as indicated below. To examine whether microbial growth occurred in fuel or water, only the DHO was filtered in water-saturated samples without an additional free water phase, whereas if a free water phase was present, it was filtered exclusively. Samples for one data point comprised 2 1-L bottles. Two bottles were sacrificed at regular intervals (1-3 times per week) to determine the cell dry weight (CDW). The bottles for a complete experiment (up to 150 bottles) were filled and treated simultaneously prior to storage.
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The bottles were kept open in the dark for 5 to 11 weeks at 20°C ± 0.1°C or, in experiments in which the influence of cultivation temperature was examined, at 8°C ± 0.1°C in an incubator (IPP800, Memmert, Schwabach, Germany) to simulate various domestic heating oil tank conditions (i.e., household and underground storage). The open bottles also mimicked the open fuel tank design including aging of DHO because of the loss of volatiles and the reported oxidation of biodiesel.33 The CDW was determined by passing the fuel and water phases separately through pre-dried and pre-weighed 0.4-µm glass fiber filters (MN-GF 5, Macherey Nagel, Düren, Germany). The filters were washed with Tween-20 and water and were dried overnight at 100°C. CDW was measured using a moisture analyzer (MAC 50/1 NH, Radwag, Radom, Poland). Values represent
the mean of 2 dry weights. Error bars indicate the deviation from the mean ( ∑| − ̅ |). Statistical significance was determined by unpaired t-test (mean, standard deviation, n), and the threshold for significance was set to P
