Abstract Bioremediation involves microbial metabolism of harmful substances. This study was undertaken to enhance oil-spill bioremediation through the addition of sodium sulfate, sodium nitrate, and sodium phosphate to samples containing Pseudomonas fluorescens and Bacillus subtilis. Since these bacteria often utilize nitrates during cellular respiration and nitrate ammonification, respectively, it was hypothesized that sodium nitrate would increase the amount of hydrocarbon metabolites present in samples more than sodium sulfate or sodium phosphate. Samples containing different combinations of bacteria, .1 and .01 molar solutions of the above nutrients, salt water, and crude-oil-substitute hexadecane (utilized to replicate the long hydrocarbon chains found in petroleum) were shaken at 27°C for seven or 14 days. Bacterial concentration was estimated by turbidity measurements. Each organic layer was analyzed with gas chromatography to identify any smaller hydrocarbons.The data from two trials partially disprove the hypothesis. While the treatment of .1 M sodium nitrate caused one of the greatest increases in bacterial concentration with P. fluorescens and B. subtilis, the chromatographic data from these treatments differed only marginally from the positive controls and other treatments. Additionally, the data suggested that substantial metabolite formation occurred only in P. fluorescens samples with .1 M and .01 M treatments of sodium phosphate. However, the .1 M nutrient combination and .01 M sodium sulfate treatments caused the greatest hexadecane disappearance. These results are relevant to the enhancement of bioremediation techniques; a need exemplified by the Deepwater Horizon catastrophe.

IntroductionMarine oil spills represent one of the most devastating accidents occurring worldwide, as demonstrated by the Deepwater Horizon explosion on April 20, 2010. Preventing these spills entirely would be practically impossible; however, preparing for them can limit the damage they cause. Increasing the rate at which oil spills in the ocean can be cleaned would save millions of organisms and acres from lasting harm. This project was completed to increase the effectiveness of bioremediation techniques by the addition of inorganic nutrients to Pseudomonas fluorescens and Bacillus subtilis for use during oil spills.

According to the EPA, almost 14,000 oil spills are reported each year1. Many different techniques are used to attempt to clean oil spills. Booms–barriers that extend about three feet below the surface of the water can be used to contain the oil, while

The Effect of Nitrogen, Sulfur, and Phosphorus Compounds on Bioremediation of Oil Spills by Pseudomonas fluorescens and Bacillus subtilisMeghan Shea1*, Sandra Litvin2, and Anastasia Chirnside3

Student1, Teacher2: Unionville High School, Kennett Square, PA 19348Mentor3: University of Delaware, Newark, DE 19716*Corresponding author: meghan.shea28@gmail.com

INTERNSHIP ARTICLE

skimmers act as vacuums to remove this contained oil1. Sorbents, or materials that absorb or adsorb oil, are also commonly used to remove the final traces of oil from an oil spill1. However, the technique that requires the least human effort and is often the most successful in the long run is bioremediation1.

According to the Civil Engineering Department of Virginia Tech, bioremediation is “the application of biological treatment to the cleanup of hazardous chemicals in the soil and surface or subsurface waters”2. Certain bacteria are able to feed on the contamination, which in this case is long hydrocarbon chains, and use the energy for growth and reproduction2. Through this process, the oil is metabolized into water and carbon dioxide. While this process usually occurs naturally, it can be stimulated through the addition of nutrients and microbes, which is known as bioaugmentation. Common organisms used in the bioaugmentation of petroleum are Pseudomonas, Proteus, Bacillus, Penicillum, and Cunninghamella.

For this experiment, two rod-shaped, aerobic, heterotrophic microbes, Pseudomonas fluorescens and Bacillus subtilis, were used to degrade oil. P. fluorescens colonizes soil, water, and plant surface environments and is motile due to multiple polar flagella3. This Gram-negative bacterium can utilize nitrate as an electron accepter in place of oxygen during cellular respiration3. B. subtilis, a Gram-positive bacterium with peritrichous flagella, also inhabits soil and water4.

In place of crude or motor oil, both of which contain additives that react with different aspects of this experiment making data collection difficult, HD (see figure 1 for abbreviations and acronyms) was utilized to replicate the long hydrocarbon chains found in petroleum. HD is an alkane hydrocarbon with a 16-carbon chain.

Often, nitrogen, phosphorus, and sulfur are the limiting factors of bioremediation in aquatic environments. Therefore, sodium nitrate (NaNO3), sodium phosphate (Na3PO4), and sodium sulfate (Na2SO4) were used as sources of the above nutrients in order to determine which has the greatest effect on the bioremediation of hydrocarbons by P. fluorescens and B. subtilis. All three inorganic nutrients are soluble in water, making them effective for aquatic oil spills. In P. fluorescens,

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Figure 1. Abbreviations and acronyms.

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Materials and MethodsInoculating liquid cultures of P. fluorescens and B. subtilis: One liter of nutrient broth was prepared using the standard procedure found on the container and sterilized using an autoclave. 150 ml of nutrient broth were added to each of three clean Erlenmeyer flasks labeled B. subtilis, P. fluorescens, and Spectronic 20 control, respectively. The three flasks were autoclaved. The flask labeled P. fluorescens was inoculated with bacteria from a pure colony from Carolina Biological, using a metal inoculating loop and standard flaming techniques to avoid contamination. The inoculation was repeated twice more with P. fluorescens, re-inoculating the same flask to ensure adequate bacterial growth. The above procedures were repeated with the B. subtilis pure colony and the flask labeled B. subtilis. All three flasks – two inoculated and one control – were incubated at 27ºC. Creating a standard spec-20 absorbance curve for each bacterium: A Spec-20 was set to a wavelength of 686 nm and left to warm up for a minimum of 15 minutes. Eleven clean culture tubes were obtained and labeled 1-11. Using different pipettes, turbid P. fluorescens liquid culture and nutrient broth were added to each tube to a total volume of 10.0 ml. The volume of liquid culture was reduced by 1.0 ml in each tube from 10.0 ml to 0.0 ml. Using a vortex mixer, each tube was mixed thoroughly for 1 – 2 seconds. After mixing, 3-4 ml of broth from tube 11 were transferred to a clean Spec-20 tube and the outside of the tube was wiped with a Kimwipe. The tube was placed in the Spec-20 and used as a blank, following the instructions for the specific machine. Starting with tube 10 and going backwards to tube 1, 3-4 ml of each dilution were transferred to the Spec-20 tube and the absorbance was measured. Using the absorbance data, a X-Y scatter plot of dilution factor vs. absorbance was created and a second order polynomial trend line was added to the plot. From the plot, the range over which absorbance was proportional to bacterial concentration was identified, and a second plot was created using only this range. A linear trend line was added to this

nitrate is often incorporated in metabolism by serving as an electron acceptor during cellular respiration3. B. subtilis uses nitrate ammonification in anaerobic conditions4. However, the specific use of these nutrients during the hydrocarbon metabolism of P. fluorescens and B. subtilis has not yet been studied. Since the two bacteria utilize nitrate during cellular respiration in anaerobic conditions and nitrate ammonification, respectively, it was hypothesized that sodium nitrate, an inorganic nutrient that provides nitrogen to the microbes, would cause the greatest HD degradation and production of smaller hydrocarbons.

To test this hypothesis, samples were prepared containing different combinations of bacteria, .1 and .01 molar solutions of the above nutrients, salt water, and crude-oil-substitute hexadecane. The turbidity of each sample was measured daily using a Spec-20. An absorbance standard curve for each bacterium coupled with serial-dilution plating facilitated the conversion of absorbance to CFU/ml. After shaking at 27°C for 7 or fourteen days, the organic layer was removed from each sample using an Acetonitrile-based Solid Phase Extraction. Each organic layer was analyzed with gas chromatography (FID) to identify any smaller hydrocarbons, the presence of which would indicate successful bioremediation.

new plot. Using the equations of the trend lines from the first and second plots, a correction formula was derived to correct for linearity at high absorbance readings. The entire procedure above was repeated with a turbid B. subtilis culture to obtain a second corrected absorbance curve. Calibrating absorbance measurements with viable cell counts: One liter of nutrient agar was prepared using the standard procedure found on the container, sterilized using an autoclave, and poured into sterile Petri dishes. 16 sterile culture tubes were obtained and labeled 10-1 – 10 -16. Using a 10-ml pipette, 9 ml of sterile nutrient broth were added to each culture tube. 32 nutrient agar Petri dishes were obtained and two were labeled 10-1, two 10-2, etc., through 10-16. Using a Spec-20 and the procedure detailed above, the absorbance of a turbid P. fluorescens culture was recorded. Using a 1-ml micropipette, 1 ml of turbid P. fluorescens culture was transferred to the 10-1 tube. The dilution was mixed thoroughly by vortexing for 1 – 2 seconds. Using a new pipette tip, 1 ml of the 10-1 dilution was withdrawn and transferred to the 10-2 tube. The dilution was mixed using a vortex, and the transferring technique above was repeated until all serial dilutions were made. Using a new pipette tip and starting with the 10-16 dilution, 0.1 ml of the dilution was pipetted onto the appropriately labeled nutrient agar plate and immediately spread with a sterile plate spreader. This procedure was repeated with the duplicate 10-16 plate and all subsequent dilutions. All plates were incubated at 27ºC for 48 hours, at which time the colonies of plates with 25 – 250 colonies were counted. Using the colony counts, the colony forming units per ml of the original culture was determined. This value was then divided by the corrected absorbance of the culture to obtain a calibration factor used to convert from absorbance to concentration. The above procedures were then repeated with turbid B. subtilis culture to obtain a second calibration factor. Creating .8 molar solutions of sodium nitrate, sodium phosphate, sodium sulfate, and a combination of the three: Using an analytical balance, 11.364 g of Na2SO4 was measured and added to a 100-ml volumetric flask along with approximately 90 ml of distilled water. The mixture was heated and stirred until the Na2SO4 dissolved completely, and which point distilled water was added to reach a final volume of 100 ml. The above procedures were repeated using 30.402 g of Na3PO4 • 12 H2O and 6.798 g NaNO3 to finish making .8 M solutions. The above procedures were also repeated using 3.835 g Na2SO4, 2.290 g NaNO3, and 10.261 g Na3PO4 • 12 H2O in the same flask to make the combination nutrient solution. All nutrient solutions were autoclaved. Plating Pseudomonas fluorescens and Bacillus subtilis: A 100 µ1 sample of turbid P. fluorescens and turbid B. subtilis cultures, respectively, were transferred to separate sterile agar Petri dishes using standard flaming techniques to avoid contamination. The Petri dishes was spun to spread the culture evenly and placed in an incubator at 27°C. Gram staining P. fluorescens and B. subtilis (figure 2): Using a sterile inoculating loop, 1 loop of P. fluorescens was smeared on a clean microscope slide. The slide and smear were flamed briefly using a Bunsen burner. The smear was covered with crystal violet, the primary stain, for 20 seconds, at which time the smear was rinsed using a wash bottle of distilled water. The smear was then covered with Gram’s iodine solution for one minute, at which time the smear was flooded with isopropyl alcohol until the solvent flowed colorlessly from the slide. Rinsing the slide with a

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wash bottle for a few seconds stopped the action of the alcohol. Then, the smear was covered with safranin, the secondary stain, for 20 seconds. The smear was washed gently for a few seconds, blotted with bibulous paper, and let dry at room temperature. The above procedures were repeated with 1 loop of B. subtilis. Both slides were viewed under a microscope using oil immersion. Preparing samples: 500 ml of Instant Ocean were prepared using the accompanying standard procedure and sterilized using an autoclave. 34 Spec-20 tubes and corresponding stoppers were labeled 1-34 and sterilized through submersion in alcohol. Using aseptic technique, the non-bacterial components in (figure 3) were added to the respective samples using the appropriate pipettes and micropipettes. After the non-bacterial components were added, the absorbance of each tube was determined using a Spec-20 at 686 nm blanked with distilled water. Using aseptic technique, the bacterial components from (figure 3) were added to the respective samples using a 500-µ1 micropipette sterilized with alcohol with sterile tips. The absorbance of each tube was determined once more using a Spec-20 at 686 nm blanked with distilled water. Setting up experimental conditions: The 34 samples were secured in an orbital shaker set at 27°C and 80 RPM. The flasks shook for 14 days, destructively sampling on day 7 and day 14. Every four days, the caps of each tube were quickly removed and replaced to aerate and samples. Every day, the absorbance of each sample was measured using a Spec-20 at 686 nm blanked with distilled water. On day 7 and day 14, the control samples were plated and Gram-stained using procedure described above. Separating waste and hydrocarbon layers using Solid Phase Extraction (SPE): 24 test tubes were placed in an Absorbex (a SPE apparatus) set to waste, and 24 Strata X columns (33-µ polymeric reversed phase, 300 mg) were attached to the top of the Absorbex. To condition, 2 ml of Acetonitrile was run through each column. To equilibrate, 2 ml of water were run through each column. At this point, the Absorbex was changed to collect. 24 samples were run through each labeled column at a rate of 1-2 ml per minute. A few samples did not filter through the columns adequately under vacuum so they were left overnight under vacuum to finish loading. Once the samples were loaded onto the columns, the samples were washed with 2 ml of 40:60 ethanol:water (v/v). Then, the samples were dried for 10 minutes under full vacuum. At this point, the Absorbex was changed to collect and samples were eluted with 4 ml of Acetonitrile. The sample test tubes were removed from the Absorbex, placed in a 40°C water bath and evaporated to dryness under a gentle stream of nitrogen gas. Then, the samples were brought back up into 5 ml of hexane, diluted (0.2 ml sample: 1 ml hexane) into GC vials and capped. The above procedures were repeated until all samples were separated. Analyzing samples using GC (figure 4): 1 microliter of each sample was injected into an Agilent 6890 N Network GC system (University of Delaware) with a Flame Ionization Detector and Agilent 25-m long x 0.2 mm diameter ultra-2 column with a .33-µ1 coating of (5% phenyl)-methylpolysiloxane. The samples were injected with a split ratio of 1/50. The oven was ramped at 8°C/min from 70°C to 200°C and then ramped at 30°C/min from 200°C to 280°C. The gas chromatograms produced were then analyzed using the Sherlock program. Fatty Acid Methyl Ester testing to verify identity of bacteria (adapted from [5]): For FAME analysis, one loop of bacterial cells from the isolated colonies on each plate was added to a 20-ml test tube and closed with a Teflon-lined screw cap. Each tube received 1 ml of saponification reagent and was mixed (vortex mixer) and heated at 100°C for 30 minutes to liberate fatty acids from the cellular lipids. Once cooled, each tube received 2 ml of methylation reagent, was mixed again and heated at 80°C for 10 minutes to form methyl esters of the fatty acids. The FAMEs were extracted from this solution by adding 1.25 ml of one part methyl-tert-butyl ether in one part hexane (v:v) (ultrapure grade, VWR Scientific Products, Bridgeport, NJ) and placing the closed tubes on an end-over-end mixer for 10 minutes. Finally, each tube was centrifuged for 10 min at 10,000g, the bottom aqueous layer removed with a pasteur pipette and the supernatant washed with 3 ml of dilute NaOH (0.27M NaOH). The extracted fatty acids were analyzed by a Hewlett Packard (HP) gas chromatograph (Wilmington,

Figure 2. A) Gram-stained B. subtilis under oil immersion before trial. B) Gram-stained B. subtilis under oil immersion after trial. C) Gram-stained P. fluorescens under oil immersion before trial. D) Gram-stained P. fluorescens under oil immersion after trial.

Figure 3. Contents of samples expressed in milliliters.

Figure 4: Sample gas chromatogram showing hexadecane peak, solvent peak, and a zoom view of the area where degradation peaks would be found.

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ResultsIn P. fluorescens samples from both trials, .1 and .01 M sodium nitrate treatments caused the greatest increase in bacterial concentration, while .1 and .01 M sodium sulfate treatments and the .1 M sodium phosphate treatment caused minimal change in concentration in trial one and trial two, respectively (figure 5). All samples containing B. subtilis exhibited far less growth than the corresponding samples with P. fluorescens (figure 6). While P. fluorescens samples increased by between 5x107 and 6x108 CFU/ml, B. subtilis samples only increased by

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DE) equipped with a HP Ultra 2 Cross-linked 5% phenyl methyl Silicone column (250 mm x 0.2 mm x 0.33 µm) and a FID using H2 as the carrier gas. The injection and FID temperatures were 250 and 300°C, respectively. The oven temperature was increased linearly from 170 to 300°C at 5°C min-1. The FAME profile was analyzed utilizing the MIDI, Inc. eukaryote program that covers the fatty acids with a larger range of chain length compared to the aerobe program. Fatty acids with chain lengths exceeding twenty carbons (characteristic of higher organisms, e.g., plants) were not considered in the analysis in an effort to focus strictly on microbial organisms.

Figure 5. A) Graph of Time vs. Change in bacteria concentration for 14 day P. fluorescens samples in Trial 1. B) Graph of Time in bacteria concentration for 14 day P fluorescens samples in Trial 2.

Figure 6. A) Graph of Time vs. Change in bacteria concentration for 14-day B. subtilis samples in Trial 1. B) Graph of Time vs. Change in bacteria concentration for 14-day B. subtilis samples in Trial 2.

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DiscussionIn both the seven-day and fourteen-day data from trial 1 and the seven-day data from trial 2 for both bacteria, the control with no HD or nutrients as well as the control with no nutrients showed the greatest change in concentration of bacteria. This suggests that the addition of nutrients, while perhaps increasing the degradation of HD, actually limits bacterial growth. It also suggests that the addition of HD, thought to be the only source of carbon in the samples, does not dramatically increase bacterial growth. Therefore, the bacteria were potentially metabolizing some component of the IO and not necessarily the HD. The nutrients found in the IO may have also contributed to the variation. The IO is created to replicate an ocean environment, which contains some quantities of dissolved nutrients. However, the IO package failed to list the nutrients present in the mixture. If the mixture contained sources of nitrogen, sulfur, or phosphorus, the addition of these nutrients as treatments may have been less effective. In fact, the nutrients may have been present in concentrations that were harmful to the bacteria, which could account for the decreased concentration of many

between 5x104 and 6x105 CFU/ml. In B. subtilis samples, a .1 M sodium nitrate treatment caused the greatest increase in bacterial concentration and no samples caused substantially reduced growth.

Qualitative analysis of the GC data failed to parallel the bacterial concentration data. Since a mass spectrum detector was not used with the gas chromatograph, a calibration standard containing octane, decane, dodecane, and tetradecane was run using the operating procedure described in the materials and methods section (figure 7). Then, the retention times of the degradation peaks found in samples could be compared to the retention times of the peaks in the calibration standard, allowing for tentative and basic identification. Gas chromatograms from trial one showed very few degradation peaks, possibly due to contamination from the rubber stoppers used in experimentation. Only the fourteen-day treatment of B. subtilis and .1 M sodium phosphate showed minimal degradation into an 11-carbon compound. However, gas chromatograms from trial two had 14-carbon, 13-carbon, and 12-carbon degradation peaks visible in all P. fluorescens samples and some B. subtilis samples. After seven days, all of the P. fluorescens treatments showed similar degradation peaks, which were smaller than the peaks in the control sample without nutrients; also, there was negligible difference between .1 M treatments and their .01 M counterparts. In contrast, all B. subtilis treatments showed slightly larger degradation peaks than the control without nutrients after seven days.

After fourteen days, P. fluorescens treatments of .1 M sodium phosphate and .01 M sodium phosphate showed the largest degradation peaks, while all P. fluorescens treatments showed an increase in degradation from the control without nutrients. Surprisingly, all three of the .01 M treatments showed greater degradation than their .1 M counterparts, bolstering the theory that the nutrients were present at too high a concentration. Of all the P. fluorescens treatments, the .1 M nutrient combination showed the least degradation; since this treatment provided the benefits of all three nutrients, it was expected to showed more degradation than some of the treatments with only one nutrient. Of the B. subtilis treatments, only one, .1 M sodium phosphate, showed degradation.

Initial quantitative analysis of the GC data also provided interesting insight (figure 8). Using a standard HD curve, the area of the HD peaks of each sample could be converted to the concentration of HD extracted from each sample. Percent recovery was determined by running spiked matrix samples (5 HD concentrations, 7 times) through the extraction procedure. While this process only yielded about a 20% HD recovery rate, it showed some degradation trends. For P. fluorescens, the controls without nutrients had the greatest HD degradation and their autoclaved counterparts typically had the smallest HD degradation, as expected. In P. fluorescens samples, treatments of .01 M sodium sulfate and .1 M nutrient combination had the greatest degradation, while in B. subtilis samples, treatments with .1 M sodium nitrate and .1 M nutrient combination had the greatest degradation.

Figure 7: Gas chromatograph from 100 ppm 8, 10, 12, 14-Carbon calibration standard run under standard conditions.

Figure 8. A) Hexadecane degradation (mg/L) by P. fluorescens in Trial 1 samples, B) Hexadecane degradation (mg/L) by B. subtilis in Trial 1 samples, C) Hexadecane degradation (mg/L) by P. fluorescens in Trial 2 samples, D) Hexadecane degradation (mg/L) by B. subtilis in Trial 2 samples.

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treatments in comparison to controls. Along with variation from initial expectations, the data also showed variation between trials. In the seven-day P. fluorescens trial 1,

a treatment of .1 M sodium sulfate showed significant change in concentration. However, the identical treatment in the trial 1 fourteen-day trial showed negligible change. In the trial 1 seven-day B. subtilis trial, a treatment of .1 M sodium nitrate showed negative change in concentration. However, the identical treatment in the trial 1 fourteen-day trial showed significant change. Trial 2 also had similar, but less obvious, discrepancies.

A common theme throughout all trials was that samples containing B. subtilis experienced far less growth than corresponding samples of P. fluorescens, which supports conclusions made in previous years of experimentation. While P. fluorescens samples changed by between 5x107 and 2x108 CFU/ml, B. subtilis samples only changed by between 5x104 and 1.5x105 CFU/ml. This may have occurred for a variety of reasons. P. fluorescens. and B. subtilis are both aerobic bacteria, meaning that they require oxygen to survive and metabolize. However in anaerobic conditions, P. fluorescens can use nitrate as an electron acceptor in place of oxygen during cellular respiration. While the flasks were aerated every four days, the bacteria may have used oxygen faster than it was being introduced, leading to anaerobic conditions. Since nitrates were most likely present in IO as well as the sodium nitrate treatment, the P. fluorescens may have been better able to survive anaerobic conditions than B. subtilis. This variation could have been avoided by aerating the flasks more frequently. Another potential explanation for the differences in results between P. fluorescens and B. subtilis is that B. subtilis needs a longer period of time to metabolize long hydrocarbons. Since the trial was only fourteen days and bioremediation often takes many years in ocean environments, B. subtilis may have shown more degradation of hydrocarbons if given additional time. This hypothesis is bolstered by the trial 2 fourteen-day B. subtilis data, which shows a relatively steady concentration of bacteria until around day 8, when the concentration begins to increase in many samples. Compared with the trial 2 fourteen-day P. fluorescens data, which began steadily increasing in concentration from day one, B. subtilis took longer to grow appreciably.

Overall, even the P. fluorescens samples didn’t appear to grow as effectively as in preliminary work and the samples never became extremely cloudy. This change may be attributed the experimental set-up. Samples were grown in small Spec-20 tubes; these tubes have a decreased surface area, which in turn decreased the diffusion of oxygen into the ocean environment. While shaking the samples should have increased diffusion, oxygen may still have been a limiting factor in the samples. Also, the Spec-20 tubes had less space available for oxygen, further limiting diffusion. Additionally, the Spec-20 tubes increased the distance between the oil layer and the bottom of the sample. While shaking was intended to create a homogenous liquid layer, visible particulate matter accumulated at the bottom of the tubes. The height of the tubes may have reduced bacterial growth near the oil layer. However, in some B. subtilis samples, bacteria growth was visible on the bottom of the oil layer; this phenomenon suggests that the bacteria were able to metabolize the oil despite the distance. Finally, the addition of nutrients to the IO may have changed the pH of the solutions. If the pH changed substantially, the bacteria may not have been able to grow as effectively as in previous years.

Variations between the qualitative and quantitative analyses can be tentatively explained by a variety of factors. While the B. subtilis samples showed no qualitative degradation peaks, the HD appeared to have been degraded in three of the treatments. B. subtilis may have cleaved the hydrocarbons into chains smaller than 12-carbons, which would have appeared within the solvent peak on the gas chromatogram, rendering them indistinguishable. This phenomenon may have occurred in other samples as well, meaning that some degradation products were not visible in the gas chromatograms. To remedy this, the gas chromatograph should be run under a different program to make the peaks within the solvent peak more visible. Also, the nutrient combination never showed the greatest qualitative degradation, but quantitatively, it appeared to have the greatest HD degradation. This variation can be attributed to the phenomenon explained above: degradation peaks may have been hidden in the solvent peak. The fact that the .01 M sodium sulfate treatment showed the greatest quantitative degradation also supports the theories that the nutrients may have been present in too high a concentration or more complete degradation occurred, leaving even smaller carbon chains; the .1 M sodium sulfate treatment may have been too highly concentrated to be effective.

Unlike preliminary work, controls containing each bacterium and no nutrients showed some degradation, a testament to the powerful metabolism of the bacteria. The peaks in all of the B. subtilis samples were considerably smaller than the peaks in P. fluorescens samples. In both the B. subtilis and the P. fluorescens samples, the treatments containing phosphate caused the largest degradation peaks; however, these peaks were only slightly larger than the peaks from the other nutrient treatments. In all degraded samples, the peaks followed a predictable pattern: the 14-carbon peak (closest to the 16-carbon HD) was always largest, followed by the 13-carbon peak and the 12-carbon peak. This suggests that the bacterial metabolism involves a systematic cleavage of one or two hydrocarbons from the end of a longer chain. Additional analysis and trials will be necessary to explain variations in hydrocarbon data and deviations from previous experimentation, which suggested that sodium sulfate and P. fluorescens caused the most degradation.

Discrepancies in data may be accounted to procedural difficulties, namely settling in samples prior to absorbance measurements and bacterial contamination. A Spec-20 was used to measure the absorbance of each sample; using a calibrated absorbance curve, this measurement was converted to a bacterial concentration in samples containing bacteria. However, the Spec-20 is incredibly sensitive, to the point where it is difficult to obtain consistent data, as evidenced by the non-smooth curves in the bacterial concentration graphs. While steps were taken to make the absorbance measurements as uniform as possible, the data were still widely variable. Even quickly removing and reinserting a sample could cause a dramatic difference in absorbance readings, potentially due to air bubbles or the uneven distribution of bacteria. While the samples were inverted before an absorbance reading was taken, the bacteria still seemed to settle near the bottom.

While plating and Gram-staining on day 7 and day 14 suggested that no contamination occurred, growth and degradation in some

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autoclaved samples suggests that the autoclave did not properly sterilize or other bacteria entered the sample. This potential contamination may have occurred in other samples as well, leading to variations in data. On both day seven and day fourteen, .1-ml samples from the liquid layers of the two control flasks without HD were grown on a sterile agar plate. After two days of growth, the colonies from each plate were Gram-stained and viewed under oil immersion. The Gram-stain and shape of the observed bacteria matched the stain and shape of the bacteria observed before the trial. Also, the bacteria appeared significantly larger, suggesting that growth was occurring. Both of these observations suggest that contamination did not cause unexpected variations in data, so perhaps the increase in absorbance in autoclaved samples was not due to bacterial growth but to some reaction that increased turbidity.

An unwanted variable also occurred that may have affected the results. When sodium phosphate was added to the trial 1 samples, an unknown chemical reaction occurred, and the clear samples became opaque and cloudy. This is most likely a result of the “salting out” effect. Since sodium phosphate tribasic was used, for every mole of phosphate ions added, three moles of sodium ions were added. The samples all contained IO, which already had an abundance of salts. Therefore, salts many have precipitated out of solution, forming the white solid. It is unknown whether this reaction affected the bacterial metabolism. However, considering the application of the results, the unwanted variable does not have a large impact. Since the goal is to determine which nutrient should be added to ocean oil spills to increase bacterial metabolism of oil, whatever reaction occurred in the flask would most likely occur in the ocean as well, so any impact the reaction had would apply to the ocean spill as well. Therefore, while the reaction could be considered an unwanted variable, it did not change the implications of the results. To remove the variable, sodium phosphate monobasic was used in trial 2 and no precipitate formed.

The use of rubber stoppers in trial 1 may have caused additional variations in data. The HD appeared to react with the rubber stopper, causing the oil layers to turn varying shades of yellow. In almost all trial 1 samples, a wide peak appeared around 15 minutes, which is likely a result of contamination from the rubber. It is unknown whether this reaction between the HD and rubber affected bacterial metabolism, but considering that trial 1 showed close to no degradation while trial 2 showed significant degradation, the rubber stoppers likely had some effect on the bacteria.

If this experiment were to be repeated, a more accurate procedure should be used to calculate bacterial concentration, such as periodic serial dilutions of each sample. Also, aeration should be increased to further simulate a surface ocean spill and ensure that oxygen is not the limiting factor in the sample environments, which contain very little air space. The pH of the samples should be monitored and controlled, as changes in pH from the addition of the nutrients may have also limited bacterial growth.

Various extensions, such as varying bacteria strains used or varying nutrients, could also be derived from this initial experiment. While the results gained from trial one and trial two show many interesting details, more trials and further GC analysis are necessary to fully verify the initial conclusions made.

No known research has been completed on the effect of sodium nitrate, sodium phosphate, and sodium sulfate on P. fluorescens and B. subtilis, so comparison with previous results proves impossible. In the wider social context, however, these initial conclusions prove incredibly valuable. Marine oil spills affect society on a local, national, and international level. In all three regards, government agencies and environmental groups rely on scientific research to determine the most effective ways to remove oil from the environment. While a miracle nutrient treatment that causes P. fluorescens and B. subtilis to degrade oil at unprecedented speed would certainly be wonderful, results from this experiment suggest that such a nutrient does not exist. While each nutrient improved bacterial metabolism in some regard, no nutrient caused substantial change from the controls without nutrients. While not the original hope of this study, this realization proves equally important. Perhaps the most effective bioremediation protocol involves manipulation of the bacteria and their environment as opposed to the addition of nutrients. If additional studies suggest that this is indeed the case, oil spill cleanup could become more economically feasible and environmentally friendly. Not only would the cost of purchasing nutrients be saved, but also the additional environmental stress of adding large quantities of nutrients to the environment would be reduced. For local and national agencies alike, results from this experiment could help establish a more cost-effective strategy for dealing with both small and large-scale marine oil spills.

Although additional trials and analyses are necessary, it can be tentatively concluded that the hypothesis–if sodium nitrate is added to P. fluorescens and B. subtilis, then the amount of smaller hydrocarbons present in the samples will increase–is partially incorrect. While the treatment of .1 M sodium nitrate consistently caused one of the greatest increases in bacterial concentration with both P. fluorescens and B. subtilis, the data from these treatments differ only marginally from the positive controls and treatments involving other nutrients. Also, a qualitative analysis of the gas chromatograms from trial two suggested that significant degradation only occurred with P. fluorescens samples. In these samples, .1 M and .01 M treatments of sodium phosphate caused the largest degradation peaks. Again, the peaks from these samples differ only slightly from the positive controls and other nutrient treatments. However, quantitative analysis of the amount of HD degraded showed that .1 M nutrient combination and .01 M sodium sulfate treatments caused the greatest HD disappearance. Overall, no nutrient treatment caused substantial improvement in comparison to controls in any assessment of bioremediation effectiveness, suggesting that perhaps the addition of nutrients is not necessary to stimulate hydrocarbon degradation. After being validated by additional trials, the results from experimentation can be applied to the bioremediation of future ocean oil spills by providing a more effective method for breaking the hydrocarbon chains into smaller compounds. If this topic were explored further, the potential exists for the identification and isolation of the specific enzymes, and later genes, involved in hydrocarbon metabolism. Future genetic engineering could create specialized bacteria designed for more effective hydrocarbon metabolism. The Deepwater Horizon catastrophe reinforced the need for a cost-effective and ecologically safe means of removing oil from ocean environments, and data gained from this experiment represent the first step towards enhancing bioremediation of ocean oil spills.

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References1. Environmentally Conscious Fossil Energy Production, John Wiley & Sons. 180. Mellor E, Elkamel A. (2010).

2. Mellor E, Landin P, O’Donovan C, Connor D. (1996). The Microbiology of In Situ Bioremediation. Groundwater Pollution Primer. 3. The Biological Metabolism of Nitrate and Nitrite in Pseudmonas fluorescens K27 Amended with Tellurium. Sam Houston State University. Tian W. (2004).

4. Hoffmann T, Frankenberg N, Marino M, Jahn D. (1998) Ammonification in Bacillus subtilis Utilizing Dissimilatory Nitrite Reductase Is Dependent on resDE. J. Bacteriol. 180.1.

5. Chirnside AEM, Ritter WF, Radosevich M. (2007). Isolation of a Selected Microbial Consortium from a Pesticide-contaminated Mix-load Site Soil Capable of Degrading the Herbicides Atrazine and Alachlor. Soil Biology & Biochemistry 39.12.

AcknowledgementsWithout the invaluable knowledge and resources of many, this study could not have been completed. Special thanks to Dr. Anastasia Chirnside and Dr. Caroline Golt of the University of Delaware for providing access to their laboratories and equipment as well as their expertise. Additionally, thanks to Sandra Litvin and the entire Unionville High School science department for providing laboratory space, supplies, and monetary support. William Anderson also provided invaluable editing. Finally, a special thanks to the Delaware Valley Science Fair for the innumerable opportunities they have provided over the years and their support of middle and high school science research.

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