Sarah Bonazza and Nicole MacDonald

Abstract:

            This project investigates the degradation rates (as well as other chemical and physical changes) of oil spills in seawater. One experiment is controlled with nutrients and the other left to natural degradation. The findings could potentially give insight on the problems with vegetable oil spills in our environment. The ideal solution to this type of research would be teaching the public of the unknown dangers of vegetable oil.

            A lot of today’s focus in biotechnology is on the environment and our global surroundings. There are many toxins in our world that may harm or pollute humans, animals, plants or the air. Vegetable oil spills are becoming more common and are potentially more dangerous than hydrocarbon spills. One spill in any place can pollute all other aspects of the ecosystem, especially because it is not easily visible.  It can spread fast between bodies of water and the land, coming in contact with humans, plants and animals. Our results will simulate the effect on the economy as well, as clean up of spills with bioremediation may cost a lot of money, but could potentially create more jobs.

 Objective:

             To understand the persistence of canola oil in the environment, and to study its short term and long term effects, as well as the oil’s chemical and physical changes. Will natural or biostimulated degradation be better for the environment?

Introduction:

            Vegetable oil is oil that has been removed from a plant or the seeds of a plant by pressing the seeds and extracting the oil with steam or water. Canola oil represents 82% of domestic vegetable oil production in Canada (other vegetable oils include soybean: 15%, flaxseed: 2%, and sunflower: 1%). In 1998, Canada produced 1.4 million tonnes of canola oil with a market value of $1.8 billion (total global production of vegetable oils was 33 million tonnes). Canola oil is a semi-drying oil. It is used as a lubricant, fuel, soap, cooking oil, and a synthetic rubber base. It comes from rapeseed, a poisonous weed in the mustard family that even insects will not eat, and is very toxic. It has a high level of monounsaturated fatty acids. Another less common name for canola oil is rapeseed oil. Vegetable oil shipments (mainly canola) through the Port of Vancouver have tripled over 10 years. Due to the increase, there have been more oil spills in the Vancouver Harbour (three). A 20 ton canola oil spill occurred in November, 1999, killing up to 2,000 sea birds. The frequency of canola oil spills is expected to increase as production, use, and transportation does.

Vegetable oils are harmful to the environment; like petroleum oils they produce similar environmental effects. In the public eye, canola oil is seen as safe because it is “edible.” Since it is almost clear in color, animals do not often see the oil and wander unknowingly into a spill. If spilled in land, vegetable oils are almost undetectable. They cause devastating physical effects, such as coating bird feathers, gills of aquatic animals, plants, sediments, and other surfaces with oil and suffocating them by oxygen depletion (the oils cause depletion of oxygen in the water column to a level below what aquatic life needs). They destroy future and existing food supplies, breeding animals, and habitats, and produce rancid odors. They create foul shorelines, clog water treatment plants, catch fire when ignition sources are present, and form products that linger in the waters and environment for many years. Canola oil (and other vegetable oils) can be toxic and form toxic products. The degree to which something is poisonous is called toxicity. Industry has claimed that as a non-petroleum and “edible” oil, canola oil is not harmful to the environment, but some of its other effects on wildlife include hypothermia and loss of buoyancy.

Canola oil is a persistent environmental pollutant, and its chemical composition and toxicity may change while it lingers in the ecosystem, possibly becoming more dangerous. Natural biodegradation does occur, and emulsification in water, but optimizing the conditions of the oil spill by regulating temperatures and adding nutrients can speed up the break down.

One component of rapeseed oil is erucic acid, found to impair responses in mammalian test populations. Other components have altered the size and metabolic activity of freshwater teleosts. When released into the environment, the toxicity of vegetable oil is altered by chemical and biological processes such as evaporation and oxidation.

 Materials and Method:

 Part I: Sampling

Materials: 3 carboys, seawater, canola oil, Marine Broth nutrients, fertilizer, air line and pump, fume hood

 Figure 1.  Schematic of experimental setup.

 Method

            Three glass carboys were filled with 8 L of seawater (Fig. 1) that had been filtered to remove any zooplankton or other creatures, but which retained some of the bacterial community. The first carboy contained only seawater and served as the experimental control.  The second carboy (natural degradation) contained 80 mL Canola oil on top of the seawater. The third (bioremediation) contained both Canola and 0.75 g Marine Broth (Difco), which are nutrients that enhance the growth of marine bacteria.  All three carboys were maintained at room temperature in a dimly-lit fume hood and each aerated by one pump.

            For sampling, a glass pipette was used to take 120 mL of the water from each carboy: 100 mL for GC-MS analysis to determine changes in the concentration of oil, 10 mL for the microtox to determine the toxicity of the aqueous phase to the marine bioluminescent bacterium (Vibrio fischeri), and 10ml to measure the pH level. Other parameters measured were temperature, salinity, and light intensity (light - only for the initial sampling date).        

Table 1. Schedule for taking samples from each of the three carboys.

  Part II: GC/MS Analysis

Equipment: 250 mL separatory funnels, 100 mL graduated cylinders, disposable glass pipettes, 200 mL Zymark concentration tubes, 15 mL graduated test tubes, TurboVap II, sonicator, heater, fume hood, vortex mixer, GC autosampler vials and caps, HP GC/MS.

 Reagents: dichloromethane (DCM), hexane, glass distilled water, methanol (MeOH), potassium hydroxide (KOH), BSTFA+TMCS 99:1(derivatizing agent), conc. hydrochloric acid (HCl).

 Extraction Step

            The entire contents of samples were transferred from the sample bottles to 250 mL separatory funnels, the sample bottles rinsed with 100 mL DCM and the rinse added to the funnel. 

            The separatory funnels were shaken vigorously by hand for 30 seconds and then placed on a rack to allow separation. The DCM layer was collected into a Zymark tube and the water was layered into a 100 mL graduated cylinder. The volume of seawater was recorded and returned to the sample bottles and stored in the freezer. DCM extracts were evaporated down with nitrogen to 1.0 mL on the TurboVap II. Samples were quantitatively transferred to pre-weighed 15 mL graduated test tubes with 3 x 1.0 mL rinses of hexane. Hexane was evaporated using the Nitrogen evaporator and the test tubes were re-weighed to determine the sample weight.

 Saponification Step

            The extracts were quantitatively transferred to graduated cylinders with 3 x 1.0 mL DCM. 30 mL MEOH/KOH solution was added to the graduated cylinders and the samples shaken, placed in a sonicator, covered with tin foil, and sonicated for 1.5 hours at ~70oC. 

            Samples were then cooled, brought to 60 mL with glass distilled water, and shaken. They were then acidified with 8 mL concentrated HCl and mixed. The samples were then extracted three times with 10 mL hexane, the top hexane layer drawn off with a glass pipette and dispensed into a Zymark tube. They were then evaporated down to 1.0 mL in TurboVap II and quantitatively transferred with 3 x 1.0 mL hexane into 15 mL graduated test tubes. The Nitrogen evaporator was then used to bring them down to 1 mL and they were diluted as necessary. 25 mL of derivatizing reagent, BSTFA+TMCS 99:1 was added to the diluted samples and they were swirled gently. The samples were then transferred to autosampler vials and sealed with crimp caps.

 Analysis with GC/MS

Run on HP GC/MS.

 Part III: Microtox Analysis

Chemicals: filtered seawater, Microtox Acute Reagent (freeze-dried bacteria, V. fischeri), Microtox Reconstitution Solution

 Materials: glass cuvettes, Microtox model M500 Analyzer, pipettes and tips (Eppendorf), Refractometer (Aquafauna)

 Method

            About 100 µL of the sample was placed on a refractometer to measure salinity. The salinity of the seawater used as diluent (dilution water- sea water that was filtered using  0.22 µm filters, funnel, clamp and vacuum flask, and vacuum pump). The vacuum was set to –34 kpa (10 inches mercury) so that phytoplankton and bacteria would be left on the filter and removed from the water. The seawater was 32 ppt salinity.

            Each sample was run in duplicate. Glass cuvettes were placed into incubator wells A1 to A5 of the Microtox M500 Analyzer for replicate test number 1, and wells B1 to B5 for replicate 2. The machine keeps these at 15ºC. At the same time, a cuvette was placed into the reagent well of the M500 Analyzer which maintains the reagent temperature at 5ºC. Using an Eppendorf pipette with a clean diposable tip, 1 mL of Microtox Reconstitution Solution was added to the cuvette in the reagent well. With an Eppendorf pipette with a clean diposable tip, 1 mL of diluent was added to the glass cuvettes in the incubator wells A1 to A4 (replicate 1) and B1 to B4 (replicate 2). The Eppendorf pipette (with another clean tip) was used to transfer 1 mL of sample to the glass cuvettes in wells A4, A5, B4 and B5.

            A series of 4 dilutions and 1 diluent control for each replicate test were made by using the pipette to sequentially mix and transfer 1 mL of sample from cuvette A4 to A3, and mix and transfer 1 mL from A3 to A2.  1 mL is removed from the mixed A2 cuvette and discarded. The same mixing procedure was carried out for the second replicate (cuvettes B4, B3 and B2).  This provides test concentrations of 0, 12.4, 24.8, 49.5 and 99% (not 100%, because of the 10 µl of acute reagent that will be added to each cuvette).  If the EC25 was lower than 12.4%, then a concentration series was prepared so that the EC25 would fall within the tested concentrations.

            A vial of Microtox Acute Reagent (freeze-dried V. fischeri bioluminescent bacteria) was reconstituted by taking the vial from the freezer and pouring into it the chilled (5 C) Microtox Reconstitution Solution from the cuvette in the reagent well. The vial was swirled 4-5 times and the reconstituted bacterial solution (reagent) was poured back into the glass cuvette. The cuvette was set back into the reagent well of the Analyzer.  This reagent was used in tests for up to 2 hours, after which a fresh vial of reagent was reconstituted for use. After 5 minutes, each cuvette of the test series was inoculated with 10 µL of the bacterial reagent, and the cuvette shaken to mix its contents.

            The software program on the computer was set up to receive and record light intensity data from the M500 Analyzer. After 15 minutes of incubation, photoluminescence of each cuvette was measured on the M500 Analyzer. Data was analyzed and EC25 values reported using the MicrotoxOmni software.

 Data:

 Part II:

SC

 SCN

 Part III:

 Ratios of toxicity from days 28 to day 0

SC – 10.963 : 1.000 (less toxic then SCN)

SCN – 22.820 : 1.000 (more toxic than SC)

 Discussion

 Part I:

The results of the temperature data taken from each sample, was that the temperature stayed relatively constant through out the whole process. Though on Day 14, it did drop drastically due to changes in the laboratory area. The temperature of SCN container was always greater than the temperature of the S and SC containers, which shows that some activity was going on. The temperatures for Day 0 were not used because of the fact that they had just been taken out of the ocean.

            The results of the Salinity data showed that for every container the salinities were approximately the same. The salinities of the S container did change more than the others, but was irrelevant due to fact that it was just sea water.

	The results of the pH level data showed a constant level in the S and SC containers. The levels of the 

SCN containers dropped over time which can be a clue as to what was happening chemically, and also what may 

have caused the toxicity to V. fischeri in the Microtox test.

 Part II:

	The results from the GC/MS testing of our samples, both SC and SCN, unfortunately did not show any 

evidence of either natural oil degradation or bioremediation of the oil with nutrients. When looking at the graphs, 

the patterns for both of the SC ions are similar in shape, but there is no apparent trend throughout the 28 days in the

abundance of oil. The two ions for SCN also showed similar patterns, but no trend could be detected. This could be 

for many reasons, probably because the experiment was not done over a long enough period of time. Usually 

experiments of this type are done over several months or even a year or more. Perhaps the ratio of oil to water 

was too high, or there was not enough aeration in the carboys. Poor sampling technique could have been a problem 

as well because excess surface oil was collected along with the sample water. Also, if the environment had been 

warmer for the water, perhaps the nutrients would have biodegraded the oil at a greater rate or the original 

enzymes in the water could have broken down the oil naturally.

 Part III:

	Our results from the toxicity testing have shown that in both the SC and SCN carboys, the oil became 

more toxic in the water. In terms of the Microtox, this means that less light could be produced by the V. fischeri in 

the samples at day 28 than at day 0. This is very dangerous to the environment and all ecosystems because all living 

things are at one point affected by seawater, and the toxic nature of this water where oil was left can harm the 

cycle of life. Since the ratio of the toxicity from day 0 to day 28 was greater for SCN than SC, the toxicity was 

greater overall for SCN. Even though biodegradation was not shown with the GC-MS, it was probably to small to 

be detected. The breakdown products of the biodegradation could have caused the increased toxicity of the SCN.

	The 100% Microtox Acute Test was used (in duplicate so that an average and standard deviation could 

be calculated) to determine the concentration of sample that causes an effect of 25% reduction in the light output of 

the bioluminescent marine bacteria, Vibrio fischeri. This 25% light loss value is called the EC25. The lower the 

EC25, the more the light output is being reduced, so the more toxic is the sample. For this particular test method, a 

sample that is not toxic will not cause any significant loss of light (compared to a control having the V. fischeri in 

seawater only), so the EC25 value cannot be calculated. This is indicated as EC25 > 99%, meaning that the 

effective concentration would be greater than the highest testable sample concentration.

 Conclusions:

               From this experiment, it has been determined that oil cannot be degraded in seawater in only 28 days, 

naturally or when biostimulated with nutrients. That is not to say that it will never degrade, but if conditions were 

more favourable and perhaps the set-up of the experiment was modified, partial or complete degradation of the 

canola oil could be possible. With this experiment, the oil became more toxic to its environment (seawater) over 

time. The light output by the Vibrio fischeri was less at day 28 than it had been at day 0, for both SC (10 times 

more toxic) and SCN (22 time more toxic). Therefore, in terms of toxicity, if this process were to be repeated in 

the environment, the oil would be better off left to degrade naturally than to have nutrients added to it. The actual 

experiment and testing would have to be performed again over a longer period of time and under different 

conditions to determine which type of degradation would have a greater rate. This could then in turn change the 

toxicity to be favourable to the environment.

THANK YOU!