Tomato induced DDT disappearance Research Paper 2011-2012 A Continuation of 2010-2011 Research Paper: Phytoremediation: to mutate or not to mutate? And A Continuation of 2009-2010 Research Paper: The effectiveness of the phytoremediation of dicofol using Lycopersiocon esculentum

William John OBrochta Research Instructors: Mrs. Cindy Bohland, Dr. John Kowalski, and Mrs. Sherry Otruba Roanoke Valley Governors School for Science and Technology

Abstract This project was designed to determine whether brt mutated tomato plants phytoremediate more than wild-type plants and if phytoremediation has any detrimental health effects on both types of plants. The hypothesis was that plants that have been genetically mutated to increase root length and size will phytoremediate more effectively, with greater negative health effects, when 1.50 grams (g) of Kelthane is applied than wild-type tomato plants. Phytoremediation ability was measured using a mustard bioassay. Plant health was determined by measuring chlorophyll concentration, leaf area, plant height, Brix concentration, plant dry mass, and root wet mass analysis. Results showed that the hypothesis was not supported as the bioassay showed that autoclaved soil alone removed 0.384 g of Kelthane, while the mutated plants removed 0.537 g, and the wild-type removed 1.140 g out of the 1.50 g added. The wildtype plants removed significantly more Kelthane than the mutated plants. The health of the mutated plants was better overall. Mutant plants had a significantly greater increase in leaf area, 123% for those with Kelthane, when compared to a -5.16% for wild-type plants undergoing phytoremediation. Plants that were not phytoremediating increased leaf area at a steadier 41% to 61% rate. Percent change in plant height showed that mutant plants grew taller without Kelthane (275%, 166%), while wild-type plants were significantly taller when phytoremediating (279%, 234%). The characteristics of the mutation show that high sucrose levels in the mutation decrease phytoremediation. Thus, decreasing sucrose levels by increasing acid invertase levels should increase phytoremediation. Introduction Plants have long been used to save our environment (Singh, et. al., 2007). Large dumps

of pesticides, heavy metals, and toxic wastes in recent years means that our need for plants has only increased (Arms, 2004; Singh, et. al., 2007). Frequently, large deposits of lead or mercury will go untreated, leaching into wastewater or preventing vegetation growth (Arms, 2004). Fortunately, individuals and governments have become more aware of their own environmental problems, but they have resorted to inaction because of the time, expense, and expertise required to mitigate or eliminate pesticide hazards (Burtt, 2009; Arms, 2004; Singh, et. al., 2007). Governments have become increasingly opposed to trying new methods of site cleanup, instead resorting to digging up all of the soil and trucking it away (Burtt, 2009). Four current methods are used to solve soil contamination issues: landfills, incineration, bioremediation, and phytoremediation. Use of landfills to transfer contaminated soil only prolongs an already bad problem (Gardea-Torresdey, 2003). Landfills combine many hazardous pesticides together to create a high concentration of dangerous chemicals and leach into groundwater, causing further contamination. Even if the landfills are lined or sequestered, it is not difficult for a cocktail of pesticides to decompose the lining. Incineration emits harmful ash that if inhaled can lead to breathing problems, making the method worse than using a landfill (Gardea-Torresdey, 2003). Phytoremediation is the new potential solution for this 1.7 trillion dollar problem (Gardea-Torresdey, 2003). Various types of plants are placed on soil that contains either chemical pesticides or heavy metals. Roots cause an increase in the number of pesticide digesting microbes by as much as 10,000 fold (Evans, 2002). Therefore, the addition of the roots allows for pesticide degradation, meaning that the amount of chemical is reduced (Evans, 2002). The reduction can be drastic, as much as 75 percent in two to three years, compared to 45 percent using bioremediation (the use of soil microbes alone to digest the pesticide) (Evans, 2002).

Aerobic bacteria bioremediation has been known to degrade many pesticides and metals and can oxidize some petroleum products (Thieman and Palladino, 2009). Phytoremediation has been claimed to be 30 percent more effective when compared to bioremediation (Greenberg, 2006). In a study on contaminated soil sites, Crane (2009) notes that phytoremediation removes between 33 and 46 percent of an oily contaminant, confirming conclusions that phytoremediation is definitely an effective clean-up method (Crane, 2009). The phytoremediation used in these experiments involved rhizodegradation, enhanced phytoremediation abilities in plant roots, and phytoextraction, chemical accumulates in the leaves of plants (Russell, 2005). Rhizodegradation involves increases in the amount of bacteria present in the rhizosphere area of the root (near the top) (Zobel, et. al., 2005). This type of phytoremediation is most common; however, some rhizosphere bacteria can harm the plant and environment, due to phytotoxicity or the maximum level of a certain pesticide that a plant can keep before its health is affected (Zobel, et. al., 2005; Russell, 2005; Weaver, 2010; Rose, 2010). Effects of rhizodegradation can include increased nutrient uptake and increased water uptake, both important in phytoremediation ability (Zobel, et. al., 2005). Phytoextraction works when phytoremediated compounds are too heavy to be released and are slowly degraded in the plant (Gerhardt, et. al., 2009). Plants that phytoextract can be removed and incinerated or left in the soil (Gerhardt, et. al., 2009). This means that incineration still may occur, but the plant matter is much less concentrated and smaller in mass than incinerating the soil. Previous research suggests that leaving the plants in the soil after phytoremediation has occurred will not affect plant health in any significant manner (OBrochta, 2011). Probably the greatest downfall for phytoremediation is not the effectiveness, but the

expense, time, and compatible plants and chemicals. In short, this method works with a few types of plants on a few chemicals and metals over a very long period of time. A typical phytoremediation application can cost up to $694,000 (Russell, 2005). The Environmental Protection Agency notes that the amount of time for phytoremediation to occur depends greatly on the type of plants and amount of dangerous pesticide present (U.S. EPA, 2001). Singh (et. al., 2007) goes so far as to claim that phytoremediation is actually a less costly alternative to traditional cleanup methods. McGrath, Dunham, and Correll provide a useful cost/benefit analysis of phytoremediation technology (Terry and Bauelos, 2000). Landfill disposal costs about $117,000 per hectare, while disposal of incinerated ash from phytoremediated plants amounts to only $70 per hectare (Terry and Bauelos, 2000). Potential spill chemicals and toxins that may be removed by phytoremediation can be broken into two groups, heavy metals and chemical compounds (Cutraro and Goldstein, 2005). Polycyclic aromatic hydrocarbons (PAH), polychlorinated biphenyls (PCB), and even dichlorodiphenyltrichloroethane (DDT) can be removed to some degree with phytoremediation (Eckley, 2001). All places have some kind of PAH contamination caused by the degradation of organic compounds in the soil (Cutraro and Goldstein, 2005). Thus PAH and Persistent Organic Pollutants (POP), chemicals defined by the EPA as having the longest half-life, are in the process of being eliminated from exported pesticides; however, removal of these chemicals from the soil will be a problem for years to come (Smith, et al., 2008). Plant type becomes the second biggest limitation of phytoremediation after chemical type. The ideal plant for the type of contamination should be selected, though no list of effective plants exists (Cutraro and Goldstein, 2005). Singh (et. al., 2007) again claims that phytoremediation is superior in this regard because the plants

evolve a tolerance level for the pesticides that are in the ground near them. Phytoremediation has produced successful results in grasses (especially fescue), legumes, aquatic plants, and metal hyperaccumulators such as alpine pennycress (Gardea-Torresdey, 2003; Zobel, et. al., 2005). A metal hyperaccumulator stores the metal in the leaves of the plant, a feat few plants can perform (Cutraro and Goldstein, 2005). The first application of phytoremediation used Saint Augustine grass and got effective results (Evans, 2002). This was probably more luck than proper plant choice. The Ford Motor Company is trying the best method available at this time to remediate former auto manufacturing plants where the soil is contaminated with oil, planting many species of plant to test which work best in their affected area (Evans, 2002). Researchers began with 55 plant species, narrowed down to 22 (Evans, 2002). Each was tested on a portion of the contaminated land and results were compared, producing the best plant for the site (Evans, 2002). This method is time consuming and inefficient, discouraging the use of phytoremediation. Time and money are also considerations when choosing to use phytoremediation and can be presented as drawbacks. An oil spill cleaned using Saint Augustine grass reduced 75 percent of the pollutants in two years (Evans, 2002). Phytoremediation does not work on a schedule, and repeated trials never take the same amount of time (Evans, 2002). The Ford project mentioned above is being implemented; however, it might have to be supplemented with old incineration or landfill techniques because the phytoremediation is taking longer than their four-year deadline (Evans, 2002). Though the cost of phytoremediation is decreasing, it is still much more expensive than conventional methods (Cutraro and Goldstein, 2005). The phytoremediation market now tops 214 million dollars per year (Evans, 2002). Even with these many problems, phytoremediation is expected to solve the environmental pollution problem (Wiley, 2007).

The ten-year-old phytoremediation phenomenon has been the subject of some small-scale research, though no real consensus exists regarding appropriate plants or which chemicals might be best suited for phytoremediation (Singh, et. al., 2007). Interest lay, therefore, in determining if mutations to tomato plants help phytoremediate land contaminated with pollutants. Additionally, little research has been done to indicate what happens to plants during phytoremediation. The projects purpose was to determine if detrimental effects occur to a plant that attempts to phytoremediate a chemical, in this case a pesticide, and whether biotechnologically mutated plants improved phytoremediation capability. The above objective is the same as a previous research project, except the goal has changed to testing mutated plants that exhibit characteristics especially helpful to phytoremediation. This project has a practical application within the realm of phytoremediation. Specific mutations can be identified that improve phytoremediation abilities. These mutations, like the one tested in this experiment, will allow phytoremediation application with fewer plants and greater effectiveness, making the technology much more attractive to businesses. Biotechnological approaches to phytoremediation have, thus far, been the source of little research. There are two goals associated with genetic modification of plants for phytoremediation increasing ability and lowering cost (Singh, et. al., 2007). For increasing phytoremediation of metals, the key is to increase the number of water and nutrient uptake sites on the roots and raise the quantity of metal transporters in the xylem (Singh, et. al., 2007). Tomato plants, known hyperaccumulators of Cadmium, need to gain biomass in order to be effective phytoremediators (Setia, et. al., 2007; Cherian and Oliveria, 2005). Transferring genes or traits from bacteria or animal systems frequently improves remediation potential (Cherian and

Oliveria, 2005). This was found to be true in genetic engineering selenium phytoremediators (Terry and Bauelos, 2000). Terry proposed engineering the Indian mustard plant to overproduce enzymes and introduce additional metabolic pathways to remediate the selenium (Terry and Bauelos, 2000). Tomato plants and dicofol miticide (Kelthane) were used to complete this phytoremediation test. Tomato plants are not known for their phytoremediation abilities (Bush, n.d.). Research showed that mutated tomato plants may phytoremediate more effectively than regular tomato plants (Bush, n.d.). This may be due to modified root structure and veins. A bushy root variety of tomato plant was selected from the University of California Davis Charles M. Rick Tomato Genetics Center for this experiment under the rationale that plants with larger roots could take up more chemical (Chetelat, 2010). Zobel (1971) located this mutation and notes that The root system is very highly branchedthe root system branches profusely within one day after emergence, in contrast to normal roots, which branch only after several days of growth (Zobel, 1971). Zobel also notes that brt mutated tomato plants germinate more slowly than non-mutated plants (Voland and Zobel, 1988). This mutant also displays increased colonization of fungus on its roots (Zsogon, et. al., 2008). Increased fungus presence could contribute to phytoremediation abilities because of the plants growing need for nutrients (Zsogon, et. al., 2008). There may be more microbial enzymes in the roots (Benedito, 2010). Overexpression of root membrane proteins in Indian mustard plants led to an increase in phytoremediation ability for removal of selenium (Terry and Bauelos, 2000). Peres (2010) notes that he has observed an increased concentration of Brix (sucrose) on the roots. Zobel (2010) confirms this observation by stating that there is an increase in starch at the base of the roots that

could be duplicated by the presence of sucrose. This sucrose is likely located on the microbial chelators, which are known to deliver nutrients to the plant, while sucrose probably is located on the top of the rizosphere (root shoot) (Gerhardt, et. al., 2009). Levels of Auxin and Gibberellin (plant growth hormones) increased in the brt mutant, when compared to non-mutated plants (Sidorova, et. al., 2002). These results were observed in pea plants with the same mutants, so the results should be similar for tomato plants (Sidorova, et. al., 2002). However, the same researcher showed that Auxin levels were actually decreased when compared to the control in a later experiment (Sidorova, et. al., 2010). Auxin hormones control stem shoot branching (Shimizu-Sato, et. al., 2008). It is reasonable to infer that the hormone levels also control root shoot branching, supporting Sidorovas (et. al., 2002) link between Auxin levels and bushy root plants (Shimizu-Sato, et. al., 2008). In a 2010-2011 research project, the experimenter found the location for the bushy root mutant on the twelfth tomato chromosome at 19.8 cM (unit length of chromosome) or 95.8 cM. The gene at this location was TG296, a Lysr transcriptional regulator protein from bacteria that was placed in the castor bean plant before being extracted by Zobel at U.C. Davis (Zobel, 1971; Voland and Zobel, 1988; OBrochta, 2011). Kelthane 50W (or WSP) Agricultural Miticide has been manufactured by Dow AgroSciences Canada Inc., Rohm and Haas Company, and Makhteshim-Agan and is a miticide that provides a high initial kill and good residual (long lasting effectiveness) (MSDS: Kelthane, 2008; Rossi, 1998). A white to gray powder, it has an odor of fresh cut hay (MSDS: Kelthane, 2008). Kelthane is composed of about 51 percent dicofol (Kelthane, 2005). Dicofol is a nonsystematic acaricide (poisonous to mites) used to control mites that damage cotton, fruit trees, and vegetables (Qiu, et al., 2005). There are few adequate alternatives to dicofol because

it is cheap and effective, however, as a result of the Stockholm Convention, it is being banned for residential use, phased out for agricultural and commercial use, and highly restricted for experimentation (Snchez, et. al., 2010). Dicofol is similar in composition to DDT (Figure 1) and, therefore, is classified a Persistent Organic Pesticide (Eckley, 2001). DDT is actually an intermediate substance in the forming of dicofol (Snchez, 2010). These two pesticides are often used interchangeably and results in a dicofol experiment should apply to DDT (Garber and Peck, 2009). The EPA notes several important distinctions between DDT and dicofol, chiefly that dicofol is more water-soluble that DDT (Rossi, 1998). Essentially, all results found for dicofol are worse for DDT and is considered less harmful than DDT (Rossi, 1998). DDT has caused huge environmental problems and was the basis for the popular Silent Spring by Rachael Carson (Eckley, 2001). It has also been linked to causing over fifty percent of breast cancer cases in women when it was in use (Watts, 2008). DDT is known to cause pancreatic cancer and neurological problems, though it is still too early to determine the exact effects, as many of DDTs problems are birth defects (Snchez, et. al., 2010). Dicofol is also extremely present in soil after long periods of treatment, with a half-life of 20-30 years (Gao, et. al., 2000). However, after only a short period of exposure to dicofol, initial degradation is somewhat exponential (Garber and Peck, 2009). This is not uncommon, though significant pesticide initially degrades; the rate of degradation slows after little additional time, but still meets or exceeds legal regulations in Italy (Cabras, et. al., 1985). Still, dicofol remains a huge problem because of its toxicity to many fish, causing mutations and decreased survival (Garber and Peck, 2009). DDT also bioaccumulates, or builds up. As predators eat prey, the concentration of DDT increases significantly (Withgott and Brennan, 2008).

Phytoremediating dicofol and DDT has been studied on a limited basis and a procedure for the remediation has been developed (Thompson, 2010; Gao, et. al., 2000). The DDT begins to be remediated when it is taken from the soil through the roots of the plant (Gao, et. al., 2000). This uptake is limited by the fact that both DDT and dicofol are hydrophobic and they resist water travel (Gao, et. al., 2000). A concentration gradient is formed near the root epidermis that is semi-permeable and absorbs some of the pesticide, transporting it to the root xylem using transport proteins (Setia, et. al., 2008). Benedito (2010, 2011) suggests that there are likely increased transport proteins in the roots of the bushy root mutated tomato plants. This suggestion is confirmed through previous research that points to a transcriptional protein gene modification that would effectively produce more transport proteins to increase the amount of DDT that could be transported from the root epidermis into the xylem. Plant metabolism transforms the DDT and degrades it significantly, first into DDD, a less hazardous pesticide, and then catalyzes the DDD using naturally occurring reagents (Gao, et. al., 2000). DDT can also form DDE through a dehalogenation, removing both halogen and hydrogen from the DDT (Gao, et. al., 2000). However, the remediation procedure in tomato plants could be significantly different than the one described since it occurred in two types of grasses (Gao, et. al., 2000). Frequently, remediated pesticides or metals will be sequestered in the leaf or stem (Setia, et. al., 2008). Either a vacuole will form around the pesticide or it will be sequestered away from any vital cell or plant process (Setia, et. al., 2008). Similar experiments have been conducted using different plants and different chemicals from this experimenter and others. A phytoremediation experiment in 2005 using rye grass to remove DDT was extremely effective (Greenberg, 2006). In fact, 30% of the DDT was removed

within 90 days, but it is noted that there is know way to know whether DDT is being degraded in the soil or in the plants, an important consideration (Greenberg, 2006). Initially, phytoremediation of DDT was deemed impossible, but was proven possible in 1977 (Russell, 2005). Chu (2006) performed a hydroponic experiment using DDT, PCBs and remediated both with rye grass (Chu, et. al., 2006). Though this test used an extremely small (ng) sample of DDT, it was remediated at a fairly fast rate and the half life determined to be only two or three days for such a small amount of DDT added (Chu, et. al., 2006). Recall, however, that this was a hydroponic test, so the DDT could have degraded in the water and not as a result of the plants (Chu, et. al., 2006). Interestingly, the DDT was mostly decomposed into DDD or DDE and the vast majority remained in the roots of the grass plants with some in the plant stems and minute amounts in the leaves (Chu, et. al., 2006). Many of the researchers and professors that the experimenter spoke to are also working on phytoremediation and genetic mutation analysis. The experimenter also performed previous research on this topic, using regular tomato plants to perform a similar test (OBrochta, 2011). This experiment involved growing mutated tomato plants and applying dicofol twice to see how much phytoremediation occurred and what the effects of the phytoremediation were on the plants. The added amounts of dicofol represented a situation where dicofol was applied yearly at a standard application rate. Tomato plants were then planted. Then a spill of dicofol was simulated with the tomato plants already in place. The independent variables in the experiment were the application of dicofol on the plants and soil and the type of plant used. Dependent variables were how much phytoremediation occurs in the plants, and the effect of this phytoremediation on the growth of the plant. Leaf area and chlorophyll content were analyzed

post-experiment to determine if there was a significant difference between average initial growth of the plants and average final growth. Plant mass, root mass, and root sucrose content was also determined. The amount of phytoremediation that occurred was measured using the bioassay method that bases germination of known amounts of pesticide against unknown amounts of pesticide. This method was determined to be effective enough for fairly precise estimations (Orcutt, 2010). The hypothesis for this experiment focused on the ability of the mutated tomato plants to phytoremediate: Tomato plants that have been genetically mutated to increase root length and size will phytoremediate more effectively, with fewer negative health effects when 1.5 g of dicofol is applied than wild-type tomato plants that have not been mutated. Materials and Methods The experiment was set-up with eight plastic plant trays, one and part of another for each variable tested. Two sunlight bulbs (40 watts, 122 cm tube) was installed in each of two fluorescent light fixtures attached to a long metal pole. The pole was taped to six Quick-Grip clamps and attached to the ends of two inch pieces of plywood (8 feet total) lined with blue plastic and elevated using sawhorses. A timer controlled the duration of light for the hours of seven in the morning to eleven at night. Temperature was controlled between 21.1 and 26.6 degrees Celsius and it was monitored using a digital thermometer. The setup was placed in an upstairs room next to a set of windows eight feet long. Sixty 5 inch diameter biodegradable (Jiffy Pots) plant pots were used in this experiment. They were purchased with two 5/16 inch holes for drainage. These holes were covered with a piece of duct tape to prevent pesticide leakage and evaporation of the pesticide. There were ten samples in each of six test groups and controls. Group A contained neither dicofol nor plants. Group B contained wild-type tomato plants without dicofol. Group C contained bushy root

mutant plants without dicofol. Group D contained dicofol, but no plants. Group E contained dicofol and wild-type tomato plants. Finally, Group F contained dicofol and bushy root mutant tomato plants. Scotts Premium Topsoil that contained organic materials and peat moss was used in the experiment. The soil was covered in foil and autoclaved at between 10 and 15 psi using one of two automated autoclaves for 30 minutes. Soil was placed in 1000 ml Pyrex beakers and autoclaved three or four at a time. Autoclaved soil was placed in sterilized plastic bins as quickly as possible and covered with aluminum foil. Using an alcohol sterilized plastic container and under a fume hood, half of the required soil (15 pots worth, 170 g per pot) was mixed with 7.5 g of dicofol. This was mixed for four minutes by hand wearing gloves and goggles. The procedure was repeated for fifteen additional pots. All soil was placed in the appropriate pots, labeled, and sealed in foil. Seeds used for this test included tomato seeds and mustard seeds (for bioassay). S. lycopersicum brt bushy root mutant tomato plants (LA2816) were obtained from the C.M. Rick Tomato Genetics Resource Center and the University of California Davis. These seeds were acid treated in 1% HCl. The wild-type tomatoes were Better Boy Hybrids from Burpee (Lot 1). Southern Giant Curled Mustard from Wetsel Incorporated (Lot 1185) was used for the bioassay. All tomato seeds were prepared before being transplanted into their soil pots. Forty of the 50 mutant seeds (quantity was very limited) and 40 wild-type seeds were placed in 2.7% sodium hypochlorite (half-strength bleach) in a 500 ml beaker for 30 minutes. Seeds were then rinsed and placed in plant trays lined with five layers of paper towel that was moistened and covered with five additional layers. Plant trays containing the seeds were placed in a warm dark location until germination. Seeds were then transplanted into soil pots, with two seeds per pot, planted

inch below the soil. Miracle Grow Water Soluble Fertilizer was prepared and added to each pot of soil every ten to fifteen days. The recommended dosage of one tablespoon of fertilizer to one square foot of soil was followed. Four thousand ml of liquid fertilizer was prepared and all was autoclaved. Five ml of the fertilizer was applied to each pot during each application. All test groups were watered with 50 ml of tap water three days a week or as needed. An alternate method of soaking the plant pots for ten seconds each (timed) using a constant stream of water was also used on occasion to keep the plants growing. As soon as plants were growing sufficiently, one plant was removed or transplanted so that only one plant remained per pot. Plants were allowed to grow for at least one month. Two different pesticides were obtained for this experiment from Dr. R. Allen Straw at Virginia Tech. Six pounds of Kelthane 50 Agricultural Miticide (Lot L2603), manufactured by Rohm and Haas Company with 50 percent dicofol and 50 percent inert ingredients was actually used in the test. Five pounds of Thionex 50 W (Endosulfan) was also obtained as an alternative to dicofol. The thionex contained 50 percent endosulfan and 50 percent inert ingredients and was manufactured by Makhteshim Agan of North America, Incorporated (Lot GM809016). Pesticide (Kelthane 50) was applied at two different times to provide the opportunity for phytoremediation. In powder form, 0.5 g of Kelthane was mixed into the soil of each test pot. After one month, an additional 1.0 g of Kelthane was added aqueously. These two applications simulated a large presence of dicofol initially and then additional dicofol being dumped at the remediation site. Thirty grams of Kelthane were added to 300 ml distilled water. The solution was heated and stirred and 2 ml of acetone forced the solution to combine. The acetone evaporated and 10 ml of the solution was added to each of the pots receiving pesticide. A pipette

pump was used to apply the solution and it was placed under the top layer of soil near the roots to minimize evaporation of the pesticide. Each day after all plants emerged from the soil, plant height was recorded. Health was also recorded using photographs for comparison purposes only. Height was measured in cm from the point where the stem meets the dirt to the last branch on the stem of the plant. The distance from where ruler starts to the zero point, when subtracted from the recorded height, gave accurate readings. At the time the solution of pesticide was added, leaf area was measured. After the solution of pesticide was added, a month went by until the plants were removed. Health was again recorded with a photograph. Final height, leaf area, chlorophyll concentration, Brix concentration, plant dry mass, and root wet mass were measured. Leaf area and chlorophyll concentration measured using below methods. Height measured using above method. Leaf area used the top leaf of the tomato plant farthest from the stem of the plant. Photographs were taken of the largest leaf on the highest petiole, removing the end leaflet. Include a square reference block in each photograph. This test used sticky notes with an area of 7.6 square cm. Imported photographs were cropped to allow plant and block to be shown. Adobe Photoshop Elements 6.0 software was used to find leaf area. Using the magnetic marquee tool, select the perimeter of each leaf. In the pallet toolbar, open the histogram. Expand and refresh. Leaf pixels should be recorded for each leaf. Select the block of known size and determine the number of pixels. Use the following equation to determine the square centimeter area of the plant: {[(Plant pixels total)/(Block Pixels)] x 7.6 sq cm}/(number of plants)=square centimeters of leaf area. These calculations were performed using Microsoft Excel. Chlorophyll content was analyzed to determine health. This required testing leaves from every plant. The leaves used for leaf area were retained and weighed for use in chlorophyll

concentration testing. These tests occurred only thirty minutes after the leaves were removed and leaves were refrigerated during this time. Put leaf tissue into a mortar and add 5 ml 91% isopropyl alcohol. Pulverize tissue with a pestle; the result is the leaf homogenate. Filter the extract and collect it in a test-tube. Let the extract settle for a few minutes. A UV/VIS NanoDrop Spectrometer was used to measure absorbance. A 2 micro liter sample from each plant was micro-pipetted in the spectrometer. The Nano-Drop was first zeroed using distilled water blank. The entire spectrum of light was recorded on a computer connected to the Nano-Drop. Wavelengths were recorded at A663 and A645 for use in Arnons equation, but general chlorophyll trends were also observed. Repeat for other extracts. Calculate using Arnons equation to convert absorbance measurements to mg Chl g-1 leaf tissue. Equation used: Chl a (mg g-1) = [(12.7 x A663)-(2.6 x A645)] x (ml alcohol / mg leaf tissue). Chl b (mg g-1) = [(22.9 x A645)-(4.68 x A663)] x [ml alcohol / mg leaf tissue]. Total Chl=Chl a+Chl b. Chlorophyll concentrations were compared and equation was computed using Microsoft Excel. Plants were removed from the soil as carefully as possible using a scoopula to minimize broken roots. The roots were separated from the plants using scissors. Plants and roots were placed in separate bags. Plants were air dried for fifteen days and then dry massed. Roots were wet massed and immediately frozen to prevent sucrose degradation. A refractometer was desired to measure the Brix (sucrose) concentration in the plant roots. This device was available at school, but the teacher lost it before it was used. Instead, because the sucrose concentration decreases with time, a hydrometer was used. The frozen roots were air warmed for five minutes. Each root was mashed with 5 ml of tap water (which was verified to contain no sucrose) in a mortar for about thirty seconds. The pulp was measured in a graduated

cylinder with 5 ml of extra water added. This solution was placed into a one-inch diameter 18inch long clear plastic tube, stopped at one end. One hundred and forty additional ml of water were added to allow the hydrometer to float and measure the Brix concentration. The amount of root pulp, the amount of added water, and the Brix reading were all recorded. This data was used to create a proportion of solution volume to Brix reading to calculate the real Brix solution of the roots as opposed to the diluted solution. The validity of this method was tested using grape juice at various dilutions and converting them to regular strength to determine if the diluted solution could provide accurate readings of the grape juice. The above procedure was used after the grape juice test was deemed valid. The soil was analyzed to see how much of the pesticide exists when compared to the control with just the miticide. The method of bioassay was used because it was deemed reliable from previous testing. The technique of using a bioassay was instrumental in the completion of this experiment. A bioassay was the main method of testing the amount of dicofol remaining in soil samples to quantitatively determine how much dicofol remained and how effective tomato plants were at phytoremediating. There is little available research about the method of bioassays. Orcutt (2010) cautions that there is not much literature that dictates proper bioassay method (Orcutt, 2010). Thus, part of this experiment was determining a proper bioassay method (Orcutt, 2010). A simple definition of a bioassay is a method for estimating the potency of a drug or materialby utilizing the reaction caused by its application to experimental subjects (Govindarajulu, 2001). The bioassay is a new method of testing, developed in the 1940s (Govindarajulu, 2001). Key to successful bioassays is creating a standard data set with known amounts of chemical for which

the sample data sets are compared (Govindarajulu, 2001). Thus, the bioassay is an inexpensive and easy method of testing soil. To prepare the bioassay, a baseline test was conducted. Pots of soil were prepared as described above. This means that 3230 g of soil (170 g per pot) were autoclaved. Nineteen pots were used. Each pot was given varying amounts of Kelthane, from 0 grams to 1.8 grams, increasing by 0.10 grams. The pesticide was massed and mixed in powder form into each sample of soil for one minute. Forty mustard seeds were added to each pot. Mustard seeds were chosen because they have been known to be effective indicators of DDT (extremely similar to dicofol) (Orcutt, 2010). The number of plants that germinated was measured for twelve days. The results were compiled and averaged and one logistic equation for each day that was representative of the data was found to allow for estimation of the amount of dicofol in soil with relation to the number of seeds that germinated. Similar testing was repeated with the pots that had unknown amounts of Kelthane. Germination of mustard seeds was recorded and using the equations found in the baseline test, an average estimated amount of dicofol remaining in the soil was obtained. A different standard equation was used for each day of germination. If the logistic curve did not fit the number of seeds germinated, results were extrapolated. For example, if the lower bound for the equation was ten plants and one pot had four plants, the pot would be recorded as having the maximum (1.5 g) of Kelthane. After recording the daily amount of Kelthane remaining, the pots that had no Kelthane were used to standardize the data. A difference was taken between the germination of the pots with no Kelthane and those with Kelthane to obtain an accurate amount of Kelthane remaining. These results were averaged and t-tests tests were run. Data was compiled and statistical analysis performed to see changes in plant growth, leaf

area, chlorophyll concentration, and mustard seed germination. Averages were performed on appropriate data sets. T-tests and error analysis was also completed. A logistic function was used to fit the bioassay results. There were many constants used in the project. They included the amount of light, amount of water, temperature, amount of soil, number of seeds, amount of chemical, method of height, area, chlorophyll content, and analysis methods. The independent variable included the presence of chemical in tomato plants or in the soil. Growth of the resulting tomato plants, the amount of phytoremediation that occurred, the amount of chemical in plant, the amount of chemical in soil, the height of the plant, the health of plants recorded using photographic comparison, and the leaf area of plants are some examples of dependant variables. In order to keep the experiment controlled, three groupings: tomato plants without added chemical, soil with no chemical, and soil with added chemical were used. Results The hypothesis that bushy root mutated tomato plants would remove more Kelthane than wild-type tomato plants, but have more negative health effects, was not supported. In fact, the exact opposite result occurred. Bioassay results showed that autoclaved soil alone removed 0.384 grams of Kelthane, while the mutated plants removed 0.537 grams, and the wild-type removed 1.140 grams out of the total 1.50 grams (g) added. Wild-type plants removed significantly more Kelthane than mutated plants while mutated plants removed more, but not a significant amount more, Kelthane than soil alone. In terms of health, the mutant plants seemed to fair best. Mutant plants had a significantly greater percent increase in leaf area, 123% for those with Kelthane added, when compared to a

5.16% decrease for wild-type plants undergoing phytoremediation. Plants that were not phytoremediating increased leaf area at a steadier 41% to 61% rate. Percent change in plant height showed a similar that mutant plants grew taller without Kelthane (275% to 166%), while wild-type plants were significantly taller when phytoremediating (279% to 234%). Though not significant, mutant plants had more chlorophyll (0.458 g without Kelthane and 0.182 g with Kelthane) when compared to wild-type plants (0.203 g and 0.177 g). Mutant plants also had the highest Brix concentrations (121% and 3.61%), though the wild-type without Kelthane was significantly higher in Brix than the wild-type with Kelthane (39.1% and -5.63%). With plant dry mass, the mutant with no Kelthane had the highest mass (0.232 g) followed by the mutant with Kelthane (0.101 g). Finally, the mutant plants had the highest root masses (2.02 g and 1.59 g) when compared to the wild-type plants (0.777 g and 1.28 g). The brt mutation was once again investigated and additional progress was made in identifying the composition of this mutation. In collaboration with Dr. Benedito (2011), the researcher was given access to a newly completed tomato protein transporters list. From previous research (OBrochta, 2011), it was determined that the brt mutation occurred on the twelfth tomato chromosome at 19.8 cM (unit length of chromosome) or 95.8 cM. The gene at this location was TG296, a Lysr transcriptional regulator protein from bacteria that was placed in the castor bean plant. The interest was determining what changes this mutation brought about. Sidorva (2002, 2010) hypothesized that the brt mutation caused increased levels of Auxin in the plant roots. Auxin, a type of Brix (sucrose), was measured in this experiment and higher levels were found in the brt mutated plants (Coombe, 1960). Using the documentation from Dr. Benedito, the bacterial SDS degradation transcriptional activation protein was matched with its

counterpart in the tomato plant. The closest match for the protein is 2.A.2.4.1 from the Glycoside-Pentoside-Hexuronide (GPH) family (Benedito, 2011). In researching this protein, it was found that it promotes importation of sucrose into the flower in order to increase pollen growth (Stadler, et. al., 1999). Thus, the original hypothesis from Sidorva (2002) is shown to be correct. The higher Brix levels in the brt mutant tomato plants are caused by protein 2.A.2.4.1, which was inserted at 19.8 cM or 95.8 cM on the twelfth tomato chromosome. Discussions and Conclusions This experiment represents a much more comprehensive look at the phytoremediation of Kelthane when compared to three previous years of research. The amount of time that the plants grew was extended by a factor of eight and the amount of Kelthane was raised to more typical levels. Bioassay testing was also much improved, with additional precision. Mutated tomato plants were healthier, sometimes statistically so, when compared to wildtype plants and test groups without Kelthane were healthier than those undergoing phytoremediation. Russell (2005) supports this conclusion and notes that plants must have phytotoxicity, or ability to withstand the presence of dicofol, a factor that wild-type tomatoes typically do not have. Weaver (2010) warns that tomato plants are usually fairly phytotoxic and are used as bioindicators meaning that their health will be adversely affected by the presence of pesticides like Kelthane. The major finding from this experiment was that more effective phytoremediation occurred in wild-type tomato plants when compared to mutated tomato plants. The (mutant) root system is very highly branchedthe root system branches profusely within one day after emergence, in contrast to normal roots, which branch only after several days of growth (Zobel,

1971). This phenomenon may have actually hurt phytoremediation ability since the root branching causes stringier and less developed roots. Zobel also notes that brt mutated tomato plants germinate more slowly than wild-type plants (Voland and Zobel, 1988). The increased Brix concentration found in mutated plants seems to have contributed to plant health, but may have made enzyme transport more difficult (Peres, 2010). More sucrose means that both the fruit health and edibility increase; however, since sucrose concentration was only tested in the plant roots, any sucrose that may have accumulated in the leaves or fruit was overlooked. Relationships between the number of microbial enzymes and their effect on phytoremediation are currently being investigated (Benedito, 2011). Dr. Benedito created a microbial enzyme transporter list that contains the location of most tomato DNA sequences and allowed the researcher to match the enzyme qualities with DNA sequences in the tomato genome. The most significant set of conclusions come from confirming that the brt mutant increases sucrose levels through the roots and their modified protein transports sucrose to pollen in the flowers of the tomato plant. While initially believed that higher levels of sucrose and more root branching would lead to increased phytoremediation ability, it was found that the wild-type tomato plants were actually more effective at removing the Kelthane from the soil. In the past, it was thought that adding additional degradation transcriptional activation proteins to the brt mutant would increase phytoremediation effectiveness even beyond the hypothesized gain from the initial mutation. This is not the case. The research suggests that removing these sucrose promoters and transporters to levels below the wild-type will allow for additional phytoremediation ability. Thus, it is not the shoot and root branching that seems to induce phytoremediation, rather the lack of sucrose in the roots. Since the wild-type tomato plants were

not modified to increase sucrose concentration, they were able to phytoremediate at a greater rate. Since it was determined that decreased sucrose concentration would allow for additional phytoremediation, the researcher tried to locate the gene or protein that causes sucrose accumulation. The Sucrose Phosphate Synthase (SPS) enzyme is found in low levels in wild-type tomatoes (Miron and Schaffer, 1991). A low SPS level would mean less concentration of sucrose and more phytoremediation ability (Miron and Schaffer, 1991; Yelle, et. al, 1991). This is linked to a low level of the acid invertase enzyme (Yelle, et. al, 1988). The low level of invertase means high levels of sucrose and no SPS, causing more starch, and lower phosphorylase, all of which lead to more phytoremediation (Yelle, et. al, 1988). The acid invertase alters composition in the gene TIV1, which is part of the sucrose accumulation group of genes (Klaan, et. al, 1996; Klaan, et. al, 1993). Chetelat (et. al, 1993) states that an increase in TIV1 or the similar gene TG102 will cause an increase in acid invertase, meaning more phytoremediation. From tomato chromosome identification, it was determined that TG102 is located on chromosome 3 at 56.79 cM and TIV1 is located between 56.62 cM and 56.96 cM (Chetelat, et. al, 1993). The DNA sequence of TG102 was run in the protein BLAST database that identified a self-incompatibility RNase protein in tomato plants that will prevent self-fertilization within the species. The overall conclusion is that increasing the amount of the self-incompatibility RNase and, thus, the TG102 DNA sequence, should cause increased phytoremediation ability. The most appropriate extension to this project is to find a tomato mutant that has been modified and to repeat the experiment with this new mutant, the brt mutant, and the wild-type tomato. So far, it has not been possible to find such a mutant, as the removal of sucrose is not

desirable for tomato fruit. That being said, the conclusion from this experiment can still be tested using the tomato mutant sucr (TGRC LA4104) that is the mutant containing increased sucrose concentration. A phytoremediation study with this mutant should, according to the results of this experiment, provide the least amount of phytoremediation of all the tested tomato groups. With the conclusions from such an experiment, the worth of mutating a wild-type tomato in order to remove all the sucrose related enzymes could be examined, since it appears that no mutant with sucr removed exists. That should, hopefully, provide evidence for the ideal tomato mutation to maximize phytoremediation of Kelthane, dicofol, and DDT. Literature Cited Arms, K. (2004). Environmental science. Austin, Texas: Holt, Rinehart, and Winston. Benedito, V. "Tomato phytoremediation of dicofol." Message to researcher. 2010. E-mail. Benedito, V. Tomato predicted transporters. Document from researcher. 2011. E-mail. Burtt, B. (2009, October 27). UW firm uses plants to clean contamination. The Guelph Mercury. Bush, C. (n.d.). Stress tolerant plants. Retrieved from http://arabidopsis.info/students/stress/stresshome.html. Cabras, P., Cabitza, F., Meloni, M., & Pirisi, F.M. (1985). Behavior of some pesticide residues on greenhouse tomatoes. 2. fungicides, acaricides, and insecticides. Journal of Agricultural and Food Chemistry, 33, 935-937. Cherian, S., & Oliveria, M.M. (2005). Transgenic plants in phytoremediation: recent advances and new possibilities. Environmental Science and Technology, 39(24), 9377-9390. Chetelat, R. (2010). Revised list of monogenic stocks. Davis, CA: C.M. Rick Tomato Genetics Resource Center, Department of Plant Sciences: University of California, Davis.

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Washington, DC: Technology Innovation Office. Voland, M.L., & Zobel, R.W. (1988). A morphologic and genetic characterization of two tomato root mutants. In C. Rick (Ed.), Report of the tomato genetics cooperative (pp. 47). Ithaca, NY: Departments of Plant Breeding and Biometry, and Agronomy: Cornell University. Watts, M. (2008, October 01). Breast cancer: the link with pesticides. Women & Environments International Magazine, 76. Weaver, M.J. "Tomato phytoremediation of dicofol." Message to researcher. 2010. E-mail. Wiley, N. (2007). Phytoremediation: methods and reviews. Totowa, New Jersey: Humana Press Inc. Withgott, J., & Brennan, S. (2008). Environment: the science behind the stories. San Francisco, CA: Pearson: Benjamin Cummings. Yelle, S., Chetelat, R. T., Dorais, M., DeVerna, J. W., & Bennett, A. B. (1991). Sink metabolism in tomato fruit: IV. Genetic and biochemical analysis of sucrose accumulation. Journal of Plant Physiology, 95, 1026-1035. Yelle, S., Hewitt, J. D., Robinson, N. L., Damon, S., & Bennett, A. B. (1988). Sink metabolism in tomato fruit: III. Analysis of carbohydrate assimilation in a wild species. Journal of Plant Physiology, 87, 737-740. Zobel, R. "Tomato phytoremediation of dicofol." Message to researcher. 2010. E-mail. Zobel, R.W. (1971). Root mutants of the tomato. Report of the tomato genetics cooperative, 21, 42. Zobel, R.W., Wright, S.F., Al-Amoodi, L.K., Barbarick, K.A., Roberts, C.A., & Dick, W.A. (Ed.). (2005). Roots and soil management: interactions between roots and the soil.

Madison, WI: American Society of Agronomy, Inc.; Crop Science Society of America, Inc.; Soil Science Society of America, Inc. Zsogon, A., Lambais, M.R., Benedito, V.A., de Oliveria Figueria, A.V., & Peres, L.E.P. (2008). Reduced arbuscular mycorrhizal colonization in tomato ethylene mutants. Scientia Agricola, 65(3), 259-267.

Acknowledgements The experimenter would like to acknowledge many researchers and professors from various institutions that provided great advice for this project. This includes Dr. R. Allen Straw who worked for months to find the appropriate pesticides for the experiment. Dr. Vagner Benedito was instrumental in completing the brt location extension. Dr. Jonathan Watkinson and Roanoke College were both extremely helpful in obtaining the correct mutant tomato seeds. Dr. David Orcutt was a huge help in determining the bioassay method and providing general advice. Mrs. Cindy Bohland was always available to answer questions or help me with procedures. Great insight and advice was also received from the following people: Mr. Glenn Ferrand and Drexel Chemical Company Incorporated, Dr. Richard Zobel, Dr. Roger Chetelat and the C. M. Rick Tomato Genetics Center at U.C. Davis, Mr. Darren Cribbes, Dr. Michael Weaver, Mr. Keith Rose, Dr. Bernard Glick, Dr. Saleh Shah, Mr. Barry Robinson, Mr. Dennis Anderson, Mr. David Richert, Dr. Andrew Thompson, Ms. Patty Webb, Dr. Victoriano Gutirrez, Dr. Lazaro E. P. Peres, Mr. Paul Foran and Dow AgroSciences, Ms. Linda Fiedler, Dr. Priscilla Gannicott, Dr. Donald Mullins, Ms. Tricia Stoss, Dr. J.O. Rogers, Mr. Greg Evanylo, Mr. Wythe Morris, Dr. Kari Benson, Dr. Jim Westwood, Dr. Darwin Jorgensen, Dr. Anthony Curtis, Dr. Linda Gooding, Mr. Steven Smith, Dr. David Glass, Dr. Audil Rashid, Dr. Laura Carreira, Dr. Zhi-Qing Lin, Dr. J.W. Scott, and many others. The experimenter would like to give special thanks to his parents and research instructor who were instrumental in the success and funding of this project.

Appendix

Graph 1: Logger Pro Generated Graph of Number of Mustard Seeds Germinated (number) vs. Amount of Kelthane (g/pot) on Day 7. Error bars of 5% error are shown. Curve was automatically fit and then tweaked so that it fit the data better. No points were stricken because the number of points above and below the curve were about equal.

Graph 2: Logger Pro Generated Graph of Number of Mustard Seeds Germinated (number) vs. Amount of Kelthane (g/pot) Through Logistic Curve Fit by generating points fitting the equation for every 0.1 g/pot. This curve was used to estimate the amount of dicofol remaining only on day 7. Different curves were generated each day using the baseline data to get a better representation of the amount of Kelthane.

Histogram 1: Histogram of Kelthane Remaining in Mutant Plant Soil. Note that the vast majority of plants had between 1.4 and 1.5 grams remaining and the table is skewed left, meaning that higher amounts remaining are more typical.

Histogram 2: Histogram of Kelthane Remaining in Wild-Type Plant Soil. Note that the vast majority of plants had between 0.0 and 0.1 grams remaining and the table is skewed right, meaning that lower amounts remaining are more typical.

Chart 1: Linear Relationship Plot of Change in Leaf Area (%) vs. Change in Plant Height (%). The mutant with Kelthane shows an opposite relationship from the other groups for the increasing percent change in leaf area is related to an increasing percent change in

plant height.

Chart 2: Linear Relationship Plot of Final Leaf Area (sq. cm) vs. Initial Leaf Area (sq. cm) All groups showed a positive relationship between initial and final area, but the mutant without Kelthane had the least directly linear relationship.

Chart 3: Linear Relationship Plot of Root Wet Mass (g) vs. Plant Dry Mass (g). All the

groups show a positive relationship between root and plant masses, but the mutant without Kelthane had the most extreme value, while the others were similar in slope.

Chart 4: Linear Relationship Plot of Brix (%) vs. Total Chlorophyll (mg). The horizontal lines show that all the groups except the mutant without Kelthane had no relation between Brix and chlorophyll. The mutant without Kelthane group shows a slight relationship, but it is probably a testing precision problem.

Chart 5: Boxplot of Amount of Kelthane Remaining in Soil. Lower numbers indicate more

phytoremediation; higher numbers indicate less phytoremediation. Seventy-five percent of Wild-type plants had more phytoremediation than only twenty-five percent of mutant plants.

Chart 6: ANOVA showing significant difference between wild-type and mutant amounts of Kelthane remaining.

Figure 1: Comparison Of Chemical Structures-Dicofol On Left, DDT On Right-To Show Their Similarities (Drawings-PubChem)

Figure 2: Inside picture of second autoclave used.

Figure 3: Bleaching tomato seeds for easier germination.

Figure 4: Kelthane powder (white) ready to be mixed in soil.

Figure 5: Autoclaved fertilizer.

Figure 6: Experimental set-up.

Figure 7: Seed germination on paper towels.

Figure 8: Growing tomato plants.

Figure 9: Kelthane liquid solution that was mixed before application.

Figure 10: Sample leaf area picture with standard sticky-note reference block.

Figure 11: Mustard bioassay used for logistic curve generation and unknown Kelthane levels.

Figure 12: Sample root used in Brix and mass testing.

Figure 13: Chlorophyll concentration curves generated by Nano-Drop Spectrometer.

Figure 14: Chlorophyll concentration set-up on Nano-Drop Spectrometer.

Figure 15: Grinding up roots for Brix concentration testing.

Figure 16: Sample root Brix concentration showing a 30% Brix concentration.

Figure 17: Drying soil pots for storage to allow for Kelthane degradation.

Figure 18: Drying plants for mass measurements.