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.
Chetelat, R. T., DeVerna, J. W., Klann, E., & Bennett, A. B.
(1986). Sucrose accumulator (sucr), a gene controlling sugar
composition in fruit of L. chmielewskii and L. hirsutum. Report of
the tomato genetics cooperative, 43, 14-16. Chu, W.K., Wong, M.H.,
& Zhang, J. (2006). Accumulation, distribution and
transformation of DDT and PCBs by Phragmites australis and Oryza
sativa L.: I. Whole plant study. Environmental Geochemistry and
Health, 28, 159-168. Coombe, B. G. (1960). Relationship of growth
and development to changes in sugars, auxins, and gibberellins in
fruit of seeded and seedless varieties of Vitis vinifera. Plant
Physiology, 35(2), 241-250. Crane, C. (2009, September 21).
Cleaning up soiled sites. Science World, 66(2), 6. Cutraro, J.,
& Goldstein, N. (2005, August 01). Cleaning up contaminants
with plants. Bicycle, 46(8), 30. Eckley, N. (2001). Traveling
toxics. Environment, 43(7), 24. Evans, LD. (2002). The dirt on
phytoremediation. Journal of Soil and Water Conservation, 57(1),
12A. Gao, J., Garrison, A.W., Hoehamer, C., Mazur, C.S., &
Wolfe, N.L. (2000). Uptake and phytotransformation of o,p-DDT and
p,p-DDT by axenically cultivated aquatic plants. Journal of
Agricultural and Food Chemistry, 48, 6121-6127. Garber, K., &
Peck, C. Office of Pesticide Programs, Environmental Fate and
Effects Division. (2009). Risks of dicofol use to federally
threatened California red-legged frog (rana aurora draytonii).
Washington, D.C. Gardea-Torresdey, JL. (2003, April 01).
Phytoremediation: where does it stand and where will it
go? Environmental Progress. Gerhardt, K.E., Huang, X-D., Glick,
B.R., & Greenberg, B.M. (2008). Phytoremediation and
rizoremediation of organic soil contaminants: potential and
challenges. Plant Science, 176(1), 20-30. Govindarajulu, Z. (2001).
Statistical techniques in bioassay. Basel, Switzerland: S. Karger.
Greenberg. (2006). Newsletter for the NSERC CRD multi-process
phytoremediation system. Phytoremediation News, 2, 1-5. Kelthane
50W agricultural miticide. (2005). Dow AgroSciences Canada. Klann,
E. M., Chetelat, R. T., & Bennett, A. B. (1993). Expression of
acid invertase gene controls sugar composition in tomato
(Lycopersicon) fruit. Journal of Plant Physiology, 103, 863-870.
Klann, E. M., Hall, B., & Bennett, A. B. (1996). Antisense acid
invertase (TIV1) gene alters soluble sugar composition and size in
transgenic tomato fruit. Journal of Plant Physiology, 112,
1321-1330. Material safety data sheet: Kelthane 50W agricultural
miticide. (2008). Dow AgroSciences Canada. Miron, D., &
Schaffer, A. A. (1991). Sucrose phosphate synthase, sucrose
synthase, and invertase activities in developing fruit of
Lycopersicon esculentum Mill. and the sucrose accumulating
Lycopersicon hirstum Humb. and Bonpl. Journal of Plant Physiology,
95, 623-627 O'Brochta, W. J. (2011). Phytoremediation comparisons
between Solanum lycopersicum wildtype and brt mutatnt using
kelthane miticide. Report of the Tomato Genetics Cooperative,
61, 5-11. Orcutt, D. "Testing plant samples." Message to
researcher. 2010. E-mail. Peres, L.E.P. "brt tomato mutant."
Message to researcher. 2010. E-mail. Qiu, X., Zhu, T., Yao, B., Hu,
J., & Hu, S. (2005). Contribution of dicofol to the current DDT
pollution in China. State Key Joint Laboratory for Environmental
Simulation and Pollution Control. Report of the tomato genetics
cooperative. (1951-2011). University of Florida, Cornell
University, University of California Davis: Tomato Genetics
Resource Center. Russell, K. U.S. Environmental Protection Agency,
Office of Solid Waste and Emergency Response: Technology Innovation
and Field Services Division. (2005). The use and effectiveness of
phytoremediation to treat persistent organic pollutants.
Washington, D.C. Rose, K. "Tomato phytoremediation of dicofol."
Message to researcher. 2010. E-mail. Rossi, L.A. United States
Environmental Protection Agency, Office of Pesticide Programs:
Special Review and Reregistration Division. (1998). Reregistration
eligibility decision (RED): dicofol (EPA 738-R-98-018). Washington,
DC. Snchez, A.I., Hernando, M.D., Vaquero, J.J., Garca, E., &
Navas, J.M. (2010). Hazard assessment of alternatives to dicofol.
Journal of Environmental Protection, 1, 231-241. Setia, R.C., Kaur,
N., Setia, N., & Nayyar, H. (2008). Heavy metal toxicity in
plants and phytoremediation. In R.C. Setia, H. Nayyar, and N. Setia
(Eds.), Crop improvement: strategies and applications (pp.
206-218). New Delhi, India: I.K. International Publishing House
Pvt. Ltd. Shimizu-Sato, S., Tanaka, M., & Mori, H. (2009).
Auxin-cytokinin interactions in the control of
shoot branching. Plant Molecular Biology, 69, 429-435. Sidorva,
K.K., Shumny, V.K., Vlasova, E.Yu., Glyanenko, M.N, Mishehenko,
T.M. (2002). The brt (branched roots) and lrt (long roots) genes
control the development of roots in peas (pisum sativum L.). Pisum
Genetics, 34, 23-25. Sidorova, K.K., Shumny, V.K., Vlasova, E.Yu.,
Glyanenko, M.N, Mishehenko, T.M., Maystrenko, G.G. (2010). Genetics
of symbiosis and breeding of a macrosymbiont for intense nitrogen
fixation by the example of pea. , 14(2), 357-374. (Translated from
Russian). Singh, R.P., Dhania, G., Sharma, A., & Jaiwal, P.K.
(2007). Biotechnological approaches to improve phytoremediation
efficiency for environment contaminants. In S. Singh and R.
Tripathi (Eds.), Environmental bioremediation technologies (pp.
223-258). Berlin, Germany: Springer Science+Business Media. Smith,
C., Kerr, K., & Sadripour, A. (2008). Pesticide exports from
U.S. ports, 2001-2003. International Journal of Occupational and
Environmental Health.Stadler, R., Truernit, E., Gahrtz, M., &
Sauer, N. (1999). The AtSUC1 sucrose carrier may represent the
osmotic driving force for anther dehiscene and pollen tube growth
in Arabidopsis. The Plant Journal, 269-278.
Thieman, W.J., & Palladino, M.A. (2009). Introduction to
biotechnology. San Francisco, CA: Pearson: Benjamin Cummings.
Terry, N., & Bauelos, G. (2000). Phytoremediation of
contaminated soil and water. Boca Raton, FL: Lewis Publishers, an
imprint of CRC Press LLC. Thompson, A. "bushy root." Message to
researcher. 2010. E-mail. United States Environmental Protection
Agency, Office of Solid Waste and Emergency Response. (2001). A
citizen's guide to phytoremediation (EPA 542-F-01-002).
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.