NITROGEN TRANSFORMATIONS IN THE VADOSE ZONE OF A SMALL, ZERO-ORDER
PIEDMONT WATERSHED IN WATKINSVILLE, GEORGIA
by
JAMES FRANK MUCKLER
(Under the Direction of John F. Dowd)
ABSTRACT
Soil water samples were collected in a small, Piedmont watershed in nine suctionlysimeters at four depths in the vadose zone: 0.35, 0.5, 1.25, and 1.75 m, and analyzed for
nitrate-N, ammonium, urea, total nitrogen, dissolved organic carbon (DOC), and ferrous and
ferric iron concentrations. Soil water potential was inferred from pressure heads measured in
tensiometers at 0.5, 1.0, and 1.5 m depths. Although nitrate-N concentrations were spatially
variable and periodically high (20 – 64 mg/L) at 0.35 and 0.5 m depths, nitrate-N consistently
decreased with depth to the deepest lysimeter at 1.75 m (0 – 6 mg/L), which was likely due to
denitrification. Also, DOC decreased with depth. Other geochemical trends were not so clear in
the limited data. Stable nitrogen isotope ratios in soils were compared to possible nitrogen
sources that found manure-source signatures in shallow soils and chemical fertilizer-source
signatures in deeper soils.
INDEX WORDS:
Nitrogen Transformations, Nitrate Profiles, Vadose Zone, Grazed Pasture,
Watershed
NITROGEN TRANSFORMATIONS IN THE VADOSE ZONE OF A SMALL, ZERO-ORDER
PIEDMONT WATERSHED IN WATKINSVILLE, GEORGIA
by
JAMES FRANK MUCKLER
B.S., University of Missouri, 2003
A Thesis Submitted to the Graduate Faculty of The University of Georgia in Partial Fulfillment
of the Requirements for the Degree
MASTER OF SCIENCE
ATHENS, GEORGIA
2010
© 2010
JAMES FRANK MUCKLER
All Rights Reserved
MEASURING NITROGEN TRANSFORMATIONS IN THE VADOSE ZONE OF A SMALL,
PRIMARY PIEDMONT WATERSHED IN WATKINSVILLE, GEORGIA
by
JAMES FRANK MUCKLER
Electronic Version Approved:
Maureen Grasso
Dean of the Graduate School
The University of Georgia
July 2010
Major Professor:
JOHN F. DOWD
Committee:
DAVID B. WENNER
DINKU M. ENDALE
DEDICATION
I would like to dedicate this to my parents and family for their support and inspiration, to
the love of my life Kristy Plattner for everything, and my friends for their support, especially
Robert Joseph McKinnon.
iv
ACKNOWLEDGEMENTS
I would like to acknowledge the USDA-Agricultural Research Service J. Phil Campbell
Sr. Natural Resource Conservation Center for permission and use of the field site known as
watershed 2. I would like to thank Dr. Dinku Endale who welcomed countless meetings in his
office to discuss even the smallest aspects of this study, met me in the field to offer help and
resources, and to provide encouragement. Also, I would like to thank Stephen Norris for all his
time and wisdom in the field.
I would like to acknowledge the USEPA-ORD for use of the Nutrients Laboratory under
the direction of Dr. Caroline Stevens. I would like to thank Dr. Caroline Stevens for use of her
budget for the lab until I almost exhausted it, for support in this project, and the knowledge she
passed along. I would like to thank Lidia Samarkina who is one of the most knowledgeable
people I have ever met in a laboratory for all her hard work and time instructing me in all the
chemical analyses. I would also like to thank Kathy Schroer for all our discussions of nitrogen,
for tips in the lab, for use of her data in comparison to mine, and for all her great help.
I would like to acknowledge Tom Maddox and the UGA Ecology Analytical Chemistry
Laboratory for analyzing the nitrogen isotopic ratios of my soil samples. The results proved to
be very interesting so thanks to Tom and his staff.
I would also like to acknowledge and thank the Miriam Watts- Wheeler Fund in the
Department of Geology for the funding of this project. I would like to thank Dr. Wenner for his
help and knowledge of geochemistry and isotopes. Also, special thanks to Dr. John Dowd for
never being too busy to discuss, help, and educate me at all stages of this project. His help in the
v
field with set up and his amazing knowledge guided me through this project.
vi
TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS .............................................................................................................v
LIST OF TABLES ...........................................................................................................................x
LIST OF FIGURES ....................................................................................................................... xi
CHAPTER
1
INTRODUCTION .........................................................................................................1
1.1. Background ........................................................................................................1
1.2. Purpose of the study ...........................................................................................4
1.3. Limitations of the study ......................................................................................5
2
LITERATURE REVIEW ..............................................................................................7
2.1. Agricultural Nitrogen Fertilizer Application ......................................................7
2.2. Non-Point Source Nitrogen Pollution from Agriculture ....................................8
2.3. Nitrogen Fertilizer Application Thresholds .....................................................10
2.4. Manure, Urea, and Nitrogen .............................................................................11
2.5. The Agricultural Nitrogen Cycle ......................................................................16
2.6. Vadose Zone Water and Nitrogen Movement ..................................................23
2.7. Wetting and Drying Cycles ..............................................................................26
2.8. Darcy’s Law and the Darcy-Buckingham Equation.........................................28
2.9. The Piedmont Region .......................................................................................29
3
MATERIALS AND METHODS .................................................................................31
vii
3.1. Experimental Site, Land Use, and Soil.............................................................31
3.2. Tensiometers and Pressure Transducers...........................................................37
3.3. Suction Lysimeters ...........................................................................................42
3.4. Backfilling with Slurry .....................................................................................45
3.5. Operation of Lysimeters ...................................................................................46
3.6. Geochemical Analyses .....................................................................................46
3.7. Soil Analyses ....................................................................................................48
3.8. Time Domain Reflectometry and Water Table Monitoring .............................48
3.9. Nitrogen Stable Isotope Analyses ....................................................................49
4
RESULTS AND DISCUSSION ..................................................................................52
4.1. Introduction ......................................................................................................52
4.2. Soil Analyses ....................................................................................................52
4.3. Fertilization of Watershed 2 .............................................................................54
4.4. Moisture Release Curves, TDR, and Tensiometric Data .................................57
4.5. Soil Water, Groundwater, and Precipitation Geochemical Results..................61
4.6. Mineralization-Immobilization ........................................................................79
4.7. Nitrification ......................................................................................................81
4.8. Denitrification ..................................................................................................82
4.9. Terminal Electron Acceptors and Oxidation-Reduction Reactions .................85
4.10. Effects of a Seasonably Elevated Water Table...............................................86
4.11. Evapotranspiration..........................................................................................88
4.12. Plant Uptake ...................................................................................................88
4.13. Non-Biological Nitrogen Transformations ....................................................89
viii
4.14. Wetting and Drying Cycles ............................................................................90
4.15. Soil Bacteria Population Cycles and Nitrogen Transformations....................92
4.16. Excessive Nitrogen Fertilizer Application in W2 ..........................................93
4.17. 15N End-Member Analyses: Manure or Fertilizer Nitrogen Source ..............95
5
CONCLUSIONS..........................................................................................................99
5.1. Summary ..........................................................................................................99
5.2. Suggestions of Future Work ...........................................................................101
BIBLIOGRAPHY ........................................................................................................................103
APPENDICES .............................................................................................................................115
A
Depth to Water Table Data for Piezometers Surrounding W2 Field Site ..................115
B
TDR and Precipitation Data for W2 Field Site ..........................................................117
C
Wiring Diagrams for Tensiometers ...........................................................................122
D
Field Site Soil pH Profiles and Particle Size Distribution Analyses .........................124
E
W2 Management: Fertilizing, Liming, Spraying, and Planting .................................126
F
Geochemical Results..................................................................................................128
ix
LIST OF TABLES
Page
Table 4.1: Particle Size Distribution Analyses ..............................................................................54
Table 4.2: USDA-ARS Watershed 2 Fertilization Management ...................................................56
Table 4.3: Soil Water Summary Statistics with Depth ..................................................................71
Table 4.4: Soil Water Summary Statistics for Each Chemical Variable .......................................72
Table 4.5: Groundwater Concentrations for Piezometers 1, 2, 3, 6, and 7 ....................................73
Table 4.6: Geochemical Analyses of Precipitation ........................................................................76
Table 4.7: Soil Water Average Concentration Values with Depth under Tree Canopy ................84
Table 4.8: Soil Water Average Concentration Values with Depth under Open Sky .....................84
Table 4.9: Soil Water Average Concentration Values with Depth in High Nitrogen Area ...........91
Table 4.10: 15N Isotope Analyses on Nitrogen Source in Field Site Vadose Zone Soils ..............97
x
LIST OF FIGURES
Page
Figure 2.1: Soil Water Nitrate-N Concentrations at (a) 60 and (b) 120 cm Depth vs. Time .........14
Figure 2.2: Soil Water Nitrate Mass Loads (Pan Lysimeters at 60 cm Depth) vs. Time ..............15
Figure 2.3: The Nitrogen Cycle for Agriculture ............................................................................16
Figure 3.1: USDA-ARS Watershed 2 Elevation Contour Map .....................................................32
Figure 3.2: Field Site in Watershed 2 ............................................................................................36
Figure 3.3: Faybishenko-Designed Tensiometer Schematic .........................................................39
Figure 4.1: Soil pH vs. Depth for W2 Field Site Soils ..................................................................53
Figure 4.2: Cecil Soil Moisture Release Curves at 0.5, 1.0, & 1.5 m Depths ...............................57
Figure 4.3: TDR Soil Moisture vs. Time vs. Depth .......................................................................58
Figure 4.4: Hydraulic Gradients and Precipitation vs. Time .........................................................59
Figure 4.5: Field Site Map in W2 ..................................................................................................62
Figure 4.6: Soil Water Nitrate-N Concentrations vs. Depth ..........................................................63
Figure 4.7: Soil Water Ammonium Concentrations vs. Depth ......................................................65
Figure 4.8: Soil Water Total Nitrogen Concentrations vs. Depth..................................................66
Figure 4.9: Soil Water Dissolved Organic Carbon Concentrations vs. Depth...............................67
Figure 4.10: Soil Water Ferrous Iron Concentrations vs. Depth ...................................................69
Figure 4.11: Soil Water Ferric Iron Concentrations vs. Depth ......................................................70
Figure 4.12: Lysimeters L1, L1A, L2, L2A, & L2B Nitrate-N Concentrations vs. Time .............77
Figure 4.13: Lysimeters L3, L3A, L4, & L4A Nitrate-N Concentrations vs. Time ......................78
xi
Figure 4.14: Seasonal Depth to Water Table Measured from All Field Site Piezometers ............87
xii
CHAPTER 1
INTRODUCTION
1.1. Background
After the U.S. Congress passed the Clean Water Act in 1972, the Environmental
Protection Agency began enforcing the act. The Clean Water Act (CWA) established the basic
structure for regulating discharges of pollutants into the water of the United States and regulating
quality standards for surface waters. The CWA was based on the Federal Water Pollution
Control Act passed in 1948, the first major U.S. law to address water pollution, and came to be
known as the “Clean Water Act” after amendments to the original law passed in 1977. The
CWA made discharge of any pollutant into navigable waters illegal unless an initial permit was
obtained (USEPA, 2009).
Phase I of the CWA focused on point source pollution such as discrete conveyances like
pipes or man-made ditches, while Phase II of the CWA focused on non-point source pollution.
In the last decade, Phase II regulations have focused more on a holistic approach to watershedbased strategies than on a program-by-program, source-by-source, pollutant-by-pollutant
approach (USEPA, 2009). The latest focus of the CWA is on watershed-based strategies
especially understanding pollution on a watershed scale, which is a major reason for this work on
investigating nitrogen transformations in the vadose zone of a primary watershed.
Nitrogen is the most common and widely used fertilizer nutrient (Follett and Walker,
1989), and nitrate-N is the most common contaminant in groundwater (Freeze and Cherry,
1979). Ground water exhibiting nitrate contamination has been noted in every state in the U.S.
1
(Hallberg, 1989). In areas of intensive farming, stream water pollution by nitrogen species,
especially nitrate, is due to overabundant organic and mineral fertilization (Beaujouan et al.,
2001).
Excessive nitrogen loading causes include, but are not limited to, fertilizer application,
nitrogen fixation by legumes, human and animal waste disposal, and fossil fuel combustion
(Vitousek et al., 1997; Peterson et al., 2001). Human alteration to the nitrogen cycle has also
approximately doubled the rate of nitrogen input into terrestrial ecosystems, increased nitrogen
oxides that drive the formation of photochemical smog, accelerated losses of plant diversity
which use nitrogen efficiently, caused losses of soil nutrients (such as calcium and potassium)
which are essential for maintaining soil fertility, greatly increased the transfer of nitrogen
through rivers to coastal oceans, increased the quantity of organic carbon stored within terrestrial
ecosystems, and contributed to the acidification of soils, streams, and lakes to name a few
(Vitousek et al., 1997).
Nitrate contributes to contamination of surfaces waters through transport of nitrate-rich
groundwater base flow to streams and lakes (Hallberg, 1989). Excessively high nitrogen
concentrations in groundwater that discharges to surface water can cause eutrophication
(Carpenter et al., 1998) and can lead to excessive algal growth (Abit et al., 2008). Excessive
algal growth can lead to less biodiversity by unbalancing an ecosystem (Horrigan et al., 2002;
Vitousek et al., 1997). Background concentrations for nitrate have been debated with consensus
being between 2 – 3 mg/L. Concentration levels above this threshold could indicate human
inputs (Madison and Brunett, 1985; Muller and Helsel, 1996; Burkart and Stoner, 2007).
Nitrogen is one of the most vital of plant nutrients, and most crops remove more nitrogen
than any other nutrient. For this reason, the amount of fertilizer-N applied far exceeds the
2
application of other nutrients. Nitrate is highly soluble and very mobile, which facilitates plant
uptake, but also makes nitrate very vulnerable to leaching through the soil with infiltrating water
(Hallberg, 1989).
Nitrogen species from zero-order watersheds have a major influence on nitrogen delivery
to headwater streams. Headwater streams in turn deliver water and nutrients to larger streams,
which discharge ultimately into the ocean. Despite the relatively small dimensions of headwater
streams, these primary streams play a disproportionately large role in N transformations on the
landscape. Headwater streams retain and transform important amounts of inorganic nitrogen,
often more than 50% of watershed nitrogen. Also, small streams, with widths of 10 meters or
less, can comprise up to 85% of total stream length in a drainage network (Peterson et al., 2001).
As watershed nitrogen loading increases, the capacity of streams to effectively retain and
transform nitrogen inputs will be overwhelmed and inorganic nitrogen will be transported much
farther, which can lead to eutrophication in streams, rivers, lakes, and estuaries (Peterson et al.,
2001).
Anthropogenic nitrogen loading in rivers is one of the main causes of eutrophication of
surface waters and seasonal zones of hypoxia in estuarine waters especially in the Gulf of the
Mississippi River (Vitousek et al., 1997; Dagg and Breed, 2003; Boesch, 2004; Booth and
Campbell, 2007; Donner, 2007). Freshwater nitrogen appears to be the driver for development
of algal blooms that start the zone of hypoxia every summer off the mouth of the Mississippi
River in the northern Gulf of Mexico. Many marine animals in this ecosystem are stressed or
killed by these hypoxic conditions. Increased agricultural-associated nitrogen loading from
surface runoff in the Mississippi River basin drains into the Mississippi River which leads to
hypoxia problems in the Gulf (Booth and Campbell, 2007; Dodds, 2006; Rayl, 2000; Ferber,
3
2004). The Mississippi River is one of the 10 largest rivers in the world, and its drainage basin
contains greater than 40% of the continental U.S., an area of 3.34 X 106 km2 (Dagg and Breed,
2003). The abundant use of nitrogen fertilizer on Midwest U.S. croplands contributes to nitrogen
loading in the Mississippi River basin, and the majority of grains grown in the Midwest are used
as animal feed (Donner, 2007).
Nitrogen inputs into the Gulf of the Mississippi River have increased dramatically in the
past 50 years which have altered coastal ecosystems (Dagg and Breed, 2003). In modeling
nitrogen inputs to the Mississippi River Basin, fertilizer runoff was found to account for 59% of
nitrogen loading, while 17% was from atmospheric nitrate deposition, 13 % was from animal
waste, and 11% was from municipal waste (Booth and Campbell, 2007). Over five years from
2001 to 2005, the average size of the zone of hypoxia off the Gulf of the Mississippi River was
15,000 km2 (Booth and Campbell, 2007).
1.2. Purpose of the Study
Critical gaps exist about our knowledge of what happens to nitrogen below the root zone
and above the water table in the vadose zone (Kosugi and Katsuyama, 2004). To date there have
not been many studies in the vadose zone especially below the root zone to determine exactly
what nitrogen transformations take place and in what quantities. Reasons given for a lack of
knowledge about nitrogen transformations in the vadose zone are the costs associated with
accurately obtaining data (Buczko et al., 2010) and lack of appropriate instrumentation (Barzegar
et al., 2004). Many of the past nitrogen studies have focused on measuring nitrate in the
saturated zone (Bottcher et al., 1990; Wilson et al., 1990; Hong et al., 2007).
The purpose of this study was to measure soil water potential, to collect soil water
samples for geochemical analyses of nitrate, ammonia, urea, total nitrogen as well as dissolved
4
organic carbon and ferric and ferrous iron at different depths in the vadose zone, and to present
trends and interpretation of these trends in the geochemical data with depth in the vadose zone.
Groundwater and precipitation samples were also examined geochemically for comparisons to
soil water samples in order to provide background concentrations from groundwater sample
averages and natural geochemical additions to the surface from precipitation. The regional water
table surrounding the study site was measured by checking the depths to water levels from a
group of eleven piezometers over several months. Time domain reflectometry (TDR) data was
collected over several months to measure soil moisture data of the vadose zone that were made
into volumetric water profiles.
1.3. Limitations of the Study
Vadose zone studies can be difficult because of the time and cost to collect and analyze
adequate samples, so decisions need to be made about where to sample and in what quantities in
order to understand nitrogen transformations taking place. Sampling in the root zone is
necessary because this is a dynamic zone where microbiological activity is very high and key
transformations occur, usually in the uppermost 0.5 m of soil. Below the root zone, the nitrogen
transformation processes are poorly understood, so sampling in the intermediate vadose zone
should be made. Sampling just above the water table is also necessary to understand what
nitrogen species are entering the unconfined aquifer below the water table and in what
concentrations, and with seasonal variations in a water table, a depth should be chosen above the
shallowest known seasonally fluctuating water table elevation.
Some other limitations of working in the vadose zone are pore-size of soil surrounding
the lysimeters and sampling frequency. The pore-size of the soil which can be sampled using
lysimeters is limited due to suction applied, which is limited by the air entry value of the
5
lysimeters. Most suction lysimeters cannot exceed one bar of suction. Soil water removed from
suction lysimeters typically comes from smaller pores, unless the porous cup of the lysimeters
are in direct contact with macropores. Too much sampling using suction lysimeters can
influence flow paths to the porous cups of the lysimeters. Excessive sampling might not produce
soil water characteristic of the depth of the sampling (Grossmann and Udluft, 1991). However,
inadequate sampling might not identify key nitrogen transformations which are occurring in the
vadose zone. The sampling campaign aimed to sample frequently: usually every two weeks
especially a few days after rain once the soil had wetted up again. Often low soil moisture
prevented soil water samples from being collected every two weeks resulting in sampling at
variable intervals throughout the study.
Drought can represent a significant limitation for sampling. Although nitrogen
transformations are still occurring in drought conditions, without enough precipitation infiltrating
the system, soil water cannot be collected to understand the geochemical processes taking place
in the vadose zone. As drought continues over several months, the matric potential in the vadose
zone increases which creates a greater gradient essentially pulling infiltrating precipitation
through the soil relatively rapidly. Often in a drought, too little soil water infiltrates to sampling
depths so that adequate geochemical analyses cannot be made.
6
CHAPTER 2
LITERATURE REVIEW
2.1. Agricultural Nitrogen Fertilizer Application
Nitrogen fertilizer can help produce economically profitable crop yields which can in turn
help in attaining a sufficient agricultural program. However, nitrogen management practices
which optimize crop yields need to take into account the leaching of possibly harmful nitrogen
species into groundwater. To insure that maximum crop yields are attained in order to maximize
profit, farmers often use excessive amounts. This approach can result in excessive nitrogen
loading so that nitrogen leaching and denitrification do not deplete all the plant available nitratenitrogen (Follett and Walker, 1989). Another agricultural approach is to attempt to lower the
rate and duration of nitrogen leaching and denitrification. This can be achieved by using
nitrification inhibitors to slow and lessen the effects of denitrification; USDA has begun using
them with applied fertilizers. Further, farmers should consider realistic yield goals that could
include more conservative fertilizer applications and different types of fertilizers. Careful
management must be used to achieve the balance between too much and too little nitrogen. A
significant portion of nitrogen that is not utilized or transported out of the system will remain in
the root zone in either immobilized or inorganic forms. The immobilized/inorganic forms of
nitrogen can be used by subsequent crops, and as long as excess water does not leach
immobilized nitrogen below the rooting depth it poses little hazard to the environment.
Agricultural management practices should make the most efficient use of nitrogen resources for
crop production (Follett and Walker, 1989).
7
2.2. Non-point Source Nitrogen Pollution from Agriculture
In general, agricultural land use is one of the most important sources of non-point source
pollution (Kronvang et al., 1995; Keeney, 1989). Sapek (2005) states that nitrate from
agriculture is the main source of groundwater pollution. In terms of groundwater pollution by
nitrate, many polluters, each contributing an unknown amount of nitrate that could have come
from several sources such as manure, nitrogen fertilizer, soil organic matter, and/or crop residue,
may eventually affect several different people in varying ways at different times (Follett and
Walker, 1989).
Keeney (1989) states that nitrate sources can be generalized as high-density animal
operations where feed is transported into a watershed and manure must be spread at rates in
excess of crop nutrient requirements, and row-crop agriculture, which uses manure fertilizer as a
source of N to supplement crop needs. Hallberg (1989) states that background nitrate
concentrations are typically < 2 mg L-1 NO3-N in shallow ground water while agricultural areas
often exhibit > 10 mg L-1 NO3-N concentrations seasonally.
Several primary factors controlling nitrate pollution of groundwater are the amount of
nitrogen available, the amount of infiltrating or percolating water, the hydraulic conductivity of
the material, depth to the water table, and the potential for nitrate reduction and/or denitrification
(Hallberg, 1989). Some common methods to help control non-point source pollution include
reducing fertilizer applications, proper timing of application of fertilizers, establishing erosion
control strategies like riparian zones and silt fencing around exposed soil, keeping livestock out
of surface water bodies and providing water troughs instead (Dodds, 2002). However, controls
on excess nitrogen application can have limited short-term benefits because of excessive
nutrients stored in watersheds from years of nutrient pollution (Bennett et al., 1999).
8
According to the USEPA nitrate/nitrite factsheet, Georgia and California were the top
two states in terms of nitrate and nitrite releases to water and land between 1991 and 1993
(USEPA, 2008). Major industries of the United States were found to have released
approximately 50.2 million pounds of nitrogenous fertilizer between 1991 and 1993, which only
includes industries that released at least 10,000 pounds or more. From 1991 to 1993, Georgia
was found to have released 12.1 million pounds of nitrate and nitrite to water, and 12.0 million
pounds of nitrate and nitrite to land (USEPA, 2008).
The International Fertilizer Industry Association estimated in 2006 that worldwide
nitrogen fertilizers were 9.09 X 107 Mg yr-1 (Abit et al., 2008). As of 1993, industrial fixation of
nitrogen for use in fertilizers is estimated to be about 80 Tg yr-1 by the Food and Agriculture
Organization (FAO) of the U.N. (Vitousek, 1997). The FAO estimated world fertilizer
production as of June, 2006, to be 155,057 X 1,000 tonnes of N, P2O5, and K2O (FAO, 2008).
Since the 1970s an increase of fertilizer use in Asia has been observed, although Western Europe
currently uses the largest unit-area amount of fertilizer on cropland at a rate of 105 kg ha-1
(Burkart and Stoner, 2007).
According to studies completed in 1998 by the National Water-Quality Assessment
(NAWQA) program, nitrate and pesticides were the most frequently detected pollutants in
shallow groundwater (less than 30 m below the land surface). Applications of fertilizers,
manure, and pesticides have degraded the water quality of streams and shallow ground water in
agricultural areas. Agricultural practices of fertilizer, manure, and pesticide application have
resulted in some of the highest concentrations of nitrogen measured in NAWQA studies. Nitrate
9
concentrations exceeding the USEPA maximum contaminant level (MCL) of 10 mg L-1 have
been found in 15 % of shallow ground water sampled beneath agricultural and urban land
(USGS, 1999).
Several sources of nitrate in groundwater have been attributed to natural geologic
deposits which have never been fertilized. One study by Boyce et al. (1976) found nitrate in
groundwater that had leached through never-fertilized Pleistocene loess in semiarid southwestern
and western central Nebraska (Keeney, 1989). High levels of nitrate have been found in
groundwater that leached through alluvium beneath the San Joaquin Valley, California (Keeney,
1989).
2.3. Nitrogen Fertilizer Application Thresholds
Fertilizer applied above certain threshold values can begin to accumulate nitrogen in the
soil, which is a reason for careful fertilizer application management. Different studies point to
different thresholds which could mean thresholds are often site-specific or soil-specific. One
study by Bergstrom and Brink (1986) spanning 10 years on an arable clay soil in Sweden found
moderate leaching of nitrate up to an application rate of 100 kg N ha-1 yr-1. However, leaching
increased rapidly after this rate threshold was exceeded. In years when this rate threshold was
exceeded, build-up of inorganic nitrogen in the soil increased the potential for future leaching
(Bergstrom and Brink, 1986).
Another study by Kolenbrander (1981) found a critical threshold range of nitrogen
fertilizer applied between 100 – 200 kg N ha-1 yr-1. Kolenbrander found this threshold range to
be consistent over many sites with different soil textures but with the same drainage of 300 mm
yr-1. Once again, when this threshold range was exceeded for nitrogen fertilizer application rate,
then nitrate leaching losses increased rapidly (Kolenbrander, 1981).
10
Two studies found that nitrogen fertilizer application rates could be reduced by half from
400 kg N ha-1 yr-1 to 200 kg N ha-1 yr-1, which reduced the nitrate leached from 100 to 27%, but
this reduced live weight of grazing cattle by 10% (Tyson et al., 1992; Scholefield et al., 1993).
These studies were on a clay loam field in Devon, UK, over a 7 year period. Management
practices required only a 10% increase in field size, from 1 ha to 1.1 ha, for the lower rate of
fertilization to reduce nitrate leaching losses by 73%. Therefore, with better management
practices, especially less nitrogen fertilizer application, and economic sacrifices possibly
reimbursed through government aid, nitrate leaching can be reduced greatly on the pasture level
(Cuttle and Scholefield, 1995).
2.4. Manure, Urea, and Nitrogen
In addition to nitrogen-based fertilizers, animal manure is a major contributing source of
nitrate to soil (Griffin and Honeycutt, 2000; Eghball, 2000; Abit et al., 2008). Manure is a point
source of nitrogen groundwater pollution when stored or cattle are confined, and manure is a
non-point source when applied to fields (Burkart and Stoner, 2007). Compared with inorganic
fertilizer nitrogen, organic wastes are preferred because nitrogen from organic wastes stays in the
mineralization/immobilization phase longer, which makes the nitrogen more slowly available
and not as susceptible to rapid loss by leaching (Keeney, 1989). Organic nitrogen in manure can
mineralize much faster than nitrogen compounds in soil organic matter (Sapek, 2005). Organic
waste application is usually added in the spring and fall when neither crops nor soil
microorganisms are active, and these seasons are also prone to substantial rainfall, which can
leach nitrate into the groundwater (Burkart and Stoner, 2007).
In North America, Asia, Western and Eastern Europe, approximately twice the amount of
nitrogen comes from inorganic fertilizer than from manure. However, in Latin America, Africa,
11
and Oceania, twice to three times the amount of nitrogen comes from manure compared with
organic fertilizer, and in the former USSR, the ratio of manure to inorganic fertilizer is about 1.7
(Burkart and Stoner, 2007).
Negative aspects of manure and urea application include that concentrations of nitrogen
are usually low in organic wastes. Also, transportation of organic waste is expensive.
Furthermore, composition and quality of organic wastes are variable. In addition, mineralization
requires some time after application before nitrogen is plant available so timing can be a problem
for rapid growing crops like corn which grows as fast as nutrients are supplied. Organic wastes
can be high in ammonia content which can be volatilized if the organic wastes are not
immediately incorporated into the soil. Organic wastes can sometimes contain toxic elements
such as heavy metals depending on the diet of the animals that produced the waste (Keeney,
1989).
Intensively managed forage and grazed grasslands can be a significant source to nitrate in
groundwater. Grasslands have annual above-ground biomass which leaves nitrate in the soil in a
cyclic fashion at times of the year when uptake by plants is low, such as autumn and spring.
This nitrate-nitrogen can be leached to the groundwater becoming a source of nitrate
contamination. Animal wastes, especially urine, are found in concentrated patches in grazed
pastures, which can lead to inefficiency of waste N use (Keeney, 1989).
One study by Adams et al. (1994) fertilized fescue pastures in Fayetteville, Arkansas,
with varying amounts of poultry litter and manure treatments to measure NO3--N concentration
in vadose water as a function of depth and time. The purpose of the study was to ascertain the
effect of application rate of poultry litter or manure on nitrate leaching in the vadose zone.
Twelve plots were made on a uniform slope (5%) Captina silt loam soil (fine-silty, siliceous,
12
mesic, Typic Fragiudult). Suction lysimeters at 60 cm and 120 cm depth and pan lysimeters at
60 cm depth were used to collect vadose water samples about every two weeks. Tensiometers
were installed at 45o angles at tip depths of 30, 60, 90, and 120 cm to monitor soil water tension
and to determine hydraulic gradients. Application rates of poultry manure (PM) and poultry
litter (PL) varied per plot from 0 Mg N ha-1 yr-1 (control) to 10 Mg N ha-1 yr-1 (PM10 and PL10)
to 20 Mg N ha-1 yr-1 (PM20 and PL20) in the first year with an additional 5 Mg N ha-1 yr-1 of
additional poultry litter applied to PL10 and PM20 in the following year (June, 1992). About 30
days after the first application, all treated plots at 60 cm depth had NO3--N concentrations greater
than 10 mg L-1 (Figure 2.1). The nitrate concentrations at 60 cm depth peaked 70 to 90 days
after the first application between 41 mg L-1 NO3--N for PM20 to 54 mg L-1 NO3--N for PL20,
and concentrations at 60 cm depth peaked at 13 mg L-1 NO3--N for PL10 about 150 days after
application.
13
Figure 2.1. Soil Water Nitrate-N Concentrations at (a) 60 and (b) 120 cm Depth vs. Time
(Modified from Adams et al., 1994)
Dry conditions prevailed for a couple of months in April and May, and afterwards nitrate
concentrations had dropped to background (control) levels (Figure 2.1). Soil water nitrate mass
loads peaked at 60 cm depth to values greater than 20 Kg ha-1 in pan lysimeter data as well
(Figure 2.2). In spite of the addition of 5 Mg N ha-1 for the PL10 plot in the second year, nitrate
concentrations went to background (control) levels 300 days after the first application and
remained there for the rest of the study (Figure 2.1). Nitrate concentrations for PL20 and PM20
dropped sharply in May and declined through the summer months below 1 mg L-1 NO3--N even
though PM20 received an additional application of 5 Mg N ha-1 in June. The authors attributed
the drop in nitrate levels to plant uptake as the fescue cover crop was growing rapidly during the
14
summer, and NO3--N concentrations increased with higher litter and manure applications rates
regardless of type of poultry waste as long as equal amounts of N were applied.
Figure 2.2. Soil Water Nitrate Mass Loads (Pan Lysimeters at 60 cm Depth) vs. Time
(Modified from Adams et al., 1994)
During the winter months, all poultry waste treatments caused increased nitrate
concentrations in the vadose zone, and matric potential measurements indicated downward water
movement in the winter. This implied that nitrate leaching was maximized during the winter
months, and nitrate could be minimized by applying poultry waste only during the spring and
early summer when microbes are actively utilizing N and less water is percolating down through
the vadose zone (Adams et al., 1994).
15
2.5. The Agricultural Nitrogen Cycle
Figure 2.3. The Nitrogen Cycle for Agriculture (Modified from Bellows, 2001)
The sources of nitrogen are the starting point for the nitrogen cycle in agriculture (Figure
2.3). These sources include elemental nitrogen gas (N2), which comprises about 78% of the
Earth’s atmosphere, through precipitation, lightning, and nitrogen fixing legumes as well as
fertilizers: inorganic and organic (manure and urea). Decay of organic matter also provides
energy and electrons needed for nitrogen transformations to occur. Decay of organic matter
(OM) that contains organic carbon, organic nitrogen, and phosphorous compounds as well as
trace elements occurs by the following chemical reaction:
Decay of OM:
Organic Matter + O2 = CO2 + NO3- + HPO42- + H2O + H+
16
Once nitrogen enters the soil, mineralization/ immobilization can convert organic nitrogen to
nitrate. Simultaneously, organic C is mineralized to CO2, while some organic C goes to form
more soil microbes. Along with carbon being released as carbon dioxide, organically combined
nitrogen is released as nitrate. Since free electrons cannot accumulate, corresponding chemical
constituents accept the electrons and these electron acceptors are oxidizing agents or chemical
constituents capable of being reduced. When oxygen is present, it accepts free electrons in a
process called aerobic metabolism:
O2 + 4H+ + 4e- = 2H2O
However, when oxygen is not present, other electron accepting chemical constituents take up the
free electrons. The chemical constituent that will take up free electrons is determined by the
redox potential, and nitrate has the next highest redox potential after oxygen. Nitrate can take up
free electrons by two reactions:
(1) Denitrification:
2NO3- + 12H+ + 10e- = N2 + 6H2O
(2) Nitrate to Ammonium also known as ammonification (2 step reaction):
2Corganic + NO3- + H2O + H+ = 2CO2 + NH3
NH3 + H2O = NH4+ + OHAmmonia released by microbial activity reacts with water to form the ammonium ion and causes
a net rise in pH. In soils with pH less than 7.5, ambient hydrogen ions convert ammonia to
ammonium. Ammonification or mobilization is the process where bacteria convert organic
matter (made up of organic nitrogen and carbon) into ammonium (Drever, 2002).
17
In a complex series of reactions, bacteria reduced nitrate by using it as the terminal
electron acceptor to oxidize organic carbon to CO2. When elemental nitrogen is the end product,
the process is called denitrification or dissimilatory nitrate reduction, and the reaction is:
5Corganic + 4NO3- + 4H+ = 2N2 + 5CO2 + 2H2O
Many bacteria reduce nitrate only as far as nitrite:
Nitrate to Nitrite:
Corganic + 2NO3- = CO2 + 2NO2-
Volatilization is when ammonium reacts with hydroxide ions resulting in ammonia exiting the
soil system as gas to the atmosphere by the following reaction:
Volatilization:
NH4+ + OH- = NH3 + H2O
(Drever, 2002)
When nitrogen is mineralized/immobilized in the soil, carbon is being consumed so the
net result is more mineral nitrogen and less C available for heterotrophic soil microbe growth.
Some of the produced ammonium will be taken up by plants, while some of the ammonium
(NH4+) will be nitrified, converted to nitrite (NO2-), and possibly further nitrified to nitrate
(NO3-). Then some of the nitrate or nitrite will be subsequently denitrified, and a small portion
of ammonium will be incorporated into recalcitrant soil organic matter (known as
immobilization) that is very slowly mineralized. Nitrate also enters the nitrogen cycle (Figure
2.3) by means of the microbial immobilization step, but heterotrophs strongly prefer ammonium,
whereas nitrate is more strongly assimilated by some higher plants (Keeney, 1989).
Burkart and Stoner (2007) ranked types of aquifers which are susceptible to nitrate
contamination: unconfined aquifers associated with agriculture, carbonate aquifers, and alluvial
aquifers. Shallow unconfined aquifers associated with agricultural systems were most
susceptible to nitrate contamination (Burkart and Stoner, 2007)..
18
Nitrification is the microbial-induced oxidation of ammonium to nitrite and further to
nitrate. Excluding some atmospheric reactions, nitrification is responsible for the sole natural
source of nitrate to the biosphere. Nitrification transforms the relatively immobile ammonium
ion to a very mobile nitrate ion, which can be leached or denitrified. Nitrification is nearly
exclusively carried out by the Gram-negative (will not retain the violet dye when stained by the
Gram method), chemosynthetic, autotrophic bacteria of the family Nitrobacteriaceae. Five
genera are recognized, but culture studies use Nitrosomonas. Nitrobacter is the dominant nitrite
oxidizer (Keeney, 1989).
Denitrification is a biological-pathway whereby nitrogen is returned to the atmosphere
as gaseous N (N2 or N2O). Denitrification is often considered a nitrogen loss and a reason that
fertilizer N is inefficient in agriculture. Biological denitrification produces N2O gas, a
photochemical oxidant, which destroys ozone (O3, a photochemical oxidant) in the stratosphere.
Denitrifying bacteria are capable of normal respiratory growth when enough oxygen is present,
but when the environment has little to no oxygen, denitrifying bacteria use nitrate, nitrite, or
nitrous oxide as terminal electron acceptors. Under a highly reduced anaerobic environment
with excess organic C, nitrate can be reduced to ammonium. At least 14 genera of denitrifying
bacteria are known and present in most soil and aquatic environments. Significant denitrification
is not known to take place in most vadose zones or aquifers due to lack of sufficient organic C
and denitrifier microbes (Keeney, 1989). Denitrification is the dominant process that can
attenuate nitrate contamination in saturated materials beneath agricultural systems (Burkart and
Stoner, 2007). Scientists are beginning to understand that the elimination of wetlands and
riparian areas has also removed important natural nitrogen traps where much of the entering
nitrate is denitrified (Vitousek et al., 1997).
19
Two processes transform elemental nitrogen (N2 gas) to biologically available forms:
lightning and biological nitrogen fixation. Biological nitrogen fixation is carried out by
microorganisms. Many microorganisms are in symbiotic relationships with algae and higher
plants, especially legumes such as beans, alfalfa, peas, clover, lentils, and peanuts (Vitousek et
al., 1997). The highest numbers of bacterial abundance are found in and around plant roots
compared with the bulk, plant-free soil, which is due to excretion of assimilates into the plant
apoplasm. Even though bacterial abundances are much higher in roots, the weight of the bacteria
account for less than 0.1 % of the root weight, and in legumes with higher bacterial abundances
the weight percentage of the root comprised of bacteria is only slightly higher (Bothe and Drake,
2007). Macropores and preferential flow paths are sites of relatively high nutrient availability
which contributes to the large abundances and heterogeneity of microbes and biologically
mediated processes in the vadose zone (Holden and Fierer, 2005).
Biological denitrification occurs when one or both of the ionic nitrogen oxides (NO3-,
NO2-) is reduced to the gaseous oxides (NO and N2O) which can be further reduced to N2 gas.
Biological denitrification is carried out by heterotrophic bacteria and fungi which use N oxides
as terminal electron acceptors and organic carbon as electron donors (McNeill and Unkovich,
2007). Denitrification is aided by high availability of organic C and NO3 --N, a low rate of
oxygen diffusion which can come from high soil moisture content or compaction, or an increase
in soil pH or temperature (McNeill and Unkovich, 2007). Soils at low elevations in watersheds
aid in denitrification in many ways such as high moisture contents, high C contents, and a low
rate of oxygen diffusion (McNeill and Unkovich, 2007; Stevenson and Cole, 1999).
Many different microorganisms are responsible for each of the transformation-pathways
that convert one form of nitrogen into another in the nitrogen cycle. Denitrification is a
20
microbially mediated process which is carried out above 10oC and in the presence of readily
available carbon or other electron donors (Burkart and Stoner, 2007). Denitrification, which
reduces nitrate and nitrite to nitrogen oxides, can be carried out by fungi and bacteria, and some
of the bacteria that are capable of denitrification include: alcaligenes, agrobacterium,
azospirillum, bacillus, flavobacterium, halobacterium, hyphomicrobium, paracoccus,
propionibacterium, pseudomonas, rhodopseudomonas, and thiobacillus. Blue-green algae are
capable of fixing N2 from the atmosphere, and plant-algal associations like gunnera, azolla, and
lichens can also fix elemental nitrogen. Examples of heterotrophic nitrifying bacteria include:
arthrobacter sp., azotobacter sp., pseudomonas fluorescens, klebsiella aerogenes, bacillus
megaterium, and proteus sp. Examples of heterotrophic nitrifying actinomycetes, actinobacteria
which play a vital role in decomposition of organic matter, include: streptomyces, nocardia, and
penicillium sp. Also, examples of fungi which are heterotrophic nitrifying microorganisms are:
aspergillus flavus and neurospora crassa (Stevenson and Cole, 1999).
Denitrification rates do not decline continuously with depth, but instead vary
considerably from microsite to microsite (Paramasivam et al., 1999; Holden and Fierer, 2005).
One study by Luo et al. (1998) found denitrification rates decrease 10 to 100-fold from 0 to
between 10 and 30 cm depth, while another study by Paramasivam et al. (1999) reported 50 to
100% reduction in denitrification rate between the surface and 90 cm (Holden and Fierer, 2005).
Extreme variability exists in nitrogen concentrations of similar types of manure, and
farmers are often uncertain as to the amount of nitrogen from manure applications that will
become available for plant uptake. About 40 to 60% of the N in manure is present as
ammoniacal-N or as urea and uric acid-N which can readily hydrolyze to ammoniacal-N, and
when left exposed to the atmosphere much of the ammoniacal-N can be lost as volatized
21
ammonia, which can happen as quickly as a day-and-a-half. Furthermore, greater uncertainty
exists in the fate of applied N in manure compared to fertilizer N because manure C is easily
decomposed and provides energy for denitrifying bacteria, which results in increased
denitrification in soils with applied manure (Schepers and Fox, 1989).
Solutes in precipitation often depend on the solute content of sea water, but the influence
of sea water has a lesser effect the further inland clouds move. The nitrogen species in
precipitation, NO3-, NH4+, and nitric acid (HNO3), come from gaseous nitrogen releases from
terrestrial vegetation, agriculture, automobile exhaust, and industrial pollution. Ammonium in
precipitation returns to the soil surface from volatilization of animal wastes. Combustion of
fossil fuels oxidizes atmospheric nitrogen into various nitrogen oxides that return to the soil as
nitric acid, an important component of acid rain. The composition of rain water at one location
may vary greatly with time. For example, the first precipitation to fall in a rainstorm may
contain most of the soluble material present in the atmosphere, while precipitation at the end of
the storm is relatively dilute (Drever, 2002).
Another way atmospheric nitrogen is delivered to the Earth’s surface is by occult
deposition, which includes dry deposition and deposition from fog and mist. Dry deposition
occurs when solutes are transferred from the atmosphere to surfaces especially surfaces on
vegetation or moist foliage. The amount of dry deposition depends on the vegetation present.
Conifers appear the most effective at dry deposition, whereas deciduous trees are second best,
and grasses are least effective. Nitrogen species (nitrate and ammonium) and sulfates are solutes
most affected by dry deposition (Drever, 2002).
22
2.6. Vadose Zone Water and Nitrogen Movement
One textbook definition of the term vadose zone is “the geologic media between the land
surface and the regional water table” (Stephens, 1996). The term vadose comes from the Latin
word vadosus, which means shallow. Vadose zone is a more encompassing term than
unsaturated zone or zone of aeration because the vadose zone includes the soil zone, the
intermediate vadose zone and the capillary fringe, which is an area encompassing the seasonally
fluctuating water table (Looney and Falta, 2000).
Two processes control water movement in the vadose zone. The first is gravity which
moves water down, and the second is a capillary process which spreads water out in all
directions, stores, and releases water. In most cases, capillary processes dominate in fine-grained
sediments such as clay and silt, and usually gravity is dominant in course-grained sediment and
large fractures. For most vadose zone sites, the boundary conditions are very dynamic and the
water content constantly changes with time. Modeling chemicals in the vadose zone can be very
complex because the chemicals interact with the soil and other constituents in the soil solution in
a dynamic way. Events and conditions in the vadose zone greatly influence the behavior of
contaminated water being discharged into an aquifer (Looney and Falta, 2000).
Many studies have proposed reasons for faster than predicted water potential and tracer
travel times through unsaturated media, such as preferential flow, bypass flow, macropore flow,
fracture flow, boundary layer flow, fractured-quartz-vein flow, mobile zone flow, finger flow,
media heterogeneities, kinematic flow, etc. These studies have aided in the collective
understanding of solute transport through unsaturated media. In an effort to obtain hydraulic and
tracer properties through undisturbed saprolite cores, rapid pressure waves were found to
propagate through the saprolite cores from short-duration irrigations, and wave velocity
23
predictions using Darcian, tracer, and kinematic models significantly underestimated observed
travel times (Rasmussen et al., 2000). In saturated flow, pressure waves propagate due to the
compressibility of the fluid, but in unsaturated flow, pressure waves move through unsaturated
media due to small changes in fluid saturation within soil pores. Pressure waves were examined
using four parametric models: Brooks-Corey, van Genuchten-Mualem, Broadbridge-White, and
the Galileo Number. Predicted pressure wave travel times were between two and fifteen times
the tracer velocity through the saprolite cores. Based on mathematical models, pressure wave
velocities should decrease with increasing depth, and at some depth, pressure wave velocities
should agree with kinematic theory. The experimental results showed that large hydraulic
diffusivities could cause rapid pressure wave propagation in homogeneous, unsaturated media
(Rasmussen et al., 2000).
Nitrate-nitrogen is a conservative ion when moving through clay-rich soils because
nitrate does not adsorb to clay surfaces. Nitrate moves with precipitation and/or irrigation water
through the soil, and due to variations in pore sizes, spatial distribution of pores, and pore
continuity, infiltrating water irregularly moves down through the soil profile. This irregular,
infiltrating precipitation/irrigation water spreads out the nitrate-N front between the preexisting
soil solution and the displacing water by hydrodynamic dispersion. Also, differences in nitrate
concentrations in the mixing soil water drive diffusive dispersion. Under intensive rainfall,
water may bypass traditional pore water channels for macropore such as structural cracks, dead
root channels, worm channels, or other macropore pathways (Vinten and Smith, 1993).
Nitrogen movement with percolating water through the unsaturated zone can be very
slow and the time required for modest inputs of nitrate to reach the groundwater reservoir may be
24
many years. Because of the slow rate of movement, contamination may persist for decades to
centuries even if input sources of nitrate decrease or are eliminated (Follet and Walker, 1989).
Topography can influence nitrogen species transformations, organic carbon
accumulation, and soil moisture content in a watershed. Significant temperature and moisture
gradients can exist between the tops and bottoms of sloped areas. Soil moisture is often
dependent on differential rates of runoff, evaporation, and transpiration. Also, soils in
depressions have a cooler and more humid microclimate which leads to higher accumulation of
carbon contents compared to knolls where the climate is drier and warmer. Furthermore,
continuously moist and poorly drained soils at the lowest point in a watershed typically have
localized areas that contain organic rich soils, and these conditions aid in biological
denitrification (Stevenson and Cole, 1999).
Anions are commonly assumed to enter the soil and move downward through the vadose
zone with infiltrating water, and then move horizontally in the groundwater. However, recent
laboratory and field studies show that water movement and transport of pollutants, especially
horizontally, can occur in the capillary fringe. Capillary fringes can range from 0 to 1 meter in
height above the water table. Nitrate was compared with another tracer, bromide, which is a
conservative, non-reactive ion not taken up preferentially by plants or used by soil microorganisms, and both were injected into the soil in the unsaturated zone of a drained bay. Nitrate
was found to persist more shallowly than bromide over a study period of several months and to
readily move in the capillary fringe. Nitrate was also found to persist longer in the capillary
fringe than below the water table allowing a greater fraction of it to be transported to greater
horizontal distances in the capillary fringe than in the shallow groundwater (Abit et al., 2008).
25
One study by Bobier et al. (1993) measured nitrate movement in a fertilized, finetextured vadose zone in southeastern Nebraska. Fertilizer was applied at rates of 336 and 448 kg
N ha-1 to selected plots annually from 1971 till 1986. Soil cores were examined first in 1985 and
again in 1990 in the selected fertilized plots, and from these cores, soil extractions for nitrate
were analyzed from 0.3 m intervals. Elevated nitrate-N beneath fertilized plots in 1985, were
statistically identified in 1990, the elevated nitrate zone moved an average of 3.81 m over 5
years. The average rate of movement of the nitrate was 0.76 m yr-1. This rate is similar to a rate
of 0.74 m yr-1 proposed by Alberts and Spomer (1985) in the loess of western Iowa, and this rate
is slightly higher than a rate of 0.62 to 0.69 m yr-1 proposed by MacGregor et al. (1974) in a
Minnesota clay loam (Bobier et al., 1993).
A study by Gehl et al. (2005) characterized nitrate movement through the vadose zone of
a sandy soil to the water table due to the health concerns once nitrate-N exceeds the maximum
contaminant level of 10 mg L-1 set by the EPA. This study used matric potentials obtained from
tensiometers to evaluate unsaturated flow, and nitrogen species concentrations obtained from soil
water samples obtained by suction lysimeters to construct a depth profile of nitrogen species in
the vadose zone. The results showed that the mass of nitrogen leached below the root zone
equaled the product of the mean concentration of the nitrogen leachate multiplied by the volume
of drainage water for a given period of time (Gehl et al., 2005).
2.7. Wetting and Drying Cycles
When compared to field-moist soils, wetting and drying cycles have been shown to cause
an increase in mineral nitrogen concentration in soil water which moves through the system.
Also, each successive cycle of wetting and drying brings about a smaller flush (Stevenson and
Cole, 1999). Nitrate loading is usually highest in the winter and spring because high nitrogen
26
concentrations are a result of soil water recharge and the increase in nitrogen mineralization that
occurs when soil drying is followed by rewetting (Heathwaite, 1993).
The flush in nitrogen in cycles of wetting and drying is believed to result from several
causes. The death of microorganisms from drying releases easily degradable nitrogen
compounds. Also, drying causes transformation of organic nitrogen to more soluble compounds,
which are used by microorganisms with release of mineral nitrogen. Furthermore, wetting and
drying leads to the dissolution of water-stable aggregates which makes new surfaces and
substrates available for microbial utilization (Stevenson and Cole, 1999).
Cabrera (1993) found that the flush of nitrogen through soils followed a model requiring
two nitrogen pools: one requiring zero-order kinetics and one requiring first-order kinetics. In
the study, soils were collected from the upper 10 cm from 3 different soils from 5 different
management practices, and results indicated that in the flush some nitrogen mineralizes quickly
while other nitrogen mineralizes more slowly (Cabrera, 1993).
One study by Mikha et al., (2005) compared soil cores that were constantly wet versus
cores that were subjected to wet-dry cycles and found that mineralization of C decreased with
time when drying and wetting cycles dominated the meteorological patterns. The study found
that soils that experience wetting-drying cycles had significantly less net N mineralization
compared to soils that were constantly wet. For each drying-rewetting period within 24 hours of
rewetting, nitrogen mineralization decreased. Repeated wetting-drying cycles reduced
cumulative N mineralization when compared to soils that were constantly wet, which has not
been seen in other studies like Cabrera (1993) where nitrogen mineralization increased after
rewetting occurred in a dry soil core. The flush of C and N upon rewetting was concluded to be
caused by microbial influence (Mikha et al., 2005).
27
2.8. Darcy’s Law and the Darcy-Buckingham Equation
In mid-eighteenth century France, an engineer named Henry Darcy made the first
systematic study of water movement through a porous medium in a pipe. Darcy found that the
rate of water flow through the sand to be proportional to the difference of the height of two ends
of the pipe the water flowed through and inversely proportional to the length of the flow path.
He determined the amount of flow is proportional to a coefficient which is dependent on the
nature of the porous medium, and this coefficient came to be known as hydraulic conductivity.
Darcy also found that flow was proportional to the cross-sectional area of the pipe, and these
terms assembled in one equation as Darcy’s law (Fetter, 2001). Darcy’s law in one dimension
(x) is:
Q=
Darcy’s law is used to model saturated flow, whereas the Darcy-Buckingham equation
can be used to describe flow in the vadose zone. Total head is the sum of pressure head ( ) and
elevation head (z). The equation of total head is:
h=z+
The Darcy-Buckingham equation was derived from Darcy’s equation for unsaturated
flow in 1907 by Edgar Buckingham. Buckingham understood the important interactions
between water and soil and the role of these interactions in describing unsaturated water flow.
Buckingham described that unsaturated hydraulic conductivity depends on capillary action. The
Darcy-Buckingham equation in one dimension (z) is:
w
h
28
In this equation, Jw is unsaturated flow and K(h) is unsaturated hydraulic conductivity as a
function of matric pressure head.
2.9. The Piedmont Region
The Southern Piedmont is a region of the southern United States encompassing an area
east of the Appalachians from Virginia to Alabama approximately 16.7 million ha (41 million
acres). Over two centuries of row-crop agriculture has ravaged the soil of the Southern Piedmont
leaving 86% classified as eroded according to the USDA NRCS. Soil moisture content is a
measure of residual moisture in the soil, and the higher soil moisture content is, the more easily
infiltrating precipitation moves through the vadose zone. In the Piedmont in general, the winter
season is a period of high average soil water content, while the summer season had the least soil
water content on average, except when influenced by intense rainstorms (Endale et al., 2006).
The Georgia Piedmont has crystalline igneous granite and metamorphic bedrock, which
generally have very little primary porosity. Secondary porosity exists in the bedrock of the
Georgia Piedmont in fractures and joints which may or may not be interconnected. Fractures in
crystalline rock develop from pressure relief due to erosion of overburden rock, shrinking during
cooling of the rock, regional tectonic stresses that result in compression and tensional forces, and
tectonic movements (Fetter, 2001).
Fetter (2001) discusses the Piedmont along with the Blue Ridge as one ground water
region that “consists of a thick mantle of weathering residuum over fractured crystalline and
metamorphic rock”. The weathered residuum saprolite lies above the metamorphic rocks and
can yield small to moderate amounts of water almost anywhere, with larger-yield wells in valleys
rather than hills and possibly on fracture traces (Fetter, 2001). Saprolite results from in-situ
weathering of parent material which makes up a significant portion of the C horizon in the
29
Southeastern Piedmont (Rasmussen et al., 2000). Radcliffe (2005) states, “The groundwater
system in the Piedmont can be characterized as an unconfined, two-layer aquifer composed of a
zone of saprolite underlain by fractured bedrock.” The saprolite layer is vital to the groundwater
system as a zone of water storage for the deeper fractures (Radcliffe, 2005).
One study by Rose (1992) found water moving through the regolith (including soil,
saprolite, and weathered rock material) in the Piedmont had an estimated residence time of
approximately 25 years. This estimated residence time was calculated from observing the
difference in tritium concentration between groundwater, precipitation from the hydrogen bomb
testing era (1960s), and modern precipitation. Unfortunately, there is no direct correlation
between tritium concentration and age, but reasonable estimates can be made.
This residence
time estimate was based on tritium concentrations of shallow groundwater that was between 28
to 34 TU (tritium units) with a precision of + / - 1 TU or better.
30
CHAPTER 3
MATERIALS AND METHODS
3.1. Experimental Site, Land Use, and Soil
The experimental site was in a 10-ha catchment, named W2, at the USDA-ARS, J. Phil
Campbell Sr. Natural Resource Conservation Center (JPC), Watkinsville, GA, USA, within the
Georgia Piedmont (Figure 3.1). The topography and soils are typical of gradual sloping, primary
catchments throughout the Southern Piedmont region. Some of the longest flow paths in the
catchment range from 390 m to 540 m in length. The highest point at the hilltop groundwater
divide along Well Brook Road is located at 241 m above sea-level to the lowest point at a large
flume measuring the primary spring-fed stream at 222 m above sea-level, and the relief was
about 19 m. Topographic slopes in W2 range from 2 to 10% (Amirtharajah et al., 2002).
31
Figure 3.1. USDA-ARS Watershed 2 Elevation Contour Map (Source: USDA-ARS-JPC)
The experimental site is located near Watkinsville approximately 55 miles East of
Atlanta, in Northeast Georgia. The Northeast Georgia region is known for a warm temperate
climate and ample rainfall, excluding the past two years in which Northeast Georgia experienced
a severe drought. Temperatures in Athens over a 65 year period from 1945 till 2009 were
recorded at Ben Epps Airport in Athens and ranged from a high average temperature of 72.2oF to
a low average temperature of 50.9oF. The high average and low average temperatures in 2007,
2008, and 2009 were, respectively, 75.4oF and 51.1oF72.9oF and 50.0oF, and 71.9oF and 50.7oF
(Hogenboom, 2010). Precipitation was observed and recorded at weather stations at the UGA
Horticultural Farm in Watkinsville across from the experimental site. Average annual
32
precipitation over a 65 year period from 1945 till 2009 was 124.46 cm (49.0 inches). In 2007,
the average annual precipitation was down from the average to 78.74 cm (31.0 inches), and in
2008, the average annual precipitation was only 88.65 cm (34.9 inches). In 2009, the average
annual precipitation was up to 169.16 cm (66.6 inches) (Hoogenboom, 2010).
The historical land use of the catchment was most likely cotton agriculture, typical for
Georgia. Terrace bench remnants can still be seen on an adjacent catchment, and muted terraces
can be seen in the experimental catchment. These terraces were used in historical agriculture as
a means to control soil erosion and to help capture overland flow. Cotton agriculture commonly
erodes landscapes. Cotton requires a large amount of nutrients which were stripped from the soil
and not replenished. In addition, all competition is removed resulting in bare soil and erosion.
The collection of land owned by the USDA-ARS, including the 10-ha catchment and study site,
located North of Hog Mountain Rd., is known as the North Unit. The North Unit is part of a
long-running experiment, spanning several decades, to stop erosion associated with certain types
of farming, e.g. cotton farming. Down-slope of the study site, a wetland naturally formed
downstream of the contact spring. In the 1940s, a dam was built on the spring fed creek that
caused a small pond. Around the 1970s, the dam holding the pond breached, and the pond was
drained. The area below the spring filled in with sediment from tillage in the watershed and
runoff, and once again became a wetland. In 1999, in joint USDA-ARS JPC-Georgia Institute of
Technology research to study water movement and transport of a cryptosporidium surrogate, the
northern edge of the wetland was channelized, which brought the watershed to its current status
(Amirtharajah et al., 2002).
The 10-ha watershed is dominated by Cecil and Pacolet soil series. These soil series are
both classified as fine, kaolinitic, thermic Typic Kanhapludults. The Pacolet soils generally have
33
less depth than the Cecil soils due to erosion of the A horizon, but the soil properties are
otherwise similar (Endale et al., 2006). The Georgia Piedmont Cecil and Pacolet soil series in
Oconee County, GA, have been analyzed in further detail by Perkins (1987). The watershed is a
fescue pasture with some bermuda grass and rye grass, which was over-seeded with cereal rye in
the winter for the grazing cattle.
The watershed is generally used as a rotational pasture with 20 to 200 head of Black
Angus cattle grazing from a few days to several weeks at a time. From 2003 through 2005, total
cattle during any one rotational grazing period varied from 52 to 152 cows. From 2005 through
present, only 24 to 100 cows were grazed in W2 at any one time (personal communication with
Dr. Dinku Endale, 2010). In addition to cows adding nutrients in manure and urea in urine to the
watershed, W2 was fertilized with 92 kg N ha-1 in 2007, not fertilized in 2008, and fertilized with
74 kg N ha-1 in 2009.
The original study area consisted of a three meter radius circle approximately 28.3 m2
portion at the bottom of the 10-ha catchment about 20 m upslope from a spring equipped with a
small flume (Figure 3.2). This study area was instrumented every 3.14 m along the 18.8 mcircumference of the circle with alternating tensiometers and suction-lysimeters to random
depths of 0.5 m, 1.0 m, or 1.5 m with porous ceramic cups in close contact with the soil.
However, these original three lysimeters ceased to function.
The three non-functioning lysimeters were replaced with nine more lysimeters, and all
are currently functioning. Six of these lysimeters were installed within approximately 5 m of the
original circle study area at depths of 0.35, 0.5, 1.25, and 1.75 m. Later, two suction-lysimeters
were installed in the study at 0.35 m and 0.5 m depths. These additional lysimeters had a
diameter of 0.0381 m (Soilmoisture Equipment Corp., Santa Barbara, California). Three more
34
lysimeters with a 0.0381 m-diameter were built from schedule 40 poly vinyl chloride body tubes
with porous ceramic cups (Soilmoisture Equipment Corp.), which were installed at depths of 0.5
m, 1.25 m, and 1.75 m. The 1.25 m and 1.75 m depth lysimeters that were built were located
two meters down-slope of the original circle study area. The constructed lysimeter (0.5 m depth)
was clustered within one meter of the 1.25 m depth and 0.35 m depth Soilmoisture Equipment
lysimeters. In total, nine suction-lysimeters were used in geochemical sampling to ensure good
spatial coverage and adequate geochemical analyses would take place where water movement
was being measured (Figure 3.2).
35
Figure 3.2. Field Site in Watershed 2
36
3.2. Tensiometers and Pressure Transducers
Tensiometers are devices that measure water potential that is the energy status of water in
the subsurface, usually expressed as a matric potential or pressure (Faybishenko, 2000a).
Tensiometers have been in use since the 1920s, but in the past century their basic parts have not
changed: a fine-porous ceramic or metal cup in contact with the soil at depth, connected through
a water-filled tube to a gauge, a manometer, or a pressure transducer. Various designs have been
made over the years, and each type of tensiometer has inherent benefits and limitations
(Faybishenko, 2000b).
Faybishenko (2000b) developed a two-cell, omni-depth tensiometer for use in soil and
rock in the unsaturated and saturated zone (Figure 3.3). Using three of these Faybishenkodesigned tensiometers at 0.5, 1.0, and 1.5 m depths around the 3 m-radius study area, pressures
were monitored every minute, averaged and recorded every 10 minutes continuously for 11
months. At several times during the winter, data collection was suspended due to freezing
temperatures. In the summer, during times of drought, data collection was suspended due to dry
soils. The tensiometer pressures were recorded from two metal tubes that exited the tops of the
two-cell tensiometers (Figure 3.3) using wet-wet 0-15 psig differential pressure transducers and
one wet-wet 0-30 psig differential pressure transducer (Omega Engineering Inc., Stamford,
Connecticut, USA). This two-cell tensiometer consisted of white poly-vinyl chloride pipe with a
diameter of 0.0222 m segmented into two pieces: an upper piece of varying length depending on
the length of the total device representing an upper cell and a lower piece with a fixed length of
0.305 m representing the lower cell (Figure 3.3). The top of the upper cell of each tensiometer is
corked with a rubber stopper with three holes where three metal tubes exit. The three metal
tubes have a 0.0025 m-diameter and are the lower cell monitoring tube which measures the air
37
pressure in air space above the water level in the lower cell, the upper cell monitoring tube that
measures the air pressure in the air space above the water level in the upper cell, and the water
supply tube (Figure 3.3). The white poly-vinyl chloride tensiometers consist of a length of pipe
approximately 0.10 to 0.35 m longer than the length at which soil water sampling occurs for ease
of any maintenance that should be needed at the top of the tensiometers where the pressure
transducers are attached to the exiting metals tubes. The bottom of the lower cell was fixed onto
a ceramic porous cup (Soilmoisture Equipment Corp., Santa Barbara, California, USA). The two
cells of each tensiometer were separated by a piece of plastic with two holes called a connector.
One of the holes in the connector was for a metal tube to monitor the air pressure of the lower
cell, and the other hole was for a metal tube to exchange fluids, water and air, between the two
cells (Faybishenko, 2000b).
38
Figure 3.3. Faybishenko-Designed Tensiometer Schematic
(Modified from Faybishenko, 1999)
When the soil is dry, water flows out of the porous ceramic cup into the surrounding soil,
and when the soil is wet, soil water can enter the porous ceramic cup and force water through the
connector into the upper cell. The upper cell usually functions as a water reservoir to maintain
39
the constant water level in the lower cell which is at the bottom of the connector tube, and just
above this water level in the lower cell an air volume exists which is under pressure
(Faybishenko, 2000b).
Before the six pressure transducers were weather-proofed, five of which were 0-15 psig
and one of which was 0-30 psig, the two types of Omega pressure transducers used at the field
site were calibrated using an oil-less, diaphragm, vacuum pump (Gast Manufacturing Inc.,
Benton Harbor, Michigan, USA) at the hydrology lab in Warnell School of Forestry and Natural
Resources, UGA. The Gast vacuum pump was initially calibrated using a mercury manometer
before being used to calibrate the pressure transducers. The Omega pressure transducers were
calibrated so that the pressure response obtained could be converted to a readable pressure. After
several trials recording the pressure response of the pressure transducers as well as the vacuum
pressure applied by the vacuum pump, calibration curves were made using Microsoft Excel. The
equations of the calibration curve were recorded for each transducer, and these equations specific
to each pressure transducer were input into the datalogger to ensure accurate pressure readings in
the field.
The pressure transducers were weather-proofed by first coating the transducers with
Scotchkote Electrical Coating (3M, St. Paul, Minnesota, USA) with up to three separate coatings,
and then put in clay casts and coated with Castin’ Craft EasyCast Clear Casting Epoxy
(Environmental Technology Inc., Fields Landing, California, USA) according to the procedure
outlined in Appendix X. After the weather-proofing, the pressure transducers were re-tested
with the Gast vacuum pump to ensure that heat produced by the EasyCast Clear Casting Epoxy
had not altered the pressure response measured by the pressure transducers. After the weather-
40
proofing process outlined in Appendix X, several pressure transducers stopped working as the
pressure responses were altered, so these malfunctioning pressure transducers were discarded.
Two metal tubes which exited the top of each tensiometer were attached by Tygon tubing
(Saint-Gobain Performance Plastics Inc., Charny, France) to weather-proofed Omega pressure
transducers, which monitored the pressure response of the air space above the water level in the
upper cell and lower cell of the two cell tensiometers. Therefore, the difference of the air
pressures inside the two cells is in equilibrium with the negative matric pressures in the vadose
zone at the depth of the porous ceramic cup (Faybishenko, 2000b).
The porous ceramic cup for each tensiometer is in tight contact with the soil surface after
the tensiometer was placed in a hole augered to the depth desired for soil water movement to be
observed. The hole is hand augered to a diameter of 0.0254 m which is only slightly bigger than
the 0.0222 m diameter of the tensiometers. Once the tensiometer is installed in the augered hole
and is observed to fit in the hole, the tensiometer is removed and soil slurry consisting of sieved
fine sand, silt, clay, and water is poured in the hole. This soil slurry helps to lubricate the device
as the tensiometer is pushed into the hole, and the soil slurry also ensures tight contact with the
surrounding soil surface after the slurry has dried. If air pockets existed around the porous
ceramic cup, then the tensiometer will record incorrect pressures. Matric pressures were
examined and compared to other studies that used tensiometers to verify the accuracy of the
pressures observed in this study (Adams et al., 1994).
The pressure transducers were wired to a CR23X datalogger (Campbell Scientific,
Logan, Utah) located at the center of the three meter-radius circle study area. The datalogger
recorded data every minute and average those data every 10 minutes. Ultimately, averaged 10
minute-data was saved and written to storage on the CR23X datalogger, and every few weeks
41
these data are collected so that new data being recorded do not write over existing data. These
pressure differences between the two cells represent the negative pressures at depth in the vadose
zone. Negative vadose zone pressures at depths along with gravity are the two main forces
moving water from the soil surface to the water table. Just below the soil surface, the water
pressure is the most negative. At the water table, the pressure is zero. Below the water table, the
pressure becomes positive as all the pore spaces are water-saturated, so the force moving water
below the water table is a positive pressure gradient. According to Faybishenko (2000b), matric
pressures (Pla) obtained from the lower cells of the tensiometers are converted to matric heads
(hs) by the following equation:
hs = (Pla / ρw * g) + hl water
In this equation, ρw is the density of water at 20oC (g cm-3), g is the gravitational acceleration
constant (cm s-2), and hl water is the height of the water column in the lower cell of the
tensiometers (cm). Matric pressures were recorded as inches of Hg and converted to dynes cm-2
so that matric heads were calculated in centimeters. Elevation heads were measured relative to
the ground surface of the field site. Matric heads added to elevation heads resulted in total head
data from each tensiometer so that gradients could be established between the three tensiometers.
3.3. Suction Lysimeters
Lysimeters are water sampling devices used to obtain soil water samples at depth by nondestructive means. Trace metals as well as kiln dust can often be found in some lysimeters from
the production of the porous cups. One way to clean the lysimeters before installation to remove
trace metals and kiln dust is to flush the lysimeters with dilute acid (Litaor, 1988; Grossmann
and Udluft, 1991), and some manufacturers of lysimeters also recommend an acid wash at least
of the porous ceramic cup (Soilmoisture Equipment Corp., Santa Barbara, CA).
42
One type of suction lysimeter used in this study is made of a pipe attached to a porous
cup at the bottom with one tube exiting the closed-top so that suction can be applied. Also
known as soil water samplers, suction lysimeters were invented in 1961 under the direction of
Dr. George H. Wagner at the University of Missouri for collecting soil water samples
(Soilmoisture Equipment Corp, 2007). Suction lysimeters can also have two tubes exiting the
closed-top of the pipe so that collected water samples can be pumped out of one tube as a
positive pressure is applied to the other tube, especially by a bicycle pump.
The suction lysimeters with two tubes exiting the closed-top were installed at 0.5 m, 1.0
m, and 1.5 m depths around the 3 m-radius circle study area, and shall be known as the original
suction lysimeters. Four additional suction lysimeters (Soilmoisture Equipment Corp., Santa
Barbara, California, USA) were also used and all of these additional suction lysimeters were
located within six meters or less outside the three meter radius circle study area at depths of 0.35
m, 0.5 m, 1.25 m, and 1.75 m. To supplement these four lysimeters, two more lysimeters were
purchased (Soilmoisture Equipment Corp.), three lysimeters were built, and these new lysimeters
were all installed. These nine suction lysimeters purchased and built (Soilmoisture Equipment
Corp.) are called the additional suction lysimeters.
3.3.1. Original Suction Lysimeters
The suction lysimeters that are emplaced along the circumference of the three meter
radius circle were constructed from clear poly-vinyl chloride pipe with a diameter of 0.0254 m.
These clear poly-vinyl chloride lysimeters consist of a length of pipe approximately 0.10 to 0.35
m longer than the length at which soil water sampling is desired to occur at depth. The bottom of
the poly-vinyl chloride pipe was fixed to a ceramic porous cup (Soilmoisture Equipment Corp.,
Santa Barbara, California, USA). The top of the clear poly-vinyl chloride pipe is corked with a
43
tight-fitting rubber stop cock, which has two holes where two tubes with a diameter of 0.006 m
exit the stopper. The lengths of the clear poly-vinyl chloride pipe for the lysimeters buried along
the three meter radius circle are 0.60 m, 1.21 m, and 1.82 m. The porous ceramic cups of these
lysimeters are located at 0.5 m, 1.0 m, and 1.5 m depths where soil water samples were
extracted.
The suction lysimeters were placed in a hole augered to the depth desired for soil water
sampling to be collected by a Giddings probe mounted on the back of an all-terrain vehicle. The
0.0381m-diameter of the augered hole for the suction lysimeters was only slightly larger than the
0.0254 m-diameter of the lysimeter.
3.3.2. Additional Suction Lysimeters
Six of the nine additional suction lysimeters were purchased (Soilmoisture Equipment
Corp., Santa Barbara, California, USA), and three of the lysimeters were built using porous
ceramic cups (Soilmoisture Equipment Corp.). Eight of the nine lysimeters were emplaced
within six meters or less distance along the peripheral of the three meter radius study area, while
one lysimeter was emplaced inside the circle near the center. All of these additional lysimeters
have a diameter of 0.048 m. Lysimeters L1 and L2 are located about five meters south of the
three meter radius circle, were approximately 0.46 m and 0.61 m in length, respectively, and the
porous ceramic cups of these lysimeters were emplaced at depths of 0.35 m and 0.50 m,
respectively. Lysimeter L4 was located about six meters south-southeast of the three meter
radius circle, was 1.82 m in length, and was emplaced at a depth of 1.75 m. Lysimeter L3 was
located three meters east of the original study area circle, was 1.82 m in length, and was
emplaced at a depth of 1.25 m. Lysimeter L1A and lysimeter L2B were clustered within one
meter distance of lysimeter L3 to observed soil water in one small area at three different depths:
44
L1A was at a depth of 0.35 m, L2B was at a depth of 0.5 m, and L3 was at a depth of 1.25 m.
Lysimeter L1A was 0.40 m in length and lysimeter L2B was 0.70 m in length. Lysimeter L2A
was located inside the original study area circle, was 0.61 m in length, and was emplaced at a
depth of 0.50 m. Lysimeter L3A and L4A were located three meters west of the original study
area circle, were 1.35 m and 2.00 m in length, respectively, and were emplaced at depths of 1.25
m and 1.75 m, respectively (Figure 3.2).
For each lysimeter, the holes were augered so that the porous cup was located at one of
the four sampling depths: 0.35 m, 0.5 m, 1.25 m, or 1.75 m depth. Each hole was dug using a
hand-auger approximately 0.044 m in diameter; each hole had to be widened slightly with the
auger to allow the nine additional suction lysimeters enough space to fit as the diameter of the
nine suction lysimeters was 0.048 m.
3.4. Backfilling with Soil Slurry
Once the lysimeters were emplaced in the augered hole and were observed to fit in the
hole, soil slurry consisting of sieved fine sand, silt, clay, and water was poured in the hole. This
soil slurry helped to lubricate the device as each of the lysimeters were pushed in the hole. The
soil slurry also ensured tight contact with the surrounding soil surface after the slurry had dried.
If air pockets existed around the porous ceramic cup, then the suction lysimeter would not work
properly. Slurry was added over several days to ensure that no air space existed around the
porous ceramic cup and the body tube of the suction lysimeters as precipitation could infiltrate
along a preferential path along the side of the suction lysimeter body tube from the surface to
depth which would be undesirable.
45
3.5. Operation of Lysimeters
A suction of 0.5 bars of pressure was applied to the tube exiting the top of the lysimeter,
and the tube was crimped and closed using a closed-jaw Hoffman screw-compressor clamp for
flexible tubing (Wards Natural Science Inc., Rochester, New York, USA) so that suction could
not escape. The operational suction range for vacuum lysimeters equipped with a porous
ceramic cup was less than 0.6 to 0.8 bars (ASTM, 1995). This applied suction pulled in a soil
water sample through the porous cup over a 24 hour period. If there was a leak in the lysimeter,
then the applied suction would escape within minutes rendering the lysimeter ineffective in soil
water sampling (ASTM, 1995), which was probably the reason for failure of the original suction
lysimeters. After the 24 hour period, the sample could be removed from the lysimeter using a
hand vacuum pump, or if the suction lysimeter had two tubes exiting the top of the closed pipe
body, then a bicycle pump could pump the soil water sample out of the device. Both methods
were used effectively to remove soil water samples from the suction lysimeters at the study site.
3.6. Geochemical Analyses
Four geochemical tests including ion chromatography (for nitrate-N), spectrophotometry
(for ammonium, urea, iron2+, and total iron used to calculate iron3+), and element-specific
machines for dissolved organic carbon and total nitrogen were used to analyze all of the soil
water samples, the rain water samples, and the piezometers groundwater samples at the Nutrients
Lab in the U.S. Environmental Protection Agency National Exposure Research Laboratory in
Athens, GA, under the direction of Dr. Caroline Stevens and lab technician Lidia Samarkina.
After the first suite of soil water samples was tested, the extremely low concentrations of urea-N
measured led to the decision to quit measuring for urea. Concentrations of urea-N were probably
volatilizing quickly from soil water as nitrogen gas before urea could be properly quantified.
46
Nitrate-N was measured in filtered water samples using a Metrohm ion chromatograph
with a Metrosep A Supp Five anion column and an 853 CO2 suppressor that used a conductivity
detector to receive the nitrate signal and make peak areas. Stock anion eluent was made of 13.56
g Na2CO3 and 3.36 g NaHCO3 in 200 mL of solution. The ion chromatograph used 10 mL of
eluent for every two liters of samples analyzed. The anion column was last replaced May 12,
2009. Urea-N and ammonium-N were measured in filtered water samples with a Hach 2010
spectrophotometer according to standard procedures of the phenate method (Clesceri et al.,
1998). In the phenate method, ammonia is reacted with hypochlorite and phenol to form indophenol blue, the intensity of which is read at λ = 640 nm using the spectrophotometer. Total
nitrogen was measured in non-filtered water samples by oxidizing all of the nitrogen species
using ultra-pure O2 gas to nitrogen oxide gas which is measured using thermal decomposition
around 720 oC and chemiluminescence with a Shimadzu total nitrogen module (TNM-1), which
has a range of 0 – 4000 mg L-1 total N, repeatability CV within three percent.
Dissolved organic carbon was measured using filtered water samples dosed with HCl
until the pH was between two and three on a Shimadzu 5050A Total Organic Carbon Analyzer
according to standard procedures (Washington et al., 2004). Iron2+ and iron3+ (from total iron)
analyses used 50 mL crimp-seal, acid washed glass serum bottles, which were acidified using 8 –
10 μL concentrated HCl and purged with grade-five N2 gas for one minute; water samples were
collected, filtered, and injected into these acidified, purged, closed serum bottles in the field. In
the lab, Fe2+ and Fe3+ were measured by the ferrozine procedure according to standard methods
(Viollier et al., 2000; Washington et al., 2004).
47
3.7. Soil Analyses
The soil water pH (pHw) was measured to determine if pH-sensitive nitrogen
transformation processes would take place. Denitrification and nitrification are pH-limited, and
soil pH needs to be above pH five for these processes to occur (McNeill and Unkovich, 2007).
Soil pH is influenced by the bedrock from which the soil is derived and the pH of infiltrating
rainwater. Acid rain by definition has a pH of 5.7 or below and is due mainly to the mineral
acids H2SO4 and HNO3 in precipitation with a minor contribution of HCl (Stevenson and Cole,
1999). Soil pH can also be decreased by fertilizing with ammonium sulfate fertilizer and the
process of nitrification can lower soil pH as well (Bohn et al., 2001).
The water pH of soil was a one-to-one ratio of water to soil which was analyzed using a
pH probe in the lab and in the field. Analyses used soil samples which were collected by auger
at depths where the porous ceramic cups were located for the suction lysimeters. In the field
once the soil was collected with an auger, the soil was combined with de-ionized water, shaken
for a minute, and the pH probe was inserted into the soil slurry where readings were recorded
after the pH readings stabilized. The same procedure was used in the lab after soils samples
were transported in air-tight containers back from the field.
3.8. Time Domain Reflectometry and Water Table Monitoring
Time domain reflectometry (TDR) probes were installed previously by the USDA-ARS
and were monitored for several months of this study to supplement moisture potential monitoring
in the tensiometers. TDR data infers water content from the dielectric permittivity of the
medium, and some TDR systems, like the ones in this study, can measure volumetric water
content of a soil at different segments of depths along the length of the waveguide, which
produces a volumetric water content profile for a certain soil (Jones et al., 2002). Four TDR
48
probes were previously installed and each probe was segmented in 0 – 15, 15 – 30, 30 – 60, and
90 – 120 cm depths. The TDR ranges of depths were reported as the deepest depth in the range
for all TDR graphs. TDR soil moistures were recorded several times a month if possible.
Moistures were read using a device provided by the USDA-ARS, which converted dielectric
permittivities immediately into soil moistures utilizing the Topp’s equation.
The water table elevations surrounding the field site were monitored several times a
month for several months by measuring depth to the water table in several piezometers which
were previously installed by the USDA-ARS. The water table elevations were measured from
the water table to the top of the casing at the ground surface for each piezometer. The device to
measure the depth to the water table had a sensor attached to a metric measuring tape at the zero
mark, and the sensor made a buzzing sound when immersed in water.
3.9. Nitrogen Stable Isotope Analyses
Nitrogen has 2 stable isotopes: 14N which makes up 99.6337% of nitrogen on Earth, and
15
N which makes up 0.3663% of nitrogen on Earth. The isotopic composition of N is expressed
as δ15N which is defined as:
δ15N = [(15N/14N)Sample – (15N/14N)Standard / (15N/14N)Standard] X 103 0/00
where the standard is atmospheric N2 gas in air. The nitrogen isotopic signatures are measured
with isotope ratio mass spectrometers which convert all nitrogen species to N2 gas (Faure and
Mensing, 2005).
The nitrogen isotope ratio 15N/14N, called natural abundance, can be changed through
processes like evaporation, condensation, freezing, melting, chemical reactions, and biological
processes (Freeze and Cherry, 1979). Trophic transfers can increase the natural abundance of an
organism by +3 to +5 0/00 (Dodds, 2002). Values of 15N from atmospheric nitrogen are constant
49
depending on elevation, and are known as the zero point of the naturally occurring nitrogen
isotope variations (Hoefs, 2004).
The isotopic composition of N derived from different sources can be used to trace
nitrogen movement through soil systems because fertilizer and animal waste have distinct δ15N
values (often a range of values). The δ15N range for nitrate from rain is -13 to +2 0/00, and the
δ15N range for ammonium from rain is -13 to +2 0/00. The δ15N range for nitrate from animal
waste is +8 to +22 0/00 (Faure and Mensing, 2005).
Stable nitrogen isotopes were determined for the two suspected sources of nitrogen to
W2, manure and chemical fertilizers, as well as four soil samples from two depths. The stable
nitrogen isotope analyses were performed at the UGA Institute of Ecology’s Analytical
Chemistry Lab by Tom Maddox. The stable nitrogen isotope, 15N, was analyzed in the manure
and chemical fertilizers to establish end-members for the analyses, and δ15N values were
compared to atmospheric N2 gas (AIR). The δ15N values were produced first for the endmembers, manure and the chemical fertilizers, and then for the soil samples and water samples.
The isotopic analyses used a Thermo-Finnigan Delta C Mass Spectrometer coupled to a
Carlo Erba CN Analyzer via Thermo-Finnigan Conflo II Interface. The Thermo pieces were
made in Bremen, Germany, and the Carlo Erba was made in Milan, Italy. The precision for
isotope analysis was +/- 0.15 per mil or better (Tom Maddox, personal communication).
A hole was augered and soil samples removed from a site carefully chosen in between
lysimeter L3 and piezometer 5, approximately 2.4 m away from lysimeter L3 in a direction
believed to be down-gradient of groundwater flow so as not to influence local hydrology and
nutrient delivery to the lysimeters in the area of high nitrogen. Due to limited funds, only four
soil samples were chosen to represent the soils in the nitrogen isotope analyses: two samples
50
from 0.5 m depth and two samples from 2.0 m depth. The samples were all removed from the
same augered hole, and the depths were picked to represent “shallow” and “deep” soils.
51
CHAPTER 4
RESULTS AND DISCUSSION
4.1. Introduction
The results of this study include soil pH and grain-size profiles to 2.0 m of the vadose
zone, USDA-ARS fertilization schedules for Watershed 2, soil water and precipitation
geochemical analyses for nitrate, ammonia, urea, total nitrogen, dissolved organic carbon, iron2+
and iron3+, summarized tensiometric data, moisture release curves for Cecil soils, isotopic
analyses of nitrogen sources to the field site, and discussion of these results. In the discussion,
several theories have been proposed to explain observed trends in the geochemical results.
4.2. Soil Analyses
The soil water pH results can be seen below in Figure 4.1 as well as in Appendix D.
Random variation of 0.1 – 0.2 pH units was allowed in replicate determinations, and could result
from using different laboratories for analyses or different instruments. Given this acceptable
range in soil pH values, these four soil samples show that low soil pH (below 5) could inhibit
denitrification and nitrification at certain depths. Specifically, the soil acidity in these values
could inhibit denitrification in Soil 1 at 0.5 m, 1.5 m, and possibly at 2.0 m depth, in Soil 2 at 1.5
m depth, and in Soil 4 at 1.5 m and 2.0 m depths, and nitrification could be inhibited in any of
the soil samples with pH below 5.5, which is a threshold below which nitrification slows.
However, precision of soil water pH can be influenced by presence of carbon dioxide gas in soil
samples, specifically causing lower than actual readings, and a common product of
denitrification is carbon dioxide (Reid, 1998). Figure 4.1 shows the changes in pH in the four
52
soil samples collected from the field site. In general, the soil pH of the field site soils decreases
with increasing depth.
Soil pH
4.6
4.8
5
5.2
5.4
5.6
5.8
6
6.2
6.4
6.6
0.0
0.2
0.4
0.6
Depth (m)
0.8
1.0
1.2
Soil 1
Soil 2
1.4
Soil 3
1.6
Soil 4
1.8
2.0
Figure 4.1. Soil pH vs. Depth for W2 Field Site Soils
Grain size as well as pore size can affect and influence vadose zone hydrology and pore
water velocity. Grain size analyses were made on several soil samples within the original three
meter radius circle study area at different depths and can be seen below in Table 4.1 (see also
Appendix D). Soil A was located at 0.5 m depth in between the original lysimeter and
tensiometer at 0.5 m depth. Soil B was located at 1.0 m depth in between original lysimeter and
53
tensiometer at 1.0 m depth. Soil C was located at 1.5 m depth in between original lysimeter and
tensiometer at 1.5 m depth. Soils D, E, and F were located at 1.6 m, 1.8 m, and 2.0 m depth,
respectively, within one meter of the field box at the center of the three meter study circle
(Appendix D).
Table 4.1. Particle Size Distribution Analyses
Sample # Depth (m) Sand (%)
A
0.5
30
B
1.0
43
C
1.5
45
D
1.6
36
E
1.8
31
F
2.0
33
Silt (%)
16
24
19
9
11
18
Clay (%)
54
33
36
55
58
49
Most of the soils sampled would be categorized as clayey soils due to the high clay
percentage of each. Particle size distributions were performed using Stoke’s law of settling
particles. Many of the samples with high clay particle percentages remained in suspension for
days. These clayey soils are not believed to be formed in place from weathering of the bedrock
as there is no decrease in clay sized particles with depth as would be seen in a naturally
developed soil profile. Instead, the clayey soils were probably moved here in the past from the
digging of a pond historically noted to be just down elevation from the spring and the field site,
which since has filled in due to the creation of a dam, or due to improper tillage practices of the
past that may have aided in clays being washed from higher elevations in the watershed to the
lower elevation field site.
4.3. Fertilization of Watershed 2
Fertilization application results can be seen below in Table 4.2. These results show type
of fertilizer in terms of N-P-K with P as P2O5 and K as K2O and quantities of raw fertilizer
applied as well as quantities of nitrogen applied. The fertilizer quantities are shown in pounds
54
per acre first as these were the units used by the fertilizer manufacturers, and secondly, the
fertilizer quantities have been converted to metric units. Other watershed management events
such as spraying of pesticides/insecticides, liming, and planting have been summarized from
January, 1992 to present in Appendix E.
55
Table 4.2. USDA-ARS Watershed 2 Fertilization Management
DATE
Febuary, 1992
April, 1992
December, 1992
March, 1993
July, 1993
November, 1993
Febuary, 1994
March, 1995
January, 1996
April, 1996
November, 1996
3/17/1997
March, 1998
October, 1998
Fall 1998
Early 1999
2/1/1999
4/1/1999
10/1/1999
2/23/2000
10/1/2000
9/26/2001
2/14/2002
9/24/2002
2/25/2003
10/7/2003
4/1/2004
1/16/2005
3/5/2005
8/20/2006
9/11/2006
8/7/2007
2008
9/10/2009
FERTILIZER TYPE
(N, P as P2O5, K as K2O)
10-10-10
34-0-0
18-0-27
18-0-27
34-0-0
14-7-14
17-0-17
10-10-10
10-10-10
15-0-15
10-10-10
10-10-10
10-10-10
17-17-17
10-10-10
FERTILIZER
APPLIED
400 lbs/ac
150 lbs/ac
225 lbs/ac
225 lbs/ac
180 lbs/ac
300 lbs/ac
300 lbs/ac
400 lbs/ac
400 lbs/ac
400 lbs/ac
400 lbs/ac
400 lbs/ac
400 lbs/ac
300 lbs/ac
400 lbs/ac
(448 kg/ha)
(168 kg/ha)
(252 kg/ha)
(252 kg/ha)
(202 kg/ha)
(336 kg/ha)
(336 kg/ha)
(448 kg/ha)
(448 kg/ha)
(448 kg/ha)
(448 kg/ha)
(448 kg/ha)
(448 kg/ha)
(336 kg/ha)
(448 kg/ha)
QUANTITY OF
NITROGEN APPLIED
40 lbs
50 lbs
41 lbs
41 lbs
61 lbs
42 lbs
51 lbs
40 lbs
40 lbs
60 lbs
40 lbs
40 lbs
40 lbs
51 lbs
40 lbs
Note: Fences removed from watershed creating modern W2
34-0-0
200 lbs/ac (224 kg/ha)
68 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
17-17-17
300 lbs/ac (336 kg/ha)
51 lbs
80 lbs N/ac (90 kg N/ha
80 lbs
Nitrogen
34-0-0
200 lbs/ac (224 kg/ha)
68 lbs
34-0-0
300 lbs/ac (336 kg/ha)
102 lbs
Urea w/ Sulfur (33-0-0)
200 lbs/ac (224 kg/ha)
66 lbs
15-0-15
200 lbs/ac (224 kg/ha)
30 lbs
Urea w/ Sulfur (33-0-0)
250 lbs/ac (280 kg/ha)
83 lbs
No Fertilizer Applied
Urea w/ Sulfur + Nutrisphere*
(33-0-0)
200 lbs/ac (224 kg/ha)
66 lbs
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
(45 kg N/ha)
(56 kg N/ha)
(46 kg N/ha)
(46 kg N/ha)
(69 kg N/ha)
(47 kg N/ha)
(57 kg N/ha)
(45 kg N/ha)
(45 kg N/ha)
(67 kg N/ha)
(45 kg N/ha)
(45 kg N/ha)
(45 kg N/ha)
(57 kg N/ha)
(45 kg N/ha)
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
N/ac
(76 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(57 kg N/ha)
(90 kg N/ha)
(76 kg N/ha)
(114 kg N/ha)
(74 kg N/ha)
(34 kg N/ha)
(92 kg N/ha)
N/ac (74 kg N/ha)
* First application with nutrisphere added to maintain ammonia-N longer in soil & to prevent volatilization losses
56
4.4. Moisture Release Curves, TDR, and Tensiometric Data
Soil water characteristic curves express the relationship between pressure head and the
soil water content. The soil water characteristic curve data was observed in a Cecil Series soil at
the USDA-ARS, and three curves were made at 0.5, 1.0, and 1.5 m depths which are the same
depths where each of the original tensiometers has measured matric pressures in this study
(Figure 4.2).
0.430
MRC at 0.5 m Depth
0.410
MRC at 1.0 m Depth
Moisture Content (cm3 cm-3 )
0.390
MRC at 1.5 m Depth
0.370
0.350
0.330
0.310
0.290
0
-100
-200
-300
-400
-500
-600
-700
-800
-900
Matric Potential (cm H2O)
Figure 4.2. Cecil Soil Moisture Release Curves at 0.5, 1.0, & 1.5 m Depths
(data from Bruce et al., 1983)
57
-1000
50.0
Soil Moisture (%)
45.0
40.0
35.0
30.0
45.0-50.0
25.0
40.0-45.0
20.0
35.0-40.0
15.0
30.0-35.0
25.0-30.0
10.0
20.0-25.0
15.0-20.0
10.0-15.0
Depth (m)
Date (mm/dd/yyyy)
Figure 4.3. TDR Soil Moisture vs. Time vs. Depth
TDR data from January, 2009, through June, 2009, for five depths were made into a
surface which shows that the soil moisture profiles have been consistently similar (Figure 4.3).
TDR soil moisture profiles from January through April, 2009, have higher than average moisture
contents , whereas TDR soil moisture profiles from May and June, 2009, have below average
soil moisture contents (Figure 4.3). Soil moisture dried considerably with lower moisture
contents from April through June, 2009 (Figure 4.3). The TDR soil moisture profiles from May
and June, 2009, especially show the beginning of increased evapotranspiration and summer
drought, and soil moisture profiles never achieved such low moisture contents in the entirety of
TDR monitoring.
58
59
11/18/09 12:00 AM
11/8/09 12:00 AM
10/29/09 12:00 AM
10/19/09 12:00 AM
10/9/09 12:00 AM
9/29/09 12:00 AM
9/19/09 12:00 AM
9/9/09 12:00 AM
8/30/09 12:00 AM
8/20/09 12:00 AM
8/10/09 12:00 AM
7/31/09 12:00 AM
7/21/09 12:00 AM
7/11/09 12:00 AM
7/1/09 12:00 AM
6/21/09 12:00 AM
6/11/09 12:00 AM
6/1/09 12:00 AM
5/22/09 12:00 AM
5/12/09 12:00 AM
5/2/09 12:00 AM
4/22/09 12:00 AM
4/12/09 12:00 AM
4/2/09 12:00 AM
3/23/09 12:00 AM
3/13/09 12:00 AM
3/3/09 12:00 AM
2/21/09 12:00 AM
2/11/09 12:00 AM
2/1/09 12:00 AM
1/22/09 12:00 AM
1/12/09 12:00 AM
1/2/09 12:00 AM
12/23/08 12:00 AM
12/13/08 12:00 AM
12/3/08 12:00 AM
11/23/08 12:00 AM
11/13/08 12:00 AM
11/3/08 12:00 AM
10/24/08 12:00 AM
10/14/08 12:00 AM
10/4/08 12:00 AM
9/24/08 12:00 AM
10.000
40
6.000
60
4.000
80
0.000
-2.000
Precipitation (mm)
9/14/08 12:00 AM
Hydraulic Gradient (cm cm-1 )
12.000
0
20
8.000
2.000
100
120
Date-Time (mm/dd/yy hh:mm)
Figure 4.4. Hydraulic Gradients and Precipitation vs. Time
Hydraulic Gradient
from 0.5 to 1.0 m
Hydraulic Gradient
from 1.0 to 1.5 m
Hydraulic Gradient
from 0.5 to 1.5 m
Rain (mm)
The hydraulic gradient is plotted between the three tensiometers: from the tensiometer at
0.5 m depth to the tensiometer at 1.0 m depth (blue line), from the tensiometer at 1.0 m depth to
the tensiometer at 1.5 m depth (green line), and from the tensiometer at 0.5 m depth to the
tensiometer at 1.5 m depth (red line) in Figure 4.4. The three lines were calculated from the
difference between the total heads measured at each tensiometer divided by the length between
the porous cups of the tensiometers. Hydraulic gradient from 1.0 to 1.5 m is the largest which
can be seen as the green line above in Figure 4.4. The smallest hydraulic gradient is from 0.5 to
1.0 m which can be seen as the blue line in Figure 4.4. Also, the hydraulic gradient from 0.5 to
1.0 m is frequently negative, which means that the direction of flow would be upwards between
the two tensiometers. The hydraulic gradient from 0.5 to 1.5 m is slightly less than the hydraulic
gradient from 1.0 to 1.5 m which is due to the addition of the negative hydraulic gradient from
0.5 to 1.0 m. Small hydraulic gradients as well as negative hydraulic gradients could be caused
by high clay contents somewhere in between the tensiometers, especially from 0.5 to 1.0 m,
which would cause the water potential to slow dramatically due to the low hydraulic conductivity
of clay. Also, precipitation increased in the last four months of the study from August till
November, 2009, which can be seen on the second y-axis in Figure 4.4. Precipitation events also
increased in duration in the last four months with events often lasting several days instead of
intermittent events which dominated the first eleven months of the study from September, 2008
till July, 2009.
60
4.5. Soil Water, Groundwater, and Precipitation Geochemical Results
The results of the geochemical analyses on the soil water samples from the lysimeters can
be seen in Figures 4.6 through 4.13, and the spatial distribution of lysimeters can be seen in
Figure 4.5. The geochemical results can also be seen in Appendix F. The nitrogen
transformation processes are discussed in greater detail below along with more specific patterns
in the geochemical results. The stable oxidation states of nitrogen within the stability of water
are nitrate, ammonia, and N2 (Bohn et al., 2001). Patterns can be seen in the concentrations of
the measured species and are discussed below in further detail. Although nitrate concentrations
were spatially variable at the study site, nitrate decreased with depth to the deepest lysimeter at
1.75 m, which is above the water table (see Appendix A for water table elevation data). Total
nitrogen concentrations were also spatially variable, but generally decreased with depth to 1.75
m just above the seasonably varying water table. Ammonium concentrations, especially where
lysimeters are clustered together in close areas, decrease with depth to 1.75 m. Urea was
measured only for the first sampling event as urea appears to be fleeting and quite likely
volatilizes as nitrogen gas.
61
Figure 4.5. Field Site Map in W2
62
Figure 4.6. Soil Water Nitrate-N Concentrations vs. Depth
Soil water nitrate-N concentrations varied most in the uppermost sampling depth at 0.35
m, and concentrations varied less with increasing depth at 0.5 and 1.25 m (Figure 4.6).
However, nitrate-N soil water concentrations varied least at the deepest sampling depth at 1.75 m
(Figure 4.6). At the shallowest sampling depth, 0.35 m, soil water nitrate-N concentrations
varied from 0.85 to 50.28 mg L-1, and the mean and median concentrations were very similar
suggesting a normal distribution of data. Also, the symmetry of the box plot about the median
suggests a normal distribution of data at 0.35 m depth. At the next shallowest sampling depth of
0.5 m, soil water nitrate-N concentrations varied from 0.89 to 64.80 mg L-1 sampled from three
separate lysimeters L2, L2A, and L2B, and the distribution of the data is outlier-prone, especially
63
in the higher concentration data. At 1.25 m depth, soil water nitrate-N concentrations varied
from 3.74 to 21.71 mg L-1, and the mean and median are very close in concentration which along
with the symmetry of the box plot about the median suggests a symmetrical distribution. At 1.75
m depth, the deepest sampling depth, soil water nitrate concentrations varied the least from 0.24
to 5.13 mg L-1, and the mean and median concentrations were very similar suggesting a normal
distribution of data. Also, the box plot was symmetrical about the median at depth 1.25 m
suggesting a symmetrical distribution of concentrations. Errors in sampling were larger than
machine errors, and are presented in Table 4.3 along with mean nitrate concentrations per depth
and results of a statistical analysis. Also, in Figures 4.12 and 4.13, nitrate concentrations per
lysimeter were plotted along with precipitation vs. time in order to examine if any trends were
present. Further discussion of nitrate concentration results and trends follows (p. 79).
64
Figure 4.7. Soil Water Ammonium Concentrations vs. Depth
Soil water ammonium concentrations varied the most at 0.5 m depth and the least at 1.75
m depth (Figure 4.7). Soil water ammonium concentrations varied from 0.025 to 0.236 mg L-1at
0.35 m depth, and the similarity of the mean and median suggests the distribution was somewhat
normal. At depth 0.5 m, ammonium concentrations varied from 0 to 0.904 mg L-1, and the
distribution was outlier-prone. Also, the mean and median at depth 0.5 m were not very similar..
At depth 1.25 m, ammonium concentrations varied from 0 to 0.376 mg L-1, 50% of the
concentration data represented by the box plot (Figure 4.7) varied the most of all the sampling
depths, and the distribution of concentrations was not normal because the mean and median were
not similar. At depth 1.75 m, soil water ammonium concentrations varied from 0 to 0.084 mg L65
1
, and the similarity of the mean and median as well as the symmetry of the box plot about the
median (Figure 4.7) suggest normal distribution of the data. Mean soil water ammonium
concentration per depth, errors, and statistics can be seen in Table 4.3. Further discussion of
ammonium concentrations and trends follows (p. 79 through 84).
Figure 4.8. Soil Water Total Nitrogen Concentrations vs. Depth
In terms of total nitrogen, soil water varied most in concentrations at 0.5 m depth and
least in concentrations at 1.75 m depth (Figure 4.8). Soil water total nitrogen concentrations
mimicked nitrate-N soil water concentrations in that total nitrogen concentrations were often
slightly higher than nitrate-N concentrations, and due to lab costs, total nitrogen was
66
discontinued from geochemical analyses after October, 2009. At depths 0.35 and 0.5 m, total
nitrogen concentrations varied and did not have normal distributions due to non-symmetrical
boxes about the medians. However, at depths 1.25 and 1.75 m depths, total nitrogen
concentrations had normal distributions due to similar means and medians and symmetry of the
boxes about the medians (more so at depth 1.75 m). Also, total nitrogen varied less at depths
1.25 and 1.75 m. Mean soil water total nitrogen concentrations per depth, statistics, and errors
can be seen in Table 4.3. Further discussion of total nitrogen follows (p.80).
Figure 4.9. Soil Water Dissolved Organic Carbon Concentrations vs. Depth
67
Soil water dissolved organic carbon (DOC) concentrations varied most in the uppermost
depth at 0.35 m depth from 85.2 to 7.4 mg L-1, and the distribution was not normal at this depth
due to the unsymmetrical box about the median value (Figure 4.9). The 0.5 m depth also varied
over a large range and was not symmetrical about the median value. The 1.25 m depth varied
over a smaller range compared to the shallower depths, but had one outlying datum of 94.63 mg
L-1 well above the 75th percentile. Soil water DOC concentrations varied the least at 1.75 m
depth from 0 to 8.193 mg L-1, and concentrations followed a normal distribution due to the
similarity of the mean and median and the symmetry of the box about the median concentration.
In general, DOC concentrations decreased with increasing depth, which is often the case in soils
due to DOC being used up by soil microbes as energy in oxidation-reduction reactions. Mean
soil water DOC concentrations per depth, statistics, and errors can be seen in Table 4.3. Further
discussion of DOC results follows (p. 83 through 85 and p. 92 through 93).
68
Figure 4.10. Soil Water Ferrous Iron Concentrations vs. Depth
Soil water ferrous iron (Fe2+) concentrations appear consistent ranging typically from 0 to
1 mg L-1 at depths 0.5 and 1.25 m, but concentrations varied more at depths 0.35 and 1.75 m
from 0 to 6.674 mg L-1 and 0 to 2.416 mg L-1 ,respectively (Figure 4.10). Ferrous iron
concentrations were only normally distributed at 0.5 m depth where mean and median
concentrations were similar and the box plot was symmetrical about the median (Figure 4.10).
Mean soil water ferrous iron concentrations per depth, statistics, and errors can be seen in Table
4.3. A discussion of ferrous and ferric iron follows as a possible terminal electron acceptor in
oxidation-reduction reactions and denitrification (p. 85).
69
Figure 4.11. Soil Water Ferric Iron Concentrations vs. Depth
Although each sampling depth has outlier-prone distributions, soil water ferric iron (Fe3+)
concentrations have similar ranges and means at each depth (Figure 4.11). The range of ferric
iron concentrations at each depth is 0.05 to about 0.30 mg L-1. More variability in soil water
ferric iron concentrations is characteristic of lysimeters at 0.35 and 0.5 m depths where the
majority of ferric iron concentrations data spans twice as large of a range, around 0.05 to about
1.7 mg L-1. Outlying high ferric iron concentrations at 0.35 m increased the mean concentration
above the 75th percentile. Ferric iron concentrations did not have a normal distribution at any
depth sampled. At depth 1.75 m, the ferric iron concentration distribution was skewed only
slightly because the box plot is nearly symmetrical about the median concentration and the mean
70
and median are somewhat similar. Mean soil water ferric iron concentrations per depth,
statistics, and errors can be seen in Table 4.3.
Table 4.3. Soil Water Summary Statistics with Depth
Chemical
Variable
Number
Minimum Maximum Mean
of
-1
-1
-1
Depth (m) Samples (mg L ) (mg L ) (mg L )
Sampling Standard
Error +/- Deviation
-1
(mg L )
-1
(mg L )
0.35
16
0.85
50.28
19.6
0.11
16.6
(NO3 -N)
0.5
1.25
1.75
17
23
29
0.89
3.74
0.24
64.80
21.71
5.13
7.01*
17.4*
2.88
2.06
1.78
0.14
20.7
4.28
1.64
Ammonium
0.35
15
0.025
0.236
0.058*
0.014
0.053
(NH4 )
0.5
1.25
1.75
22
23
29
0.000
0.000
0.000
0.904
0.376
0.084
0.047*
0.084*
0.035
0.010
0.013
0.005
0.207
0.111
0.021
Total N
(TN)
0.35
0.5
1.25
1.75
0.35
0.5
1.25
1.75
0.35
8
11
12
20
14
20
23
29
13
2.643
2.86
4.606
0.3203
7.40
2.5
0.41
0.00
0.005
33.42
76.83
22.029
5.770
85.2
40.6
94.63
8.193
6.674
17.49
8.498*
18.87*
3.86
37.5
12.8
2.17*
1.23*
0.351*
0.137
0.63
0.149
0.321
0.00
0.30
1.72
0.94
0.001
10.8
28.5
4.68
1.63
24.68
13.09
19.36
2.048
1.792
0.5
1.25
1.75
0.35
21
20
24
13
0.000
0.000
0.000
0.062
0.636
1.112
2.416
1.666
0.046*
0.024*
0.214*
0.115*
0.317
0.286
0.229
0.009
0.180
0.292
0.665
0.500
0.5
1.25
1.75
* = value excludes outliers
21
20
24
0.052
0.055
0.052
1.537
0.726
0.647
0.081*
0.108*
0.134*
0.738
0.138
0.223
0.327
0.159
0.152
Nitrate-N
-
+
Dissolved
Organic
Carbon
(DOC)
Ferrous
Iron (Fe 2+)
Ferric
Iron (Fe 3+)
Mean soil water concentrations and statistics per depth (Table 4.3) can be compared to
mean values for each chemical variable and statistics per chemical variable (Table 4.4). Mean
concentrations per depth have outliers removed from calculations in order to obtain a more
accurate mean value not influenced by outliers (Table 4.3). However, when examining mean
71
concentrations for each chemical variable, all data were used which may increase mean values
(Table 4.4) compared with mean values for each depth (Table 4.3).
Table 4.4. Soil Water Summary Statistics for Each Chemical Variable
Number
Minimum Maximum Mean
of
Chemical
Variable Samples (mg L-1 ) (mg L-1 ) (mg L-1 )
Sampling
Error +/-1
(mg L )
Standard
Deviation
-1
(mg L )
2
R *
F
Value*
-
85
0.242
64.80
11.93
2.06
13.46
0.254
9.20
NH4
Total N
DOC
+
89
51
86
0.000
0.320
0.000
0.904
76.83
94.63
0.077
12.90
11.43
0.014
0.63
1.72
0.122
15.34
19.38
0.096
0.230
0.395
2.23
4.69
17.9
2+
78
0.000
6.674
0.325
0.317
0.848
0.091
2.47
NO3 -N
Fe
3+
78
0.052
1.666
0.195
0.738
0.286
0.035
0.90
*Statistical analysis (ANOVA) was run in SAS using a model comparing each chemical variable to depth
2
in order to see if depth influenced the chemical variable (where R and F come from)
Fe
Also, mean soil water concentrations per depth for each chemical variable (Table 4.3)
and mean concentrations for chemical variables, in general, (Table 4.4) can be compared to
groundwater concentrations in five piezometers across the field site (Table 4.5). Groundwater
concentrations in piezometers 1, 2, 3, 6, and 7 can be thought of as background concentrations
from across the field site because these concentrations are random groundwater samples. Also,
groundwater concentrations can be considered as a mixture of soil water from across the
watershed which has infiltrated past the sampling depths in the vadose zone and then past the
water table. Groundwater concentrations are lower than soil water concentrations especially for
nitrate-N (Table 4.5). Groundwater samples were removed after emptying the piezometers twice
using a vacuum pump, then piezometers were allowed to refill again with what was assumed to
be groundwater alone, and then samples were collected and analyzed (Table 4.5).
72
Table 4.5. Groundwater Concentrations for Piezometers 1, 2, 3, 6, and 7
Fe 3+
Depth to
(Totl. Fe Total Dissolved
Ground+
2+
Fe 2+)
N
Organic C Fe
Piezometer [NO3 -N] [NH4 ]
water
(m)
(mg/L)
(mg/L)
(mg/L)
(mg/L)
(mg/L)
(mg/L)
Location
Date
Up-Slope of
4/28/2009 Site Piez. 7
3.45 0.055 3.561
0.44 0.031 0.068
1.99
6/10/2009
3.81 0.013 3.618
0.764 0.334 1.031
2.13
11/19/2009
4.51 0.149
NA
1.572
NA
NA no data
On-Site
4/28/2009
4.60 0.043 5.322
0.14 0.093 0.128
1.83
Piez. 2
6/10/2009
4.69 0.026 5.103
0.631 0.013 0.088
1.85
11/19/2009
4.41 0.031
NA
0.678
NA
NA no data
Down-Slope
Piez. 3
4.95 0.111 5.665
0.75 0.171 0.287
1.42
4/28/2009
6/10/2009
4.14 0.024 4.287
0.753 0.300 1.281
1.52
11/19/2009
4.51 0.026
NA
0.416
NA
NA no data
Side of Site
2.97 0.059 3.004
1.177 0.305 1.088
2.48
6/10/2009
Piez. 1
11/19/2009
6.87 0.078
8.129
NA
NA no data
Side of Site
6/10/2009
4.24 0.019 4.483
0.604 0.615 0.908
1.70
Piez. 6
11/19/2009
3.31 0.025
NA
0.369
NA
NA no data
NA = no analysis
Stream water samples were collected from 10 sampling locations over two years in both
streams that flow through the wetland and from a flume down-stream of the confluence of both
the wetland streams at the very bottom of Watershed 2. These stream water geochemical data
are presented for comparison with soil water geochemical data since soil water at the field site
eventually percolates to the groundwater which feeds the two streams flowing through the
wetland (personal communication with Katherine Schroer, UGA PhD Geochemistry student,
EPA-ORD technician, 2010).
Stream water nitrate-N concentrations varied from 0.025 to 11.56 mg NO3--N L-1 from
162 data with a mean of 3.69 mg NO3--N L-1 and a standard deviation of 2.27 mg NO3--N L-1.
When compared to mean nitrate-N concentrations for the soil water of 11.93 mg NO3--N L-1 with
73
an error of 2.06 mg NO3--N L-1 and a standard deviation of 13.46 mg NO3--N L-1, the soil water
has higher nitrate-N concentrations than down gradient in the streams. The large nitrate-N
standard deviation for soil water samples was likely due to the outlier-prone distribution of data.
Also, the soil water has a much larger range of nitrate-N concentrations compared with the
stream water, which only ranges up to about the mean soil water concentration (Table 4.4).
Stream water ammonium concentrations varied from 0.003 to 0.392 mg NH4+ L-1 from
130 data with a mean of 0.087 mg NH4+ L-1 and a standard deviation of 0.078 mg NH4+ L-1.
When stream water ammonium is compared to soil water ammonium, the range of soil water
ammonium is more than twice the range of stream water ammonium (Table 4.4). The mean soil
water ammonium concentration was 0.077 mg NH4+ L-1 plus or minus 0.014 mg NH4+ L-1 error
and a standard deviation of 0.122 mg NH4+ L-1, which is lower than the mean ammonium
concentration of stream water samples. Also, the standard deviation of soil water ammonium is
very large compared to the standard deviation of the stream water ammonium, which is likely
due to the outlier prone distribution of data. Furthermore, the soil water ammonium standard
deviation is larger than the soil water mean ammonium concentration.
Stream water total nitrogen (TN) concentrations varied from 1.52 to 10.5 mg TN L-1 from
28 data with a mean of 4.6 mg TN L-1 and a standard deviation of 2.22 mg TN L-1. Soil water
total nitrogen varied from 0.32 to 76.83 mg TN L-1 from 51 data with a mean of 12.90 mg TN L-1
plus or minus an error of 0.63 mg TN L-1 and a standard deviation of 15.34 mg TN L-1. Although
the standard deviation was high for soil water TN (likely due to outliers), the mean and range of
soil water TN was much higher than for stream water.
Stream water dissolved organic carbon (DOC) concentrations varied from 0.072 to 10.08
mg DOC L-1 from 110 data with a mean of 1.95 mg DOC L-1 and a standard deviation of 1.56
74
mg DOC L-1. For comparison, soil water DOC concentrations varied from 0.000 to 94.63 mg
DOC L-1 from 86 data with a mean of 11.43 mg DOC L-1 plus or minus an error of 1.72 and a
standard deviation of 19.38 mg DOC L-1. Lower DOC concentrations in the stream water were
similar to soil water DOC concentrations from deeper depths (1.75 m) probably due to the trend
of DOC decreasing with depth. On average from 86 data soil water DOC was an order of
magnitude larger than the mean DOC concentration from stream water samples. Large standard
deviations of soil water DOC data were likely due to the outlier-prone distribution.
Stream water ferrous iron concentrations varied from 0.007 to 5.90 mg Fe2+ L-1 from 122
data with a mean of 1.02 mg Fe2+ L-1 and a standard deviation of 1.21 mg Fe2+ L-1. In soil water,
ferrous iron concentrations varied from 0.000 to 6.674 mg Fe2+ L-1 from 78 data with a mean of
0.325 mg Fe2+ L-1 plus or minus an error of 0.317 mg Fe2+ L-1 and a standard deviation of 0.848
mg Fe2+ L-1. The range and mean of ferrous iron was higher in the stream water compared with
the soil water.
Stream water ferric iron concentrations varied from 0.011 to 2.34 mg Fe3+ L-1 from 122
data with a mean of 0.466 mg Fe3+ L-1 and a standard deviation of 0.438 mg Fe3+ L-1. For
comparison, soil water ferric iron concentrations varied from 0.052 to 1.666 mg Fe3+ L-1 from 78
data with a mean of 0.195 mg Fe3+ L-1 plus or minus an error of 0.738 mg Fe3+ L-1 and a standard
deviation of 0.286 mg Fe3+ L-1. The ranges of soil water and stream water were very similar, but
the mean soil water ferric iron concentration was lower than the mean stream water
concentration, which was probably due to the skew of the soil water ferric iron data toward lower
concentrations. Stream water data were used for comparison to soil water data from this study
(personal communication with Katherine Schroer, UGA student, EPA-ORD, 2010).
75
Table 4.6. Geochemical Analyses of Precipitation
Total Dissolved
+
Precipitation [NO3 -N] [NH4 ]
N
Organic C Sample
Location
(mg/L) (mg/L) (mg/L) (mg/L) Number
Date
10/15/2009 ARS-Main lot*
0.19 0.008 0.212
3.64
51
10/15/2009
0.19 0.008 0.211
2.58
52
10/15/2009
0.19 0.005 0.212
2.67
53
10/15/2009
0.19 0.010 0.202
3.02
54
10/26/2009 W2 Field Site
0.45 1.303
11.7
70
10/26/2009
0.44 1.367
12.8
71
10/27/2009
0.18 0.034
0.95
72
10/27/2009
0.18 0.035
0.60
73
10/27/2009
0.18 0.037
0.68
74
10/27/2009
0.18 0.030
0.56
75
11/12/2009
0.22 0.126
1.715
89
11/12/2009
0.23 0.124
1.120
90
11/12/2009
0.23 0.104
1.144
91
11/12/2009
0.22 0.127
1.074
92
11/19/2009
0.37 0.106
1.606
114
11/19/2009
0.37 0.180
4.752
115
* Multiple Precipitation Events Collected in One Sample Bottle at This Site
Nitrate-N, ammonium, and total nitrogen concentrations were relatively low in all of the
precipitation samples (Table 4.6). Dissolved organic carbon concentrations varied over a range
in precipitation samples from 0.56 to 12.8 mg L-1 (Table 4.6). Geochemical additions from
precipitation to soil water were relatively minor except for certain precipitation events in which
dissolved organic carbon added up to 12.8 mg L-1, which could be considered outlying data
because both large concentrations of dissolved organic carbon happened on October 26, 2009.
76
80
0
20
60
40
50
60
40
80
30
Precipitation (mm)
Nitrate Concentration (mg L -1 )
70
100
20
Precipitation
120
10
Nitrate at 0.35 m (L1)
Nitrate at 0.35 m (L1A)
11/13/2009
10/19/2009
9/24/2009
8/30/2009
8/5/2009
7/11/2009
6/16/2009
5/22/2009
4/27/2009
4/2/2009
140
3/8/2009
0
Nitrate at 0.5 m (L2)
Nitrate at 0.5 m (L2A)
Nitrate at 0.5 m (L2B)
Date (mm/dd/yyyy)
Figure 4.12. Lysimeters L1, L1A, L2, L2A, & L2B Nitrate Concentrations vs. Time
After a period of only infrequent precipitation of low total volume (less than 25 mm)
from May till August, 2009, nitrate concentrations began to increase significantly in the surface
lysimeters located at depths of 0.35 and 0.5 m, except for lysimeter L2A for reasons unknown
(Figure 4.12). During this same time of small, infrequent precipitation (less than 25 mm) from
May till August, 2009, nitrate concentrations also increased in lysimeter L3A at 1.25 m depth
(Figure 4.13). Most of the soil water sampled from lysimeters increased in nitrate concentrations
during this period after August, 2009, except for soil water sampled from lysimeters at the
deepest depth of 1.75 m (L4 and L4A) and lysimeters L2A at 0.5 m depth and L3 at 1.25 m
depth, which remained the same or decreased in value. Also, precipitation throughout the study
with highlights on dates for soil water sampling is shown in Appendix B.
77
40.00
0
20
30.00
40
25.00
60
20.00
80
15.00
Precipitation (mm)
100
10.00
120
5.00
Precipitation
Date (mm/dd/yyyy)
11/13/2009
10/19/2009
9/24/2009
8/30/2009
8/5/2009
7/11/2009
6/16/2009
5/22/2009
4/27/2009
140
4/2/2009
0.00
3/8/2009
Nitrate Concentration (mg L -1 )
35.00
Nitrate at 1.25 m (L3)
Nitrate at 1.25 m (L3A)
Nitrate at 1.75 m (L4)
Nitrate at 1.75 m (L4A)
Figure 4.13. Lysimeters L3, L3A, L4, & L4A Nitrate Concentrations vs. Time
78
4.6. Mineralization-Immobilization
The largest pool of nitrogen in the plant root zone is organic nitrogen in soil organic
matter, but organic nitrogen is not available to plants. This organic nitrogen can be released
through the process of mineralization to make plant available or mineral nitrogen.
Mineralization results in the release of ammonium (NH4+) or ammonia (NH3) by heterotrophic
soil microbes under aerobic and anaerobic conditions (McNeill and Unkovich, 2007).
Mineralization starts the nitrogen cycle in this study by transforming organic nitrogen
into ammonia by means of microbes in the upper-most soil layers, especially in the top 0.05 m of
soil where most of the dead and decaying plant (especially grasses, leaves, roots, and fallen tree
branches) and animal matter (mostly manure and urea) are located (Stevenson and Cole, 1999).
The mineralization process is evident as nitrate and ammonium were not added to the system
substantially in rain, but nitrate is often found in large concentrations at a depth of only 0.35 m.
Manure and urea from cow urine are the main sources of nitrogen to the system. As manure and
urea are decomposed by aerobic and anaerobic microorganisms in the upper-most soil layers,
organic nitrogen is transformed into ammonia, most of which is in turn transformed into nitrate
by the nitrification process. However, because mineralization is usually accompanied
simultaneously by immobilization, concentrations of ammonium and nitrate cannot be used to
calculate a mineralization rate nor can concentrations of nitrate be used to determine a
nitrification rate because these rates would not accurately quantify all of the transformations
(Stevenson and Cole, 1999).
The immobilization process is when ammonia is transformed back into organic nitrogen,
some of which is incorporated into soil microbes by microbial reproduction. The term
“immobilization” implies that the nitrogen forms are not available to plants. The immobilization
79
and mineralization processes are constantly transforming nitrogen between organic and inorganic
forms in the soil system. Immobilized nitrogen can be seen as part of the percentage of total
nitrogen which is not nitrate or ammonia or N2 gas. At a depth of 0.35 m, the percentages of
total nitrogen which exclude nitrate are 0%, 11.0%, 10.6%, 85.1%, and 56.5%. These
percentages represent other forms of total nitrogen such as ammonia and N2 gas, and organic
nitrogen, which excluding ammonia are forms of nitrogen that are difficult to quantify and were
not measured in this study. However, non-nitrate forms of nitrogen in general are expected to
increase with depth unless nitrogen gases escape the soil system into the atmosphere. This is
because in general denitrification increases with depth, and this general rule was observed in
most of the lysimeter soil water samples except for the soil water samples collected from the L3
lysimeter at 1.25 m depth.
At the L3 lysimeter, two more lysimeters were clustered to observe the consistently high
total nitrogen and nitrate-N concentrations of around 18 mg NO3-N L-1. These clustered
lysimeters are within one meter spatially of L3 and are L1A (0.35 m depth) and L2B (0.5 m
depth) lysimeters. Over several sampling events, high total nitrogen and nitrate nitrogen have
been observed in the shallower L1A and L2B as well. This area encompassing the eastern-most
part of the instrumented field site appears to have higher nitrogen concentrations, but specifically
this area has higher nitrate concentrations more than any other nitrogen species. This means this
area has higher mineralization accompanied by higher nitrification since ammonia concentrations
are some of the lowest average values compared with soil water samples from any other
lysimeters at the field site, even lysimeters in other spatial areas at the same depths as L1A and
L2B.
80
4.7. Nitrification
Nitrification is limited by soil temperature, soil pH, soil moisture, ammonia
concentration, bacteria with the metabolic ability, and concentrations of oxygen or other terminal
electron acceptors used as energy by the soil microbes (McNeill and Unkovich, 2007). Along
with ammonification, nitrification is considered the second half of mineralization and is a twostep biological process: first Nitrosomonas converts ammonium into nitrite, and second
Nitrobacter converts nitrite into nitrate (Stevenson and Cole, 1999). Only aerobic
microorganisms oxidize ammonia to nitrate in the nitrification process so zones without oxygen
inhibit the nitrification process and promote ammonia accumulation (Stevenson and Cole, 1999).
Nitrification slows below a soil pH of 5.5 (Bohn et al., 2001).
Total nitrogen represents all the inorganic and organic forms of nitrogen present in the
soil water. Percentages of nitrate in total nitrogen in each soil water sample from Lysimeter L1
at 0.35 m depth were 105.4%, 89.0%, 89.4%, 14.9%, and 43.5% for five samples with an
average percentage of 68.4%. In one soil water sample from Lysimeter L1 nitrate exceeded total
nitrogen (105.4% of total nitrogen), which was due to loss of total nitrogen from improper
storage procedures.
Percentages of nitrate in total nitrogen per soil water sample from
Lysimeter L1A at 0.35 m depth were 97.0%, 96.7%, and 98.7% with an average of 97.5%. On
average at the shallowest sampling depth, nitrate comprises 79.3% of total nitrogen, which was
averaged from two lysimeters, L1 and L1A, whereas ammonium represents a very small
percentage of the total nitrogen. Most of the ammonium is most likely being transformed into
nitrate through the nitrification process at the 0.35 m depth and at most of the sampled depths.
Nitrification can happen at any depth given the proper conditions (warm enough
temperatures, high enough soil moisture, etc.), but nitrification happens very readily in the upper
81
soil layers especially within the top 0.05 m of soil where most of the dead and decomposing
plant and animal matter is located (Stevenson and Cole, 1999). As soon as organic nitrogen is
converted to ammonium or ammonia by the process of mineralization, ammonia is transformed
by nitrifying bacteria.
Lysimeters L1A (0.35 m depth), L2B (0.5 m depth), and L3 (1.25 m depth) which appear
to be in an area of high nitrate and total nitrogen show trends of high nitrification because nitrate
makes up a large percentage of total nitrogen and ammonia makes up a very low percentage of
total nitrogen. Lysimeters L1A and L2B have some of the lowest ammonia concentrations over
several sampling events as well as the lowest ammonia average concentrations at sampling
depths of 0.35 and 0.5, respectively. Lysimeter L3 (1.25 m depth) has some of the higher
ammonia concentrations and higher average ammonia concentrations, but this is likely the results
of the product of denitrification. Piezometer 2 which is down-gradient from the L1A-L2B-L3
cluster has groundwater with nitrate concentrations at 3.45 and 3.81 mg NO3-N L-1 on two
separate sampling events, which implies denitrification in the soil below lysimeter L3 as well as
denitrification in the groundwater below the water table or dilution from groundwater.
4.8. Denitrification
Denitrification is limited by soil pH, soil temperature, soil moisture, bacteria with the
metabolic ability, nitrate concentration, and concentrations of O2 or other terminal electron
acceptors used in the denitrification processes as a source of energy (McNeill and Unkovich,
2007). When examining soil water samples with depth, high nitrate concentrations in near
surface soil water samples were never seen in the deepest soil water samples, which could imply
denitrification was lowering nitrate concentrations in soil water samples collected in the deeper
lysimeters. High nitrate and total nitrogen concentrations around 18 mg L-1 were observed in
82
soil water samples collected over several months from lysimeter L3 at 1.25 m depth, and soil
water samples from lysimeter L4 at 1.75 m depth, which was 7.70 m distance from L3,
consistently had concentrations of nitrate and total nitrogen around 4 – 5 mg L-1 over the same
time period, which demonstrates denitrification at least to some degree probably in the soil as
well as below the seasonally fluctuating water table between lysimeters L3 and L4, at depths
1.25 and 1.75 m, respectively. Denitrification increasing from around 0.5 m to 1.25 m depth
agrees with a study by Linden et al. (1984) in which denitrification increased five-fold in a
vadose zone under a corn crop from depths 0.6 m to 1.25 m.
Groundwater nitrate concentrations averaged from 5 different piezometers over three
sampling events (Table 4.5) showed an average nitrate concentration of 4.34 mg L-1, which
agreed well with nitrate concentrations measured from groundwater samples collected from
monitoring wells and the spring on average which was 4 – 5 mg L-1 (personal communication
with Dr. Caroline Stephens, USEPA). Therefore, as nitrate in soil water percolating downward
reaches a depth of 1.75 m, denitrification was most likely occurring that was lowering nitrate
concentrations especially in surface soils were microbial activities were highest.
Increased soil carbon contents can also promote denitrification such as in long-term
manure applications (Webster and Goulding, 1989). Grazing cows at W2 could then add to the
nitrate contamination and help relieve excessive nitrate by denitrification. Dissolved organic
carbon was measured during the study, and in a typical pattern in soils, dissolved organic carbon
decreased with depth. Dissolved organic carbon comes from decaying plant material, manure,
urea {(NH2)2CO}, and organic acids which can wash off trees and plants during rainfall, also
known as throughfall.
83
Table 4.7. Soil Water Average Concentration Values with Depth under Tree Canopy
[NO3--N] [NH4+]
Sample
Depth (m) Lysimeter (mg/L) (mg/L)
0.35
L1
20.42
0.074
0.50
L2
6.33
0.048
1.25
L3A
12.69
0.048
1.75
L4A
1.28
0.015
Dissolved
Urea Total N Organic
Fe2+
(mg/L) (mg/L) C (mg/L) (mg/L)
-0.011 13.488
52.485
1.085
-0.005
4.189
29.041
0.074
12.929
15.020
0.094
2.112
3.611
0.026
Fe3+
(Totl. Fe - Fe2+)
(mg/L)
0.301
0.071
0.178
0.093
The majority of high DOC concentrations were seen in the 0.35 m and, to a lesser extent,
the 0.5 m lysimeters, and average values for lysimeters under tree canopy were higher than
average values under open sky (Tables 4.7 and 4.8). Average dissolved organic carbon
concentration values at 0.35 m and 0.5 m depth under tree canopy were 52.485 and 29.041 mg
DOC L-1, respectively (Table 4.7). When compared to average DOC concentrations under open
sky at 0.35 m and 0.5 m depth (two lysimeters located at 0.5 m depth) that were 11.123, 4.237,
and 4.030 mg L-1, respectively (Table 4.8), the presence of more carbon from organic acids being
washed off plants or increased decaying organic matter under trees can be seen (Table 4.7).
More dissolved organic carbon in soils under tree canopy could mean more denitrification
occurring in soils under the trees.
Table 4.8. Soil Water Average Concentration Values with Depth under Open Sky
-
+
[NO3 -N] [NH4 ]
Sample
Depth (m) Lysimeter (mg/L) (mg/L)
0.35
L1A
18.36
0.060
0.50
L2A
2.48
0.343
0.50
L2B
42.87
0.019
1.25
L3
19.40
0.127
1.75
L4
3.73
0.045
Urea Total N
(mg/L) (mg/L)
24.158
3.891
50.039
-0.007 20.060
-0.008
4.439
Dissolved
Fe2+
Organic
C (mg/L) (mg/L)
11.123
0.281
4.237
0.164
4.030
0.094
1.013
0.182
0.855
0.597
Fe3+
(Totl. Fe - Fe2+)
(mg/L)
0.344
0.185
0.249
0.153
0.219
Trees next to the study site may have influenced denitrification around many of the
lysimeters in the soil. Most of the lysimeters at the study site are located within 20 m of the trees
which grow around the spring down-slope from the study site, except for lysimeters L1A, L2B,
and L3 positioned at a depths of 0.35, 0.5, and 1.25 m, respectively, which were located farthest
84
away from the riparian zone trees at an approximate distance of 15 to 20 m. Microbes tend to
work in a symbiotic relationship around roots. Meding et al. (2001) found up to four times
higher denitrification rates in riparian zones compared to upland zones under the same
background conditions.
4.9. Terminal Electron Acceptors and Oxidation-Reduction Reactions
The factors affecting microbes in the vadose zone are similar to the factors affecting
microbes everywhere: water, source of carbon, energy, terminal electron acceptors, and other
nutrients and environmental factors such as pH and temperature (Holden and Fierer, 2005).
Deep and near surface vadose zone sediments host microbes that are desiccation tolerant,
meaning that the microbes are frequently exposed to low water content and low water potential
(Kieft et al., 1993; Fierer et al., 2003a; Holden and Fierer, 2005). Water transports C and
microbes below the surface such that high recharge areas harbor different culturable bacteria
compared to low recharge areas (Brockman et al., 1992; Holden and Fierer, 2005), and
infiltrating water transports surface soil bacteria to deeper depths depending on the amount of
recharge an area receives, which implies areas of greater recharge have larger abundance of
bacteria at deeper depths in the vadose zone (Holden and Fierer, 2005). Oxygen is at
atmospheric levels at the surface of a soil but can decrease to about 20% at 5 m depth depending
on the soil (Wood and Petraitis, 1984; Holden and Fierer, 2005).
Iron is present in most rocks of Earth’s crust, and the Pacolet and Cecil soil series are
known to have large amounts of iron present as goethite and hematite clay minerals. Hematite is
responsible for the dominant red color of many Georgia/Piedmont soils. Oxidants should react
spontaneously with reductants of lower potential (pe), and oxidants react dominantly with
reductants which are present in highest concentrations (Washington et al., 2004). With
85
denitrification, oxidation-reduction reactions should lower oxygen or, in an anaerobic
environment, another terminal electron donor such as ferric iron should decrease (Endale at al.,
2003). However, neither a significant decrease in ferric iron nor an accompanying rise in ferrous
iron was seen in soil water collected from the deepest lysimeters at 1.75 m depth, which should
accompany denitrification since iron is most likely the reductant present in highest concentration
besides oxygen (Figures 4.10 and 4.11). Iron as a reductant likely dominates in areas in the
vadose zone such as clay lenses and saprolite zones where oxygen does not exist or is quickly
used up in by oxidation-reduction reactions. Ferrous and ferric iron varied at each sampling
depth over the same ranges and at the same sampling depth in different lysimeters only meters
apart with no clear trends such as increasing or decreasing with depth.
4.10. Effects of a Seasonally Elevated Water Table
One explanation for high nitrate/total nitrogen concentrations in near-surface soil water
samples that decreases significantly at a depth of 1.75 m to background concentrations is dilution
by seasonally high capillary fringe water or groundwater. The two lysimeters at 1.75 m depth
were able to collect very large amounts of soil water compared to the other lysimeters, usually at
least twice as much volume. The volume of water in the saturated zone is far greater than the
volume of water in the vadose zone, and this dilution in a greater amount of water below the
water table helps explain the low concentrations of groundwater compared to many nitrogen
concentrations measured from the lysimeter soil water samples. Also, large volumes of soil
water at the deepest sampling depth (especially lysimeter L4) could point toward a fluctuating
capillary fringe because nearby piezometers had water levels about 0.5 m below this 1.75 m
depth, water levels approximately around 2.25 m depth below the surface (Appendix A). Larger
volumes of water present in a capillary fringe could dilute soil water in proximity to the porous
86
ceramic cup of the deepest lysimeter. Also, as depth increases in the vadose zone usually clay
content also increases which means higher water contents at greater depths as clays take more
time to release soil water. Fluctuating seasonal depth to the water table in meters (Figure 4.14)
has a major influence on proximity of lysimeters at 1.75 m depth (L4 and L4A) to the capillary
fringe (if one is present) or the water table.
Figure 4.14. Seasonal Depth to Water Table Measured from All Field Site Piezometers
Figure 4.14 shows smallest minimum depth to water table and largest maximum depth to
water table from all depth to water table data measured in piezometers 1, 2, 3, 4, 5, 6, 7, 8, 9, 10,
and 12 (see Figure 4.5 for piezometer layout at field site). Another effect of a seasonally high
water table (seasonal minimum water table depth data in Figure 4.14) could be horizontal
movement of nitrate-rich soil water across the site in a capillary fringe as was noted in a study by
Abit et al. (2008). High total nitrogen after several months of drought-like conditions from June
87
till September could have brought on nitrogen flush conditions with significant September rains.
However, another explanation could be horizontal transport approximately 11 m by means of a
capillary fringe from the eastern-most side of the field site at L3 to the western-most side at L3A
at the same sampling depth of 1.25 m. These high nitrate concentrations were not seen at
lysimeter L4A at a lower sampling depth of 1.75 m located 3.75 m south of lysimeter L3A, but
the deepest lysimeters could have been diluted by groundwater due to high water table which
made the high capillary fringe possible (Figure 4.14 and Appendix A).
4.11. Evapotranspiration
Evapotranspiration can play an influential part in nitrogen concentrations. For a given
input of water and nitrogen flux, evapotranspiration may remove more water than nitrogen which
would increase nitrogen concentrations in soil water which did not transpire into plants or
evaporate into the atmosphere. The opposite could occur in winter or spring months, where
evapotranspiration occurs at a lower rate so soil water could become relatively, seasonally
diluted from this process compared to summer soil water (Green et al., 2008).
4.12. Plant Uptake
In acid soils, plants that can take up ammonia have an advantage over plants that require
nitrate because nitrification is slow below pH 5.5 (Bohn et al., 2001). Other plants that can only
take up nitrate can transform ammonia to nitrate using microbes that live in close proximity to
plant roots in the rhizosphere, although this process is more energy intensive. Some plants can
take up nitrogen as organic nitrogen, which is advantageous when organic matter is in high
supply in the root zone in certain environments like bogs, some forests, or a pasture with grazing
animals.
88
4.13. Non-Biological Nitrogen Transformations
Chemical transformations of nitrogen include ammonium fixation on the surfaces of
clays, fixation of ammonia on soil organic matter, and nitrite organic matter reactions.
Ammonium is a cation that is attracted to negatively-charged sites in the expandable lattice of
clay particles. Certain clays, such as vermiculite, illite, and montmorillonite, are more likely to
fix ammonium due to isomorphic substitution of Al for Si in the tetrahedral layers, which is the
source of the negative charge. Ammonium becomes fixed in the voids created between the
tetrahedral layers of the clay particles. In soils dominated by kaolinite, such as the soils at the
study site, almost no fixation of ammonium occurs. Also, highly acidic soils (< 5.5) generally
fix little ammonium, and approximately half of the soils tested in this study would be considered
highly acidic (Stevenson and Cole, 1999).
Ammonia fixation by organic matter was facilitated by oxidation, uptake of O2, and
occurred more readily in alkaline soils. Application of aqueous or anhydrous ammonia alkaline
fertilizers could result in considerable fixation. Ammonia fixation rates are very comparable to
the organic matter content of a soil. Also, studies indicated that fixed ammonia was not available
to plants (Stevenson and Cole, 1999). Dissolved organic carbon concentrations measured up to
94.53 mg L-1 at 1.25 m depth indicate wide-spread presence of organic matter in the soil water at
this site, although the trend in general was decreasing dissolved organic carbon with depth.
However, with the pH of the acidic soils present in this study averaging around 5.5, ammonia
fixation to organic matter was most likely very low.
89
Reaction of nitrite with lignin, fluvic, and humic acids produces nitrogen gases.
However, nitrite is very fleeting in most soil conditions and is thought to quickly nitrify or
denitrify. Reactions of nitrite with organic acids occur in trace amounts in normal soil conditions
(Stevenson and Cole, 1999).
4.14. Wetting and Drying Cycles
North Georgia was in a drought during this study from 2007 till the fall of 2009. Winter
and spring rain events can act as wetting cycles after periods of drought or drying, which bring
flushes of nitrogen. Many soil water sampling events followed winter and spring rain events in
this study. For example, rain events preceding soil water sampling on the 23 of September,
2009, were the first significant precipitation greater than 2 cm rainfall since early June at the W2
site.
In soil water collected from the 0.35 m and 0.5 m depth lysimeters over several months,
total nitrogen concentrations decreased with increasing time due to flushing from the numerous
rewetting cycles caused by winter/spring precipitation events. This trend of decreasing total
nitrogen concentrations over time continued until evapotranspiration increased at the beginning
of summer, and soil moisture decreased rapidly until air entry matric pressure was reached
making soil water collections impossible.
After a drought in the summer that lasted from the start of June until mid-September, soil
water concentrations in several lysimeters were found to have high total nitrogen that could have
been due to a nutrient/nitrogen flush accompanying the significant September and October rains.
High total nitrogen concentrations around 15 mg N L-1 or higher were measured in soil water
samples from L1A, L2B, L3, and L3A during the September 23, 2009, and the October 16, 2009,
90
sampling events. Total nitrogen reached 75 and 76 mg N L-1 in two soil water samples taken
from lysimeter L2B on October 16, 2009.
Table 4.9. Soil Water Average Concentration Values with Depth in High Nitrogen Area
Dissolved
[NO3--N] [NH4+] Urea Total N Organic
Sample
Fe2+
Depth (m) Lysimeter (mg/L) (mg/L) (mg/L) (mg/L) C (mg/L) (mg/L)
0.35
L1A
18.36 0.060
24.158
11.123 0.281
0.50
L2B
42.87 0.019
50.039
4.030 0.094
1.25
L3
19.40 0.127 -0.007 20.060
1.013 0.182
Fe3+
(Totl. Fe - Fe2+)
(mg/L)
0.344
0.249
0.153
From the first and second rain events following the summer drought, nitrate
concentrations of soil water were some of the highest concentrations measured during the length
of the study at 20 mg NO3--N L-1 or more in lysimeters L1A, L2B, and L3, which are all located
in the high nitrogen area of the field site at the East end (Table 4.9). Also, from the first and
second rain events following summer drought, nitrate concentrations in soil water at the West
end of the field site were the highest recorded for the L3A lysimeter at 14 mg NO3--N L-1 or
more at a depth of 1.25 m. These high nitrate concentrations, especially in the L3A lysimeter
since the concentrations are the highest for this lysimeter and this area of the field site during the
length of the study, could represent a flush through the vadose zone after a period of drought, but
another lysimeter in this West side of the field site 4 m away at a depth of 1.75 m had low soil
water concentrations for these two storms between 3.11 and 0.28 mg NO3--N L-1. However,
another interpretation could be that throughfall has increased the nitrate of the soil water for the
L3A lysimeter for these two storm events.
In surface soils, drying-rewetting cycles stress soil microbes (Fierer et al., 2003b). This
stress is shown in lower C and N mineralization rates compared with unstressed subsurface
microbes which occupy the same substrate (Rovira and Vallejo, 1997). However, according to
91
Rovira and Vallejo (1997) in surface soils, high organic matter helps reduce the stress of low
water on soil microbe activity (Holden and Fierer, 2005).
4.15. Soil Bacteria Population Cycles and Nitrogen Transformations
Another hypothesis as to why nitrate-N and total N concentrations lessen in soil water
with depth is the natural attenuation resulting from seasonal fluctuations of the abundance of soil
microbes. As precipitation lessens and evapotranspiration increases over the constantly warm,
sunny, summer months, the nutrient delivery, especially oxygen, dissolved organic carbon,
nitrate, Fe2+, and other terminal electron acceptors, lessens as well until the soil dries and kills
the populations of microbes. Conversely, when precipitation is high and evapotranspiration is
low, during the winter and spring months, microbial activity is highest and nitrogen
transformations, especially denitrification and, to a lesser extent, nitrification, are being carried
out in the soil of the vadose zone at the W2 site.
Soil microbial biomass is generally highest near the surface and decreases rapidly with
depth for all types of bacteria abundance counting methods (Balkwill et al., 1998; Taylor et al.,
2002) mostly due to a decrease in organic C concentrations with depth (Holden and Fierer,
2005). Also, in shallow vadose zones, the majority of total microbial biomass occurs in the
highest cell densities in surface soils and in the capillary fringe (Holden and Fierer, 2005). Nonbacterial biomass, which includes plants, algae, fungi, and diatoms, decreases more rapidly with
depth than bacterial biomass according to data that used the phospholipid fatty acids (PFLA)
total biomass counting method. Direct counts of fungal populations also decline more rapidly
with depth than direct bacterial counts (Taylor et al., 2002; Holden and Fierer, 2005). The
abundance of microbes in the vadose zone varies more within a single soil profile than
abundance of microbes varies in different soils (Fritze et al., 2000; Holden and Fierer, 2005).
92
One study showed a correlation between microbial abundance and soil texture (Konopka
and Turco, 1991); however, in another study microbial abundance did not correlate well with
water content or water potential (Balkwill et al., 1998), which is related to soil texture. A strong
correlation exists between microbial biomass and soil C content over a range of soils with
different textures and for many different counting methods. As organic C decreases rapidly with
depth so do microbes which require organic C for growth (Holden and Fierer, 2005). Bacteria,
the most abundant microbes in the vadose zone, can colonize the air-water interface (Wan et al.,
1994), live freely in water, or exist as biofilms on surfaces (Oades, 1984), although Else et al.
(2003) dispute the ability of microbes to exist as biofilms on vadose zone materials (Holden and
Fierer, 2005).
The factors that aid in microbial nitrogen transformations are clearly at the highest rates
during the winter and spring months in Georgia. The winter likely experiences the most
microbial nitrogen transformations of any season due to the least amount of evapotranspiration,
the most infiltrating soil water, the highest delivery of nutrients to the microbes with the
infiltrating water, and the decay of plant matter which in turn delivers more nitrogen back to the
soil.
4.16. Excessive Nitrogen Fertilizer Application in W2
As previously discussed, fertilizer thresholds have been defined many different times in
the literature. A fertilizer threshold from a study on an arable clay soil in Sweden that spanned
10 years found moderate leaching of nitrate up to a rate application of 100 kg N ha-1 yr-1
(Bergstrom and Brink, 1986). Due to the length of the Bergstrom and Brink (1986) study, this
threshold was examined for the present study. As can be seen from Table 4.2, the years during
which W2 exceeded this nitrogen fertilizer application threshold were: 1992-1993, 1996, 199893
2000, 2002-2003, and 2005-2006. These years which exceed this fertilizer rate threshold are
from the total N applied in a year which often involved split applications. Also, these years of
excessive application of nitrogen can and most likely did accumulate inorganic nitrogen for
leaching years after the fertilizers were applied.
In certain years, the USDA-ARS was budget-limited in fertilizer application to W2, and
this practice resulted in no fertilizer application during 2008. Other years W2 was fertilized up
to three times during the year. The fertilizer applications did not account for cow manure and/or
urine which were applied naturally to the field whenever cows were allowed to graze the
watershed so these additions add further nitrogen to the system. The years with the highest rates
of fertilization were: 1999 and 2005 (tied 1st with 190 kg N ha-1 yr-1), 1993 (2nd with 162 kg N
ha-1 yr-1), and 1996 (3rd with 157 kg N ha-1 yr-1), and these fertilizer application rates are well
above the threshold application rate of 100 kg N ha-1 yr-1 described by Bergstrom and Brink
(1986). However, the study by Bergstrom and Brink (1986) in Sweden was on arable land
compared to W2 which has been a pasture for decades so the rate of 100 kg N ha-1 yr-1 may not
be the right threshold rate for W2, but W2 fertilizing rates should still be examined due to
occasional excessive nitrate-N in soil water. Another study recommended a threshold rate for a
single application at or below 120 kg N ha-1 per application in areas susceptible to leaching
(Nangia et al., 2008). Only once since 1992 did a single application of nitrogen fertilizer
approach 120 kg N ha-1: March 5, 2005 at 114 kg N ha-1. The threshold rate of 120 kg N ha-1
described for areas susceptible to leaching found that below this threshold nitrate leaching was
moderate, but above this threshold application rate, nitrate leaching rates increased steeply
(Nangia et al., 2008). However, the Florida Cooperative Extension Service recommends single
nitrogen fertilizer applications of 165 - 220 kg N ha-1 (Nangia et al., 2008).
94
In Georgia over a growing season, for top quality Coastal Bermuda (good beef
production) fertilizer rates are 224 kg N ha-1 (200 lbs N ac-1) as top dressing in split applications
(Johnson, 1967). Also, hay production requires a fertilizer rate of 336 to 448 kg N ha-1 (300 to
400 lbs N ac-1) (Johnson, 1967). These rates necessary for production exceed the Bergstrom and
Brink fertilizer threshold rate of 100 kg N ha-1 yr-1 (1986), which likely means fertilizer threshold
rates cannot be universally applied to watersheds.
The instrumented field site lies outside of a fence which separates the grazing and
fertilizable portion of the watershed from the spring area at the bottom of the watershed. The
fence was put up by the USDA-ARS JPC in 1999, and excluded all of the field site but
lysimeters L1A, L2B, and L3A at depths 0.35, 0.5, and 1.25 m. This group of three lysimeters is
also the area in which nitrate-N and total N were consistently high (above 10 mg L-1). Upon
request, the USDA-ARS JPC moved the fence back further in 2006 to help avoid damage to field
instruments, after which time the field site was entirely outside the fence.
4.17. 15N End-Member Analyses: Manure or Fertilizer Nitrogen Source
Soil samples were analyzed in nitrogen stable isotope end-member analyses from two
different depths augered from the same hole to see if the source of nitrogen could be determined
from the concentration of 15N relative to air. The area on the East side of the field site around
lysimeters L1A, L2B, and L3 where nitrogen concentrations are the highest was targeted for soil
samples since this area may act as a source of nitrogen to other parts of the vadose zone across
this part of the field site. Also, the East side of the field site has always produced high nitrogen
concentrations so the 15N signal would likely be strongest in a soil with high total nitrogen and
nitrate concentrations.
95
Chemical fertilizers were applied September 10, 2009, to the pasture which was 15 days
before the soil samples were collected from an area of the field site 15 m down-gradient from the
pasture portion of W2. Also, fertilizers have not been applied in W2 since August 7, 2007, until
the application on September 10, 2009. The chemical fertilizers applied to Watershed 2 were a
combination of ½ of the N-P-K (25-0-0) chemical fertilizer and ½ of the N-P-K (46-0-0)
chemical fertilizer measured out by fertilizer weight percentages. Also, the fertilizer mixture
applied in 2009 had a product called Nutrisphere (S.F.P., Leawood, KS, USA) applied to prevent
ammonia volatilization losses. Nutrisphere was reported to prevent ammonia volatilization and
kept the nitrogen in the ammonium/ammonia phase longer so that maximum plant uptake was
achieved. The current position of the cattle fence in W2 that has separated the spring at the
bottom of the watershed from the grazing portion has been in place since 2006. The fence has
been outside of most of the field site since 1999 except for the high nitrogen portion of the field
site where lysimeters L1A, L2B, and L3 are located at 0.35, 0.5, and 1.25 m depth. The portion
of the field site where soil samples were removed from for the stable nitrogen isotopic analysis
was only fertilized in 2006, 2007, and 2009.
96
Table 4.10.
15
N Isotope Analyses on Nitrogen Source in Field Site Vadose Zone Soils
Sample
Weight (mg) δ15N vs. air
Atom % 15N
Date
Nitrogen Source
9/11/2009
Cow Manure
0.748
4.35
0.367809
9/15/2009
Chemical Fertilizer
(N-P-K) = (25-0-0)
0.512
-1.33
0.365986
9/15/2009
Chemical Fertilizer
(N-P-K) = (46-0-0)
0.524
-0.96
0.366123
Sample
Weight (mg) δ15N vs. air
Atom % 15N
Date
Soil Sample
9/25/2009
0.5 m depth soil
35.730
4.54
0.368129
9/25/2009
0.5 m depth soil
32.314
4.85
0.368244
9/25/2009
9/25/2009
2.0 m depth soil
2.0 m depth soil
52.243
59.229
-0.55
-0.95
0.366273
0.366124
Date
Reference Sample
Bovine Liver
Bovine Liver
Bovine Liver
Bovine Liver
Sample
Weight (mg) δ15N vs. air
2-2.5
7.52
2-2.5
7.36
2-2.5
7.58
2-2.5
7.53
Atom % 15N
0.369217
0.369158
0.369238
0.369223
Nitrogen isotopic analyses showed that the manure δ15N signature vs. air was 4.35 and
the 0.5 m depth soil samples δ15N signatures vs. air were 4.54 and 4.85. The chemical fertilizer
with an N-P-K of 25-0-0 and the chemical fertilizer with an N-P-K of 46-0-0 had δ15N signatures
vs. air of -1.33 and -0.96, respectively, and the 2.0 m depth soil samples had δ15N signatures vs.
air of -0.55 and -0.95 (Table 4.10). The 0.5 m shallow soils were very similar to the δ15N
signature of manure. The 2.0 m deeper soils were very similar to the δ15N signature of both of
the chemical fertilizers.
The results showed that at the time of sampling the shallow 0.5 m soils probably had a
manure nitrogen source, whereas the deeper soils around a depth of 2.0 m likely originated from
a chemical fertilizer nitrogen source (Table 4.10). These results could imply influence of a
capillary fringe at 2 m depth close to the lysimeter cluster of L1A-L2B-L3 and piezometer 5
because the δ15N signatures of the chemical fertilizer were transported to this deep depth without
97
influencing the δ15N signatures at the shallower depth. Due to negative pressure in a capillary
fringe, some groundwater could have been pulled up to a depth of 2 m which could have had
similar δ15N signatures as the chemical fertilizer if the fertilizer had already infiltrated into the
groundwater below the soil sampling site before the soils were sampled.
98
CHAPTER 5
CONCLUSIONS
5.1. Summary
In this study, soil water potential was measured and soil water samples were collected
and analyzed for nitrate, ammonia, urea, total nitrogen as well as dissolved organic carbon and
ferric and ferrous iron at different depths in the vadose zone. Geochemical trends were presented
and interpreted by depth and across time in the vadose zone. Groundwater and precipitation
samples were compared to soil water samples in order to provide background concentrations
from groundwater sample averages and to provide natural geochemical additions to the surface
from precipitation. The regional water table was also observed at the field site from a nest of
eleven piezometers over several months, and TDR data was also collected over several months to
measure soil moisture of the vadose zone. Volumetric water profiles were made from TDR data
measured in the vadose zone soils of the field site.
Nitrogen transformations are difficult to measure directly in the vadose zone, and
attempts in this study were spread across an area approximately 300 m2 in size (20 m NorthSouth by 15 m East-West) at four different depths in the vadose zone and in groundwater
samples collected from piezometers. The transformation processes observed in the data obtained
in this study area included mineralization, immobilization, ammonification, nitrification, and
denitrification to a certain extent. Studies involving the nitrogen cycle cannot often interpret all
of the nitrogen processes going on in an area, but only speculate as many transformation
processes overlap over and over again through time.
99
Nitrate-nitrogen concentrations observed in this study were at times very high in the soil
water (< 64 mg NO3N L-1) at lysimeter L2B at 0.5 m depth. However, groundwater leaving the
watershed through the spring and downstream of the spring in a pond often showed low
concentrations (< 6 mg NO3--N L-1) which implies denitrification at some point in the system
whether above or below the water table, in the watershed, in the stream, or in the pond. The
watershed and resulting stream and pond are not overly contaminated with nitrate-nitrogen, and
the watershed system uses much of the nitrogen applied as manure and chemical fertilizer.
Watersheds often need monitoring for decades to keep fertilization and grazing in check so that
contamination problems do not occur in the future.
Factors affecting nitrogen transformations in the vadose zone soils at the field site were
surprising. Throughfall precipitation brought increased dissolved organic carbon and possibly
increased nitrogen to lysimeters located underneath the tree canopy, which influenced
denitrification. Oxidation-reduction patterns believed to be occurring in the soils were not
observed with depth as iron concentration of the soil water samples showed no clear pattern with
depth or through time. Flushing events accompanying wetting-drying cycles were observed after
a period of no precipitation from June, 2009, till September, 2009, and these flushing events
brought surges of high nitrogen, especially nitrate-N, through the vadose zone soils even to
depths as deep as 1.25 m. These flushing events produced the highest nitrogen concentrations
observed in the soil water during the length of the study.
Isotopic analyses, although sample limited due to funding and time, demonstrated two
separate sources of nitrogen to the vadose zone soil samples from one area of the field site: a
manure source in the shallow soils at a depth of 0.5 m and a chemical fertilizer source in soils at
a depth of 2.0 m. The manure in the study area is from years past as this study area has been
100
fenced off from the pasture and grazing cows for 3 years. Chemical fertilizers have also
accumulated in the soils over many years as the nitrogen-rich fertilizers have been applied in this
watershed since 1992. With more isotope work, specifically with δ18O analyses of soil water and
groundwater, the exact extent of denitrification could be determined as 18O and 15N both
fractionate when denitrification occurs. However, this study was time and budget limited, and
these isotope analyses must be left for future work.
5.2. Suggestions of Future Work
As mentioned above, an extensive stable isotopic study soil, soil water, and groundwater
samples from W2 could show the extent of denitrification as well as nitrogen sources all over the
watershed.
18
O and 15N stable isotopes signatures could be analyzed in groundwater samples
collected from across the watershed. Lysimeters could be constructed to help facilitate more
groundwater samples at different locations across W2 like lysimeters L1A, L2B, L3A, and L4A
which were built in this study (only longer). Isotope lab analyses are relatively low cost for
UGA students at the UGA Institute of Ecology Analytical Chemistry Lab compared with other
labs and non-student prices.
Other work should involve clustering of lysimeters around established existing lysimeters
in the study area. New areas of interest could be explored for installation of lysimeters each
testing new theories of nitrogen transformations, like surface channels or small depressions. The
emphasis of clustering should be on shallow and deep lysimeter placement together in one area
separated by little more than 1 m. With shallow and deep lysimeters clustered together, more
can be determined about nitrogen transformations with depth than from speculation across
several meters spatially. Lysimeters could be installed right next to the spring and spring-fed
creek which leaves W2 to examine denitrification before groundwater enters the stream as
101
stream recharge. Lysimeters could be installed under tree canopies to examine effects of
throughfall on nitrogen in the forest soils.
Other factors could be observed to see if there is an effect on nitrogen in soil water such
as alternative fertilizer applications like poultry litter, pig manure, or other animal feces could be
examined for their effect on nitrogen transformations. This could mean examination of nitrogen
transformations in another watershed where alternative fertilizers were in use.
A similar site at another USDA-ARS watershed could be instrumented in a similar
fashion to this study to compare side-by-side. Improvements on clustering of lysimeters could
be implemented. Perhaps more shallow depths could be examined for instrumentation of
lysimeters in the root-zone. Soil pits could be dug and lysimeters and tensiometers could be
installed horizontally to examine any differences to the field site in W2.
102
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114
APPENDIX A. DEPTH TO WATER TABLE DATA FOR PIEZOMETERS
SURROUNDING W2 FIELD SITE
Groundwater levels from top of casing (t.o.c.) to the water table in meters
All piezometers are located around the flume and around my 3 tensiometers and 9 lysimeters
Date
1/7/2008
1/24/2008
3/3/2008
3/11/2008
3/21/2008
3/28/2008
4/4/2008
4/10/2008
4/18/2008
4/25/2008
5/5/2008
5/9/2008
5/20/2008
5/30/2008
6/6/2008
6/12/2008
6/20/2008
7/3/2008
7/11/2008
7/24/2008
8/1/2008
8/29/2008
9/4/2008
9/12/2008
9/18/2008
9/29/2008
10/9/2008
10/31/2008
11/7/2008
11/17/2008
1/5/2009
1/23/2009
1
2.63
2.37
2.51
2.43
2.49
2.52
2.51
2.49
2.41
2.55
2.64
2.68
2.76
2.88
2.99
3.09
3.18
3.25
3.25
3.33
3.42
3.44
3.44
3.55
3.56
3.63
3.26
3.46
3.58
3.57
3.27
3.09
2
2.06
1.89
1.85
1.86
1.86
1.90
1.92
1.94
1.94
2.01
2.13
2.21
2.27
2.39
2.55
2.67
2.85
3.03
2.96
3.20
3.27
3.28
3.24
3.41
3.47
3.55
3.17
3.41
3.47
3.45
2.90
2.60
Piezometers
3
4
5
2.48 1.97
2.31 1.94
2.22 1.94
2.17 1.92
2.22 1.93
2.24 1.96
2.27 1.98
2.28 1.99
2.28 1.99
2.36 2.04
2.42 2.10
2.44 2.15
2.50 2.23
2.60 2.36
2.71 2.48
2.80 2.58
2.90 2.72
3.02 2.91
3.03 2.92
3.12 3.05
3.17 3.10
3.23 3.18
3.23 3.07
3.28 3.15
3.31 3.24
3.38 3.33
3.10 3.33
3.37 3.15
3.39 3.20
3.41 3.26
3.14 2.73
1.88 2.87 2.39
6
7
2.06
2.10
2.16
2.22
2.34
2.42
2.51
2.64
2.59
2.72
2.74
2.77
2.75
2.84
2.89
2.98
2.61
2.88
2.91
2.92
2.60
2.39
2.35
2.25
2.19
2.23
2.27
2.30
2.32
2.29
2.37
2.42
2.44
2.49
2.57
2.64
2.68
2.76
2.89
2.92
3.01
3.03
3.11
3.10
3.15
3.18
3.25
3.00
3.25
3.26
3.29
3.02
2.63
115
8
1.85
1.94
1.99
2.02
2.10
2.18
2.27
2.34
2.41
2.52
2.46
2.55
2.57
2.61
2.60
2.62
2.65
2.72
2.67
2.64
2.73
2.73
2.32
2.26
9
10
12
1.75
1.72
1.87
1.97
2.08
2.14
2.24
2.35
2.32
2.43
2.44
2.47
2.45
2.54
2.55
2.64
2.28
2.57
2.61
2.62
2.36
1.92
2.05
1.99
1.96
2.02
2.06
2.06
2.06
2.06
2.09
2.11
2.13
2.17
2.20
2.26
2.29
2.37
2.47
2.49
2.59
2.57
2.67
2.66
2.69
2.71
2.78
2.52
2.79
2.82
2.85
2.64
2.40
2.45
2.11
2.20
2.28
2.32
2.43
2.53
2.59
2.59
2.59
2.61
2.57
2.64
2.68
2.75
2.39
2.70
2.73
2.74
2.50
2.30
Data Continued
Date
2/2/2009
2/9/2009
2/20/2009
2/26/2009
3/9/2009
3/19/2009
3/26/2009
4/3/2009
4/9/2009
4/20/2009
5/7/2009
6/10/2009
1
3.15
3.19
3.08
3.17
2.51
2.40
2.51
1.88
2.31
2.30
2.31
2.48
2
2.70
2.75
2.69
2.74
1.84
1.83
1.84
1.77
1.83
1.83
1.84
1.85
3
1.95
1.98
1.93
1.97
1.47
1.41
1.47
1.23
1.43
1.42
1.45
1.52
Piezometers
4
5
2.93 2.45
2.97 2.51
2.84 2.55
2.97 2.56
2.22 1.91
2.09 1.91
2.22 1.92
1.48 1.78
2.00 1.90
2.01 1.90
2.03 1.91
2.16 1.95
6
2.46
2.48
2.46
2.49
1.70
1.60
1.69
1.40
1.59
1.60
1.63
1.70
116
7
2.82
2.88
2.81
2.89
2.15
2.16
2.19
1.67
1.98
1.99
2.03
2.13
8
2.30
2.34
2.33
2.42
1.72
1.65
1.73
1.42
1.66
1.66
1.72
1.78
9
2.05
2.13
2.17
2.22
1.48
1.41
1.51
1.09
1.41
1.41
1.45
1.55
10
2.44
2.48
2.47
2.50
1.99
1.91
2.02
1.68
1.90
1.90
1.96
2.04
12
2.37
2.40
2.36
2.41
1.74
1.65
1.76
1.24
1.64
1.65
1.68
1.80
APPENDIX B. TDR & PRECIPITATION DATA FOR W2 FIELD SITE
TDR Probe Location Notes:
Probes 1, 2, 3, and 4 measure discretely along sections of depth listed
TDR Probe 1 is located between piezometer # 3 and the spring
TDR Probe 2 is located between the spring and piezometer # 1
TDR Probe 3 is located closest to lysimeters L1 & L2 (at 0.35 & 0.5 m depth, respectively)
TDR Probe 4 is located closest to piezometer # 6
Bad data not represented (blank)
TDR Probes (data in % moisture)
Depth (m)
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
1
23.4
26.6
18.7
22.5
23.8
18.7
23.1
25.6
19.0
22.5
24.0
17.8
21.1
25.3
2
22.1
28.1
32.3
40.4
41.8
21.3
26.3
30.9
37.9
40.7
22.1
28.7
32.0
39.5
41.3
16.9
26.6
29.5
38.8
39.9
22.4
27.8
32.6
40.0
42.0
3
33.4
21.2
Date
4 Sampled:
25.6
1/6/09
23.1
2/9/09
24.7
2/20/09
24.0
2/26/09
24.6
3/9/09
25.0
18.0
30.3
18.8
26.2
19.1
52.4
28.8
19.6
44.5
117
Avg.
(% Moisture)
27.8
25.8
28.2
30.8
41.8
21.7
24.4
25.9
28.0
40.7
23.7
25.9
27.4
29.2
41.3
20.7
24.6
25.8
29.0
46.2
23.0
24.5
27.5
29.8
43.3
TDR Probes (data in % moisture)
Depth (m)
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
1
22.7
20.8
26.1
20.1
24.0
24.3
23.9
25.8
26.9
21.0
19.6
26.1
23.3
25.8
27.8
16.6
15.6
17.9
2
23.3
28.1
32.0
40.0
42.3
23.6
29.2
33.0
40.8
41.8
24.5
30.1
33.0
40.2
42.1
21.6
27.2
31.7
38.5
42.0
26.2
29.5
33.0
41.5
43.0
20.1
28.4
25.5
32.7
34.7
3
27.6
21.2
24.7
29.7
18.9
39.9
30.8
23.2
26.3
27.9
20.7
42.3
33.4
24.8
23.7
30.3
23.4
42.6
118
Date
Avg.
(% Moisture)
4 Sampled:
24.5
24.5
24.9
3/19/09
27.7
30.6
33.5
24.5
26.6
24.1
3/26/09
27.1
29.9
40.9
26.4
28.0
26.6
4/3/09
28.8
31.7
34.2
23.5
23.4
24.6
4/9/09
27.5
29.6
42.2
27.6
27.7
27.1
4/14/09
29.3
33.2
33.4
22.3
22.0
20.9
5/4/09
21.4
28.1
38.7
Depth (m)
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
0 - 0.15
0.15 - 0.30
0.30 - 0.60
0.60 - 0.90
0.90 - 1.20
TDR Probes (data in % moisture)
1
2
3
15.5
16.9
30.5
15.6
22.3
17.2
22.7
30.9
21.2
33.7
41.4
12.3
11.1
18.7
12.9
13.2
14.1
17.6
27.3
15.2
27.2
41.3
Average TDR Soil Moisture
0 - 0.15
19.1
0.15 - 0.30
21.0
0.30 - 0.60
23.5
0.60 - 0.90
0.90 - 1.20
Values
20.9
26.6
29.7
37.6
39.4
Avg.
Date
4 Sampled: (% Moisture)
21.0
19.0
22.7
5/11/09
20.9
26.1
37.6
14.0
13.1
15.4
6/10/09
15.7
21.3
34.3
28.7
23.7
20.4
37.9
119
Averaged
from all
data points
22.9
23.5
25.5
28.9
38.6
Day March
1
2
3
4
5
6
7
8
9
10
0
11
0
12
0
13
0
14
9.9
15
35.8
16
9.6
17
0
18
0
19
0
20
0
21
0
22
0
23
0
24
0.3
25
1.5
26
13.5
27
34.3
28
38.6
29
0
30
0
31
0.8
2009 Precipitation (mm)
(Hogenboom, 2010)
April May June July August September October November
13.2
2.3
0
0
5.6
0
0
0.3
41.7 13.5
0
0
0.8
0
0
0
0.5
1
0
0
0
0
0
0
0 13.2 23.6
0
0
0
8.1
0
0
5.8
3.8
5.6
0
0
31.2
0
0.3
5.8
0
0
0
0
0
0
0 18.3
0 21.1
0
0
5.1
0
0
0
0
0
0
0
0
0
0
0
0
0
0
3
0
0
6.6
0
0.3
0
0
0
0
81
0
8.1
0.3
0
29.5
0
0
22.6
0
0
1
5.1
5.1
0
102.1
0
25.6
0
0
2.8
0
0
0
0
24.6
0
0
0
0
0
22.4
0
0
0
0
0
10.9
18.3
2.3
0
0
8.4
0
0
0
18.5
0
0
0
5.1
4.3 10.9
10.9
63.2
4.1
6.1
0
0
9.9
0
0
17
0
0.5
5.1
0
0
0
0
50.8
0
0.5
2.8
0
0
0
3.8
23.4
0
0
0
0
0
5.1
62.7
0
0
1
0.5
0
1.3
0
0
0
2
0
0
0
0
3.3
0
1.8
0
0
0
0
0.3
0
0.5
0
0
0
0
0
0
0.8
0
0
0
32.8
0
0
0
0
0
0
0
39.4
0
0
0
0.3
17.3
0
0
0
0
0
0
0.3
0
0
0
0
0 14.5
8.9
0
1.8
0
4.1
7.9
5.1
= Sampling date when soil water samples were removed from lysimeters
120
TDR Moisture vs. Depth vs. Time
50.0
Soil Moisture (%)
45.0
40.0
35.0
30.0
45.0-50.0
25.0
40.0-45.0
20.0
35.0-40.0
15.0
30.0-35.0
25.0-30.0
10.0
20.0-25.0
15.0-20.0
10.0-15.0
Depth (m)
Date (mm/dd/yyyy)
121
APPENDIX C. WIRING DIAGRAM FOR TENSIOMETERS
The three tensiometers are emplaced around the circumference of the original 3 m circle study
area at 0.5, 1.0, and 1.5 m depths
0.5 m depth Tensiometer wiring diagram:
Cell
wire color location in CR23X datalogger Pressure Transducer
Green
6 High
Upper
White
6 Low
"White T" (0-15 psig)
Cell
(blueRed
Excitation 1
marked wire) Black
Ground
Lower
Green
5 High
White
5 Low
"White X" (0-15 psig)
Cell
(non-marked Red
Excitation 1
wire)
Black
Ground
1.0 m depth Tensiometer wiring diagram:
wire color location in CR23X datalogger Pressure Transducer
Upper
Green
2 High
White
2 Low
"Newest 0-15 psig"
Cell
(blueRed
Excitation 3
marked wire) Black
Ground
Green
1 High
Lower
White
1 Low
"Newest 0-30 psig"
Cell
(non-marked Red
Excitation 4
wire)
Black
Ground
1.5 m depth Tensiometer wiring diagram:
wire color location in CR23X datalogger Pressure Transducer
Green
4 High
Upper
White
4 Low
"Red 2" (0-15 psig)
Cell
(blueRed
Excitation 2
marked wire) Black
Ground
Green
3 High
Lower
White
3 Low
"White Δ" (0-15 psig)
Cell
(non-marked Red
Excitation 2
wire)
Black
Ground
122
Notes about Tensiometer hook-up to Pressure Transducers:
0.5 m depth Tensiometer--used "butt splices" (2-sided female-female crimps) to wire transducer
to tensiometer wires with 3M's Scotchkote
1.0 m depth Tensiometer--used "butt splices" and "nipple crimps" to wire transducer to
tensiometer wires with 3M's Scotchkote
1.5 m depth Tensiometer--used "nipple crimps" to wire transducer to tensiometer wires with
3M's Scotchkote
123
APPENDIX D. FIELD SITE SOIL PH PROFILES AND PARTICLE SIZE
DISTRIBUTION ANALYSES
Soil pH determined by placing pH probe in a soil: water (1:2) slurry made on site augering to depths
Particle size distribution analyses made at 6 different depths by hand augering to sample depths
and using undisturbed samples from soil cores recovered
Data Table from Figure 4.1.
Table of pHw (from soil:water slurry) for W2 Soils
Date
2/23/2009 2/24/2009 2/25/2009 2/26/2009
Sampled:
Depth (m)
S1
S2
S3
S4
0.0
*
6.33
6.25
6.05
0.5
4.98
5.33
5.37
6.15
1.0
6.31
5.53
6.11
5.54
1.5
5.07
4.77
5.56
5.10
2.0
5.16
*
5.34
5.15
* = no data
Table 4.1. Particle Size Distribution
Sample # Depth (m) Sand (%) Silt (%)
Clay (%)
A
0.5
30
16
54
B
1.0
43
24
33
C
1.5
45
19
36
D
1.6
36
9
55
E
1.8
31
11
58
F
2.0
33
18
49
Position soil samples excavated from in Table 4.1:
Sample A excavated between 0.5 m lysimeter & 0.5 m tensiometer
Sample B excavated between 1.0 m lysimeter & 1.0 m tensiometer
Sample C excavated between 1.5 m lysimeter & 1.5 m tensiometer
Sample D excavated next to field box
Sample E excavated next to field box
Sample F excavated next to field box
124
Soil pH
4.6
4.8
5
5.2
5.4
5.6
5.8
6
6.2
6.4
0.0
0.2
0.4
0.6
Depth (m)
0.8
1.0
1.2
Soil 1
1.4
Soil 2
1.6
Soil 3
Soil 4
1.8
2.0
Soil pH vs. Depth for Four W2 Field Site Soils
125
6.6
APPENDIX E. W2 MANAGEMENT: FERTILIZING, LIMING, SPRAYING, AND
PLANTING
USDA-ARS Watershed 2 Pasture Management
Fertilization (N, P as P2 O5 , K as K2 O), Liming, Pesticide/Herbicide Spraying, and Planting
DATE
TREATMENT
1992
Febuary, 1992
Fertilizer
April, 1992
Fertilizer
April, 1992
Spray
December, 1992
Fertilizer
1993
March, 1993
Fertilizer
July, 1993
Fertilizer
November, 1993
Fertilizer
1994
Febuary, 1994
Fertilizer
1995
March, 1995
Fertilizer
1996
January, 1996
Fertilizer
April, 1996
Fertilizer
November, 1996
Fertilizer
1997
3/17/1997
Fertilizer
5/16/1997
9/8/1997
November, 1997
1998
March, 1998
October, 1998
Fall 1998
Early 1999
2/1/1999
2/26/1999
4/1/1999
10/1/1999
Fall 1999
Spray
Lime
Plant
Fertilizer
Fertilizer
Fertilizer
Fertilizer
Spray
Fertilizer
Fertilizer
Plant
TYPE
No plant
10-10-10
34-0-0
2-4-D
18-0-27
No plant
18-0-27
34-0-0
14-7-14
No plant
17-0-17
No plant
10-10-10
No plant
10-10-10
15-0-15
10-10-10
No plant
10-10-10
FERTILIZER
QUANTITY
NITROGEN
QUANTITY
400 lbs/ac (448 kg/ha)
150 lbs/ac (168 kg/ha)
1 qt/ac (2.34 L/ha)
225 lbs/ac (252 kg/ha)
40 lbs N/ac (45 kg N/ha)
50 lbs N/ac (56 kg N/ha)
225 lbs/ac (252 kg/ha)
180 lbs/ac (202 kg/ha)
300 lbs/ac (336 kg/ha)
41 lbs N/ac (46 kg N/ha)
61 lbs N/ac (69 kg N/ha)
42 lbs N/ac (47 kg N/ha)
300 lbs/ac (336 kg/ha)
51 lbs N/ac (57 kg N/ha)
400 lbs/ac (448 kg/ha)
40 lbs N/ac (45 kg N/ha)
400 lbs/ac (448 kg/ha)
400 lbs/ac (448 kg/ha)
400 lbs/ac (448 kg/ha)
40 lbs N/ac (45 kg N/ha)
60 lbs N/ac (67 kg N/ha)
40 lbs N/ac (45 kg N/ha)
400 lbs/ac (448 kg/ha)
40 lbs N/ac (45 kg N/ha)
8 ac (3.24 ha) along
Wellbrook Rd.
41 lbs N/ac (46 kg N/ha)
Grazon
1qt/ac (2.34 L/ha)
Lime
1 ton/ac (2242 kg/ha)
Rye
2 bu/ac (174 L/ha)
No plant
10-10-10
400 lbs/ac (448 kg/ha) 40 lbs N/ac (45 kg N/ha)
17-17-17
300 lbs/ac (336 kg/ha) 51 lbs N/ac (57 kg N/ha)
10-10-10
400 lbs/ac (448 kg/ha) 40 lbs N/ac (45 kg N/ha)
Note: Fences removed and modern W2 created
34-0-0
200 lbs/ac (224 kg/ha) 68 lbs N/ac (76 kg N/ha)
Weedone
2 qt/ac (4.7 L/ha)
17-17-17
300 lbs/ac (336 kg/ha) 51 lbs N/ac (57 kg N/ha)
17-17-17
300 lbs/ac (336 kg/ha) 51 lbs N/ac (57 kg N/ha)
Rye
2 bu/ac (174 L/ha)
126
DATE
2/23/2000
8/22/200
10/1/2000
3/27/2000
11/1/2000
11/1/2000
9/26/2001
Fall 2001
2/14/2002
3/28/2002
9/24/2002
10/2/2002
Fall 2002
2/25/2003
3/28/2003
10/7/2003
11/1/2003
2004
4/1/2004
4/1/2004
11/1/2004
1/16/2005
3/5/2005
Fall 2005
2006
8/20/2006
9/11/2006
10/23/2006
2007
8/7/2007
2008
2008
2009
9/10/2009
TREATMENT
Fertilizer
Spray
Fertilizer
Spray
Plant
Plant
Fertilizer
Plant
Fertilizer
Spray
Fertilizer
Spray
Plant
Fertilizer
Spray
Fertilizer
Plant
Spray
Fertilizer
Lime
Fertilizer
Fertilizer
Plant
Fertilizer
Fertilizer
Lime
Fertilizer
Urea w/ Sulfur
+ nutrisphere
TYPE
17-17-17
Sevin
17-17-17
Grazon
Rye
Crimson C lover
17-17-17
Rye
17-17-17
Grazon
17-17-17
Sevin
Rye
17-17-17
Grazon
17-17-17
Rye
No plant
Grazon
Nitrogen
Lime
34-0-0
34-0-0
Rye
No plant
Urea w/ Sulfur (33-0-0)
15-0-15
Lime
No plant
Urea + Sulfur (33-0-0)
No plant
No fertilizer
No plant
33-0-0
127
FERTILIZER
QUANTITY
300 lbs/ac (336 kg/ha)
1.25 lbs/ac (1.40 kg/ha)
300 lbs/ac (336 kg/ha)
1.5qt/ac (3.5 L/ha)
2 bu/ac (174 L/ha)
15 lbs/ac (17 kg/ha)
300 lbs/ac (336 kg/ha)
2 bu/ac (174 L/ha)
300 lbs/ac (336 kg/ha)
1.5 qt/ac (3.5 L/ha)
300 lbs/ac (336 kg/ha)
1.25 lbs/ac (1.40 kg/ha)
2 bu/ac (174 L/ha)
300 lbs/ac (336 kg/ha)
1 qt/ac (2.34 L/ha)
300 lbs/ac (336 kg/ha)
2 bu/ac (174 L/ha)
NITROGEN
QUANTITY
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
51 lbs N/ac (57 kg N/ha)
1 qt/ac (2.34 L/ha)
80 lbs N/ac (90 kg N/ha) 80 lbs N/ac (90 kg N/ha)
1 ton/ac (2242 kg/ha)
200 lbs/ac (224 kg/ha) 68 lbs N/ac (76 kg N/ha)
300 lbs/ac (336 kg/ha) 102 lbs N/ac (114 kg N/ha)
2 bu/ac (174 L/ha)
200 lbs/ac (224 kg/ha)
200 lbs/ac (224 kg/ha)
1 ton/ac (2242 kg/ha)
66 lbs N/ac (74 kg N/ha)
30 lbs N/ac (34 kg N/ha)
250 lbs/ac (280 kg/ha)
83 lbs N/ac (92 kg N/ha)
200 lbs/ac (224 kg/ha)
66 lbs N/ac (74 kg N/ha)
APPENDIX F. GEOCHEMICAL RESULTS
Geochemical Results for Soil Water, Groundwater, & Precipitation
Soil Water collected from suction lysimeters installed at 4 depths in the vadose zone
L2B, L3A and L4A hand-made using porous ceramic cups (Soil Moisture Inc., Santa Barbara, USA)
& potable water-grade pvc body tubes with rubber stopper tops with glass & tygon tubing
NA = no analysis
Geochemical Analyses of Soil Water from Lysimeters
Sample
Total Dissolved
Depth
+
Organic C Fe 2+
Date
(m) & [NO3 -N] [NH4 ] Urea
N
Sampled Lysimeter (mg/L) (mg/L) (mg/L) (mg/L) (mg/L) (mg/L)
3/10/2009
0.35
35.24 0.048 0.000 33.420
39.2
NA
3/24/2009
(L1)
15.06 0.027
16.926
32.4 0.209
3/31/2009
7.83 0.025
8.754
43.5 0.372
4/15/2009
0.85 0.044
5.697
85.2 0.076
4/28/2009
1.15 0.236
2.643
50.8 0.143
5/12/2009
1.00 0.119
NA
67.44 0.848
6/9/2009
no volume
9/23/2009
no volume
10/16/2009
43.80 0.076
NA
6.674
50.28 0.025
NA
63.0 1.047
10/29/2009
11/13/2009
44.46 0.064
NA
45.072 0.005
11/19/2009
4.55 0.077
NA
45.756 0.391
5/12/2009
0.35
no volume
6/9/2009 (L1A) no volume
9/23/2009
23.84 0.044
24.585
9.490 0.953
10/16/2009
23.29 0.043
24.081
9.25 0.028
10/16/2009
dupl.
23.51 0.071
23.808
9.25 0.026
10/29/2009
16.15 0.053
NA
7.40
NA
11/13/2009
2.75
NA
NA
NA
11/19/2009
20.60 0.093
NA
16.500 0.115
128
Fe
3+
(Total Fe Fe 2+)
Sample
(mg/L) Number
NA
1
0.090
7
0.086
12
0.104
16
0.078
21
0.163
28
1.666
0.259
0.156
0.111
55
76
93
105
1.156
0.062
0.080
NA
NA
0.077
44
56
57
88
102
107
Geochemical Analyses of Soil Water from Lysimeters
Sample
Depth
+
Date
(m) & [NO3 -N] [NH4 ] Urea
Sampled Lysimeter (mg/L) (mg/L) (mg/L)
3/10/2009
0.5
6.31 0.025 0.000
3/24/2009
(L2)
2.50 0.026
3/31/2009
1.81 0.034
4/15/2009
0.93 0.029
4/28/2009
0.89 0.064
5/12/2009
1.21 0.060
6/9/2009
no volume
9/23/2009
no volume
10/16/2009
no volume
10/29/2009
no volume
11/13/2009
no volume
11/19/2009
30.63 0.099
5/12/2009
0.5
3.17 0.469
6/9/2009
(L2A) no volume
9/23/2009
no volume
10/16/2009
3.12 0.119
10/16/2009
dupl.
3.24 0.114
10/29/2009
2.19 0.904
11/13/2009
1.77 0.308
11/19/2009
1.40 0.145
9/23/2009 0.5 (L2B)
22.79 0.036
9/23/2009
dupl.
23.19 0.039
10/16/2009
60.69 0.009
10/16/2009
dupl.
64.80 0.029
10/29/2009
NA 0.000
10/29/2009
dupl.
NA 0.007
11/13/2009
NA 0.017
11/13/2009
dupl.
NA 0.016
11/19/2009
NA 0.032
129
Fe 3+
(Total Fe -
Total Dissolved
2+
Fe 2+)
Sample
N
Organic C Fe
(mg/L) (mg/L) (mg/L) (mg/L) Number
7.631
25.4
NA
NA
2
4.065
23.0 0.063
0.082
8
3.338
29.5 0.217
0.085
13
3.052
40.6 0.009
0.052
17
2.860
35.6 0.109
0.063
22
NA
32.52 0.038
0.087
29
NA
NA
16.665
8.034
0.011
0.566
0.058
0.293
106
32
3.843
3.939
NA
NA
NA
23.394
24.360
75.57
76.83
NA
NA
NA
NA
NA
NA
NA
2.85
2.840
3.222
5.960
5.625
4.55
5.15
2.70
2.50
3.160
2.941
3.682
0.274
0.016
0.000
0.000
0.137
0.636
0.002
0.000
0.050
0.051
0.043
0.000
0.000
0.081
0.489
0.072
0.074
0.092
0.090
1.537
0.062
0.071
0.093
0.079
0.067
0.087
0.125
0.119
60
61
79
94
109
45
46
58
59
77
78
95
96
108
Geochemical Analyses of Soil Water from Lysimeters
Sample
Depth
+
Date
(m) & [NO3 -N] [NH4 ] Urea
Sampled Lysimeter (mg/L) (mg/L) (mg/L)
3/10/2009
1.25
19.60 0.148 0.000
3/24/2009
(L3)
17.06 0.376
3/31/2009
17.91 0.245
4/15/2009
18.27 0.097
4/28/2009
15.30 0.149
5/12/2009
17.51 0.301
6/9/2009
18.09 0.253
9/23/2009
no volume
10/16/2009
21.39 0.016
10/16/2009
dupl.
21.71 0.042
10/29/2009
20.74 0.020
10/29/2009
dupl.
21.43 0.004
11/13/2009
20.93 0.033
11/13/2009
dupl.
21.24 0.031
11/19/2009
20.44 0.067
5/12/2009
1.25
3.74 0.235
6/9/2009 (L3A) no volume
9/23/2009
14.18 0.086
10/16/2009
14.06 0.015
10/16/2009
dupl.
13.93 0.018
10/29/2009
14.48 0.000
10/29/2009
dupl.
14.18 0.000
11/13/2009
14.10 0.019
11/13/2009
dupl.
10.55 0.020
11/19/2009
14.96 0.042
130
Fe 3+
(Total Fe -
Total Dissolved
2+
Fe2+)
Sample
N
Organic C Fe
(mg/L) (mg/L) (mg/L) (mg/L) Number
19.084
3.12
NA
NA
3
19.792
1.33 0.019
0.117
9
19.972
1.03 0.585
0.246
14
19.840
0.75 0.011
0.070
18
18.838
0.68 0.057
0.074
23
19.190
1.080 1.112
0.365
30
NA
2.23 0.000
0.084
34
21.732
22.029
NA
NA
NA
NA
NA
4.606
0.56
0.62
0.41
0.44
0.782
0.560
0.588
94.63
0.023
0.116
0.015
NA
0.000
0.000
0.262
0.019
0.087
0.332
0.057
NA
0.127
0.157
0.116
0.726
62
66
80
87
97
98
110
33
17.090
15.025
14.995
NA
NA
NA
NA
NA
9.11
6.05
6.00
2.80
3.00
3.304
6.748
3.542
0.000
0.048
0.620
0.072
0.005
0.000
NA
0.000
0.055
0.167
0.164
0.082
0.061
0.095
NA
0.076
47
63
67
81
82
99
103
111
Geochemical Analyses of Soil Water from Lysimeters
Sample
Depth
[NO3 --N] [NH4 +] Urea
Date
(m) &
Sampled Lysimeter (mg/L) (mg/L) (mg/L)
3/10/2009 1.75 (L4)
4.13 0.057 0.018
3/10/2009
dupl.
4.11 0.058 0.000
3/24/2009
4.99 0.046
3/24/2009
dupl.
5.02 0.045
3/31/2009
5.13 0.044
4/15/2009
5.10 0.033
4/15/2009
dupl.
5.05 0.037
4/28/2009
4.75 0.068
5/12/2009
4.48 0.031
6/9/2009
3.81 0.021
6/9/2009
dupl.
3.84 0.030
9/23/2009
2.84 0.020
9/23/2009
dupl.
2.77 0.022
10/16/2009
2.35 0.051
10/16/2009
dupl.
2.27 0.051
10/29/2009
2.08 0.041
10/29/2009
dupl.
2.18 0.043
11/13/2009
2.63 0.070
11/19/2009
3.27 0.084
6/9/2009
1.75
no volume
9/23/2009
(L4A)
2.84 0.020
9/23/2009
dupl.
3.11 0.017
9/23/2009
dupl.
3.00 0.017
10/16/2009
0.28 0.018
10/16/2009
dupl.
0.24 0.019
10/29/2009
0.38 0.000
10/29/2009
dupl.
0.37 0.000
11/13/2009
0.84 0.017
11/13/2009
dupl.
0.94 0.016
11/19/2009
0.79 0.040
Trip
Blanks
(DI Water)
0.03 0.004 0.000
NA = no analysis
131
Fe 3+
(Total Fe -
Total Dissolved
2+
Fe2+)
Sample
N
Organic C Fe
(mg/L) (mg/L) (mg/L) (mg/L) Number
4.731
0.61
NA
NA
4
4.758
0.00
NA
NA
5
5.058
0.54
0
0.076
10
5.167
0.66
NA
NA
11
5.549
0.48 0.207
0.133
15
5.717
0.26
0
0.052
19
5.770
0.35
NA
NA
20
5.388
0.35 0.042
0.087
24
5.516
0.856 0.325
0.268
31
3.911
0.835 1.038
0.260
35
3.918
0.567 0.581
0.301
36
3.319
0.789 0.008
0.069
42
2.677
0.601 0.217
0.515
43
2.545
0.59 2.416
0.647
64
2.561
0.86 2.091
0.197
68
NA
0.57 0.917
0.107
83
NA
0.55 1.002
0.152
84
NA
5.892 0.001
0.342
100
NA
0.983 0.128
0.074
112
3.227
3.385
3.295
0.3328
0.3203
NA
NA
NA
NA
NA
1.971
1.792
1.794
4.02
3.71
2.93
2.96
8.193
6.323
2.414
0.136
0.008
0.000
0.008
0.014
0.021
0.040
0.000
NA
0.022
0.159
0.055
0.065
0.068
0.063
0.099
0.131
0.136
NA
0.062
48
49
50
65
69
85
86
101
104
113
0.056
0.000
NA
NA
6
Soil Water Summary Statistics with Depth
Chemical
Variable
Number
Minimum Maximum Mean
of
-1
-1
-1
Depth (m) Samples (mg L ) (mg L ) (mg L )
Sampling Standard
Error +/- Deviation
-1
(mg L )
-1
(mg L )
0.35
16
0.85
50.28
19.6
0.11
16.6
(NO3 -N)
0.5
1.25
1.75
17
23
29
0.89
3.74
0.24
64.80
21.71
5.13
7.01*
17.4*
2.88
2.06
1.78
0.14
20.7
4.28
1.64
Ammonium
0.35
15
0.025
0.236
0.058*
0.014
0.053
0.5
1.25
22
23
0.000
0.000
0.904
0.376
0.047*
0.084*
0.010
0.013
0.207
0.111
1.75
0.35
0.5
1.25
1.75
0.35
0.5
1.25
1.75
0.35
29
8
11
12
20
14
20
23
29
13
0.000
2.643
2.86
4.606
0.3203
7.40
2.5
0.41
0.00
0.005
0.084
33.42
76.83
22.029
5.770
85.2
40.6
94.63
8.193
6.674
0.035
17.49
8.498*
18.87*
3.86
37.5
12.8
2.17*
1.23*
0.351*
0.005
0.137
0.63
0.149
0.321
0.00
0.30
1.72
0.94
0.001
0.021
10.8
28.5
4.68
1.63
24.68
13.09
19.36
2.048
1.792
0.5
1.25
1.75
0.35
21
20
24
13
0.000
0.000
0.000
0.062
0.636
1.112
2.416
1.666
0.046*
0.024*
0.214*
0.115*
0.317
0.286
0.229
0.009
0.180
0.292
0.665
0.500
0.5
1.25
1.75
* = value excludes outliers
21
20
24
0.052
0.055
0.052
1.537
0.726
0.647
0.081*
0.108*
0.134*
0.738
0.138
0.223
0.327
0.159
0.152
Nitrate-N
-
+
(NH4 )
Total N
(TN)
Dissolved
Organic
Carbon
(DOC)
Ferrous
2+
Iron (Fe )
Ferric
3+
Iron (Fe )
132
Soil Water Summary Statistics for Each Chemical Variable
Sampling
Number
Minimum Maximum Mean
Error +/Chemical
of
-1
Variable Samples (mg L )
-
-1
(mg L )
-1
(mg L )
-1
(mg L )
Standard
Deviation
-1
(mg L )
F
Value*
2
R *
NO3 -N
85
0.242
64.80
11.93
2.06
13.46
0.254
9.20
NH4 +
Total N
DOC
89
51
86
0.000
0.320
0.000
0.904
76.83
94.63
0.077
12.90
11.43
0.014
0.63
1.72
0.122
15.34
19.38
0.096
0.230
0.395
2.23
4.69
17.9
2+
78
0.000
6.674
0.325
0.317
0.848
0.091
2.47
Fe
3+
78
0.052
1.666
0.195
0.738
0.286
0.035
0.90
*Statistical analysis (ANOVA) was run in SAS using a model comparing each chemical variable to depth
2
in order to see if depth influenced the chemical variable (where R and F come from)
Fe
Soil Water Average Concentration Values with Depth under Tree Canopy
Fe3+
Sample
Depth (m) Lysimeter
0.35
L1
0.50
L2
1.25
L3A
1.75
L4A
-
+
[NO3 -N] [NH4 ] Urea
(mg/L) (mg/L) (mg/L)
20.42 0.074 0.000
6.33 0.048 0.000
12.69 0.048
1.28 0.015
Total Dissolved
N
Organic Fe2+
(mg/L) C (mg/L) (mg/L)
13.488
52.485
1.085
4.189
29.041
0.074
12.929
15.020
0.094
2.112
3.611
0.026
(Totl. Fe Fe 2+)
(mg/L)
0.301
0.071
0.178
0.093
Soil Water Average Concentration Values with Depth under Open Sky
Fe3+
Sample
Depth (m) Lysimeter
0.35
L1A
0.50
L2A
0.50
L2B
1.25
L3
1.75
L4
-
+
[NO3 -N] [NH4 ] Urea
(mg/L) (mg/L) (mg/L)
18.36 0.060
2.48 0.343
42.87 0.019
19.40 0.127 0.000
3.73 0.045 0.000
133
Total Dissolved
N
Organic Fe2+
(mg/L) C (mg/L) (mg/L)
24.158
11.123
0.281
3.891
4.237
0.164
50.039
4.030
0.094
20.060
1.013
0.182
4.439
0.855
0.597
(Totl. Fe Fe 2+)
(mg/L)
0.344
0.185
0.249
0.153
0.219
Soil Water Average Concentration Values with Depth in High Nitrogen Area
Fe3+
(Totl. Fe -
+
[NO3 -N] [NH4 ] Urea
Sample
Depth (m) Lysimeter (mg/L) (mg/L) (mg/L)
0.35
L1A
18.36 0.060
0.50
L2B
42.87 0.019
1.25
L3
19.40 0.127 0.000
Total N
(mg/L)
24.158
50.039
20.060
Dissolved
Fe 2+)
Organic Fe2+
C (mg/L) (mg/L) (mg/L)
11.123 0.281
0.344
4.030 0.094
0.249
1.013 0.182
0.153
Piezometer Groundwater Samples Geochemical Results
Fe 3+
Depth to
(Totl. Fe GroundTotal Dissolved
+
2+
Fe 2+)
water Sample
N
Piezometer [NO3 -N] [NH4 ]
Organic C Fe
(m)
Number
(mg/L) (mg/L) (mg/L) (mg/L) (mg/L) (mg/L)
Location
Date
Up-Slope of
4/28/2009 Site Piez. 7
3.45 0.055 3.561
0.44 0.031
0.068
1.99
27
6/10/2009
3.81 0.013 3.618
0.764 0.334
1.031
2.13
37
11/19/2009
4.51 0.149
NA
1.572
NA
NA no data
116
On-Site
Piez. 2
4.60 0.043 5.322
0.14 0.093
0.128
1.83
26
4/28/2009
6/10/2009
4.69 0.026 5.103
0.631 0.013
0.088
1.85
39
11/19/2009
4.41 0.031
NA
0.678
NA
NA no data
118
Down-Slope
4.95 0.111 5.665
0.75 0.171
0.287
1.42
25
Piez. 3
4/28/2009
6/10/2009
4.14 0.024 4.287
0.753 0.300
1.281
1.52
41
11/19/2009
4.51 0.026
NA
0.416
NA
NA no data
120
Side of Site
2.97 0.059 3.004
1.177 0.305
1.088
2.48
38
Piez. 1
6/10/2009
11/19/2009
6.87 0.078
8.129
NA
NA no data
117
Side of Site
4.24 0.019 4.483
0.604 0.615
0.908
1.70
40
Piez. 6
6/10/2009
11/19/2009
3.31 0.025
NA
0.369
NA
NA no data
119
NA = no analysis
134
Geochemical Analyses of Precipitation
Dissolved
Total Organic
+
Precipitation [NO3 -N] [NH4 ] N
Sample
C
(mg/L) (mg/L) (mg/L) (mg/L) Number
Location
Date
ARS-Main
10/15/2009*
lot
0.19 0.008 0.212
3.64
51
10/15/2009*
0.19 0.008 0.211
2.58
52
10/15/2009*
0.19 0.005 0.212
2.67
53
10/15/2009*
0.19 0.010 0.202
3.02
54
10/26/2009 W2 Field Site
0.45 1.303
11.7
70
10/26/2009
0.44 1.367
12.8
71
10/27/2009
0.18 0.034
0.95
72
10/27/2009
0.18 0.035
0.60
73
10/27/2009
0.18 0.037
0.68
74
10/27/2009
0.18 0.030
0.56
75
11/12/2009
0.22 0.126
1.715
89
11/12/2009
0.23 0.124
1.120
90
11/12/2009
0.23 0.104
1.144
91
11/12/2009
0.22 0.127
1.074
92
11/19/2009
0.37 0.106
1.606
114
11/19/2009
0.37 0.180
4.752
115
* Multiple Precipitation Events C ollected in O ne Sample Bottle at This Site
135