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My Other Home is a Mesocosm:
A Water Quality Analysis of Three Different Rearing Treatments for Rana pretiosa
Brianna Lorraine Morningred
Submitted in partial fulfillment of the requirements for the degree of
Master of Environmental Studies
June 2015
ii
©2015 by Brianna Lorraine Morningred. All Rights Reserved
iii
This Thesis for the Master of Environmental Studies Degree
by
Brianna Lorraine Morningred
has been approved for
The Evergreen State College
by
________________________
Dr. Dina Roberts
Member of the Faculty
iv
ABSTRACT
My Other Home is a Mesocosm: A Water Quality Analysis of Three Different
Rearing Treatments for Rana pretiosa
Brianna Lorraine Morningred
Rana pretiosa populations have been in decline for many years. As a result, there are
many captive rearing programs that are working to boost the local populations by
rearing these frogs in captivity and releasing them back into the wild. Despite the
hundreds of healthy young frogs that have been released into the wild, there has not
been a corresponding increase in egg mass numbers during the survey season. It is
theorized that this may be due to over-stimulation in captivity, and as a result the
scientists working on the R. pretiosa recovery plan wanted to explore the possibility
of rearing the young frogs in a Mesocosm environment. Mesocosms are meant to be
a self-sustaining ecosystem unit, with balanced chemical and nutrient cycles that
mimic the natural environment as closely as possible. It is hoped that this type of
environment will reduce the human contact with the frogs during captivity,
reducing their stimulation and keeping their predator-evasion response times fast.
However, there are still many factors of the mesocosm environment that we must
learn before rearing Rana pretiosa in them. This research project looked into
gathering baseline data for water quality parameters, comparing the three potential
rearing habitats for the Oregon Spotted Frog: Traditional, Mesocosm and Prairie
Wetland. The water quality measurements taken and compared were pH, dissolved
oxygen, temperature; and nutrient concentrations of chloride, sulfate, phosphorous
and nitrate. In each of these parameters both captive rearing treatments
(Traditional and Mesocosm) were significantly different than the Prairie wetland
environment. These results indicate not only that the Prairie wetland environment is
much more variable than either captive rearing environment, but this may also
mean that more variability can be applied to the captive rearing environments to
better prepare the juvenile frogs for their release into the wild. More extensive
research is needed to explore the water quality—especially measurements of
chemistry—fluctuations of a Mesocosm environment, however providing a more
variable rearing environment for Rana pretiosa may prove beneficial..
v
Table of Contents
Chapter 1: Literature Review………...………………………………..p.1
Chapter 2: Introduction………………………………………………..p.10
Methods……………...…………………………………….p.14
Results……………………………………………………...p.20
pH, Dissolved Oxygen, Temperature …………………….p.20
Nutrient Analysis ……………………………...………p.22
Discussion………...………………………………………..p.27
Captive Rearing Management Recommendations…...…p.34
Sources……………………………………………………..p.36
Appendix A: Materials List…………………………………………….p.44
Appendix B: Full Run List for Ion Chromatograph…………………p.45
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List of Figures and Tables
Figures
Figure 1: Woodland Park Zoo Captive Rearing Tanks (image)……………………….p.15
Figure 2: pH data (graph)………………………………………………………………p.22
Figure 3: Dissolved oxygen data (graph)………………………………………………p.22
Figure 4: Temperature data (graph)……………………………………………………p.23
Figure 5: Phosphorous concentrations (graph)………………………………………...p.24
Figure 6: Nitrate concentrations (graph)……………………………………………….p.25
Figure 7: Chloride concentrations (graph)……………………………………………..p.26
Figure 8: Sulfate concentrations (graph)……………………………………………….p.27
Tables
Table 1: Sampling Plan for Nutrients………………………………………………….p.16
Table 2: Stock Standard Solution Recipes…………………………………………….p.18
Table 3: Average Phosphorous Concentrations……………………………………….p.24
Table 4: Average Nitrate Concentrations……………………………………………...p.25
Table 5: Average Chloride Concentrations……………………………………………p.27
Table 6: Average Sulfate Concentrations……………………………………………...p.28
Acknowledgements
This thesis research project was made possible by several partners. I would like to
graciously acknowledge Dr. Dina Roberts, Dr. Marc Hayes, Julie Tyson, Dr. Jennifer
Pramuk, Alyssa Borek, and the Sustainability in Prisons Project. I was a hopeful nerd
when this began, and now I feel like a real scientist: I could not have done this without
all of you and I am so grateful. May all of you continue to inspire many more hopeful
nerd for years to come. I would also like to acknowledge my incredible family and
friends: particularly my nerdtastic parents, Wendy and Daniel Morningred, my
wonderful brother and sister-in-law Aaron and Heidi, my Nerd—the love of my life, and
my dear friends who are too numerous to name here. I am proud to have accomplished
this great feat, but I could not have dreamed about doing so without you by my side, at
my back, and sometimes pushing me forward. Nothing I am, or ever will be, would be
possible without you.
vii
CHAPTER 1. LITERATURE REVIEW
Introduction
Oregon Spotted frog Ecology
Rana pretiosa, the Oregon Spotted frog, is one of 46 amphibian species native to
Washington State (Washington State DNR, 2009). R. pretiosa was once vastly distributed
from northern California to southern British Columbia. However, it has since disappeared
from 70-90% of its historic geographical range (Pearl & Hayes, 2005). Currently its
range is restricted to isolated sites in western and south-central Washington and the east
Cascades region of central and south-central Oregon. R. pretiosa is listed as an
endangered Species both in Washington State and as threatened federally under the
Endangered Species Act of 1973 (WDFW 2013).
R. pretiosa is medium-sized (roughly 4.4-10.5cm length vent to snout) and is
classified as a highly aquatic frog that is closely associated with permanent water
(Conlon et al., 2011). Females are typically larger than males and can reach up to 100
millimeters (4 inches) (Leonard et al. 1993).While there is variation due to age, this
species is distinguished from other PNW Ranids by the following characteristics:
“The dark spots have ragged edges and light centers, which are usually
associated with tubercles or raised areas of skin; these spots become larger
and darker and the edges become more ragged with age. Body color also
varies with age. Juveniles are usually brown or, occasionally, olive green
1
on the back and white or cream with reddish pigments on the underlegs
and abdomen. Adults range from brown to reddish brown, but tend to
become redder with age; large, presumably older individuals may be brick
red over most of the back. Red increases on the abdomen with age, and the
underlegs become a vivid orange-red. This red coloration can be used to
distinguish the spotted frogs from other native frogs,” (USFWS, 2014)
R. pretiosa frequents heavily vegetated wetlands, though the mechanisms behind
this habitat preference are not well understood (Watson, McAllister, & Pierce, 2003).
Pearl, 2005), Pearl and authors (2005) observed Oregon Spotted frogs in typical basking
positions, eyes above the water with body partially submerged, on or among floating
vegetation mats consisting mainly of algae and bladderwort (Utricularia sp.). Studies
conducted on the predator evasion techniques of Anuran species (including R. pretiosa)
observed frogs in a “frozen” or motionless posture in the water column (Rand, 1952;
Heatwole, 1961; Gans and Rosenberg, 1966; Hedeen 1972).
Current Population Decline Theories
Amphibians are now in greater peril than at any time in recent geologic history
(Lannoo, 2005). These are perilous times for amphibians, as evidenced by fully one-third
of all amphibians worldwide are now considered threatened (Stuart et al., 2004). Native
herpetological diversity in northwestern North America is in part a result of the complex
geological processes that formed the massive mountain ranges and large plains of the
region and subsequently split historical species ranges, fragmented habitats, and altered
climates (Nussbaum, Brodie, & Storm, 1983). It is thus speculated that northwestern
2
amphibian species are reflective of current landscape diversity (Olson, 2009). Due to
species habitat preferences outlining distributions, a species may not occupy all suitable
habitats within its range due to many factors including, but not limited to, stochastic
events affecting current population dynamics and lingering after-effects of historical
disturbance events (Olson, 2009). Habitat loss and extirpation from historic ranges
necessitate species-specific conservation plans for at- risk species.
The scientific community, in an effort to understand why amphibian species—
even those on protected lands—were disappearing, hypothesized a list of the six most
influential causes of amphibian decline. In no particular order: habitat
destruction/modification, commercial over-exploitation, non-native species introduced to
native habitat, environmental contaminants, global climate change and emerging
infectious diseases (the most concerning being chytrid fungus Bactrachochytrium
dendrobatidis (Collins & Storfer, 2003).
Habitat loss and alteration/degradation are considered among the most likely
causes of the decline of the R. pretiosa (McAllister & Leonard, 1997). The loss and
degradation of shallow breeding wetlands are particularly concerning as R. pretiosa is
reliant on habitat that stays inundated year-round. Watson et al. (2003) observed the
Oregon Spotted frog during an animal behavior study using a variety of different aquatic
habitats depending on the time of year. For example, they used shallow pools with stable
water levels for egg deposition and tadpole development during mating season in the
spring, deep pools for juveniles and adults in the dry seasons (suitable for temperature
regulation during the hotter months and for predator evasion), and finally shallow water
overlaying emergent vegetation during the winter rainy/icy season (Watson et al., 2003).
3
Due to their specific habitat uses and needs, they are very susceptible to habitat changes
and as a result population declines
Pollution of groundwater by agrochemicals (chemical runoff from agricultural
practices (Hayes & Jennings, 1986) is a prevalent problem in wetland habitats. Even now
many people see wetlands only as wastelands and have to place their current needs over
the preservation of important habitats for the future (Aber, Pavri, & Aber, 2012). As
such, pollution that reaches the wetlands originates from many different sources that
aren’t controlled (i.e. agricultural run-off, storm water run-off, etc). R. pretiosa spends its
whole life in the aquatic environment, and thus is vulnerable to direct exposure of
chemicals that are in the water. The egg-stage is especially susceptible to siltation and
water pollution (Bugg & Trenham, 2003). Due to their highly permeable skin, the
transdermal movement of toxins—absorption of toxins through the skin—can happen
easily. Deterioration in water quality can therefore have potentially lethal or sub-lethal
effects on amphibians (Boyer & Grue, 1995).
Mesocosms
Ecology is studied across varying geographic scales: from whole system scales to
mesocosms, to microcosms. Mesocosms are moderately sized (i.e., smaller than whole
ecosystem studies, bigger than microscopic ecosystem studies) man-made ecosystems
that are used as tools in ecological research—allowing a certain amount of control over
natural complexity through smaller scale, simplified studies. They are also used in
applied research and educational development (Kangas & Adey, 1996). They combine a
technological component related to the form of container structures with environmental
management and control of boundary exchanges with living populations (Kangas &
4
Adey, 1996). The mesocosm is an extension of the microcosm method, first developed by
EP Odum in 1960 (Beyers & Odum, 1993). It is a wonderful research tool as it allows the
researcher to replicate natural conditions as close as possible, while still exhibiting the
controls of a laboratory to aid replication and statistical validity. Kangas and Adey (1996)
put it best by stating that “mesocosms are special ‘windows’ along the spatial scale of
ecosystems for examining ecological questions.” The mesocosm was used initially in the
1970s as a basic terrarium in school classrooms, and now it is now often used when
conducting semi-experimental studies in aquatic ecology (Odum et al., 1993). It is
important to stress, however, the importance of replicability when using mesocosms. This
factor will prove crucial for any evaluation of ecosystem dynamics, including those
related to the factors I researched for my thesis. A suitable balance between mesocosm
replicability and ecological realism of the mesocosm must be found when using
mesocosms in research, preferably at reasonable costs (Kraufvelin, 1998).
Amphibian Rearing Practices
Current Water Quality Standards
The water quality parameters that can have potentially negative effects on
amphibian species are dissolved oxygen, temperature, pH, salinity/water conductivity,
organic carbons and pollution. All of these factors and the way they interact with each
other can affect survival, growth, maturation and physical development of amphibian
species (Dodd, 2010).
5
Members of the family Ranidae (i.e. Rana pretiosa) assimilate oxygen through
dermal uptake, gills, and lungs (Dodd, 2010). In hypoxic conditions, larvae with lungs
swim to the surface and gulp air into the lungs. Though this risks increased predator
exposure, the alternative is hypoxia which can induce similar physiological responses in
amphibians as in other vertebrates including changes in blood pH, build-up of lactate in
muscles, lethargy and death (Dodd, 2010). There are natural diurnal cycles of dissolved
oxygen in wetland environments, however. These cycles correspond to the rates of
photosynthesis versus respiration and varies seasonally as well as regionally (Dong, Zhu,
Zhao, & Gao, 2011).
Amphibians are ectotherms meaning they rely on elements in their environment to
regulate their temperature (i.e. shade when they’re hot, sunlight when they’re cold, etc).
Therefore water temperature is extremely important in metabolic function, physiological
processes and behavior. For most amphibian species, including R. pretiosa, between 10°
and 40°C each 10° increase in ambient temperature increases metabolism by 1.4-2.4
times (Rome, Stevens, & John-Alder, 1992). Licht (Licht, 1971) tested the range of
tolerance for R. pretiosa embryos in extreme temperatures. He reported a lethal
minimum, the temperature at which the eggs’ survival is less than 50% is approximately
6°C. There is also a general relationship between temperature and dissolved oxygen.
Generally speaking as temperature increases dissolved oxygen decreases (Dodd, 2010).
The final water quality parameter that I would like to address with my research is
pH. pH is the negative log of Hydrogen ion concentrations in water , scaling from 0-14,
with 6.0-7.5 considered “safe” or “neutral” conditions for most freshwater aquatic species
(Dodd, 2010). Natural factors affecting pH are the bedrock found in the area and the
6
concentration of organic acids (Dodd, 2010). However, anthropogenic sources of acidity
are acid deposition from the production of nitrous oxides and sulfates during the
processing of fossil fuels and acid mine drainage (Dodd, 2010). pH can affect successful
amphibian development at all life stages. For example, at pH values lower than 4.5,
embryonic development may cease entirely, whereas at high pH values development
continues but hatching is disrupted (Dunson & Connell, 1982). The critical pH or that
which can cause significant increases in mortality for amphibian embryos ranges from
5.0-3.5 (Freda & Dunson, 1985). The primary effects of being exposed to low pH waters
are interference with ion transport, compromised immune systems, inability for embryos
to hatch, reduced growth and delayed metamorphosis (Brodkin et al., 2003).
Issues with Captive Rearing
Captive rearing programs have been successfully raising R. pretiosa for a number
of years and releasing them back into their native habitat in hopes of boosting local
populations. Currently there are partnerships with government agencies (such as
Washington Department for Fish and Wildlife) and private organizations such as the
Oregon, Pt. Defiance and Woodland Park Zoos that foster captive rearing projects for the
rearing and reintroduction of the now endangered Rana pretiosa. However, despite years
of successful rearing and release, there has not been a physical confirmation of increased
populations in the form of egg masses which leads to one question—what is happening to
the frogs once they’re released?
Kyle Tidwell, currently a doctoral student at Portland State University wondered
the same question and began exploring the possibility of lowered predator responses
being responsible for the Oregon Spotted frog not bouncing back after successful captive
7
rearing programs. Kyle Tidwell, as well as his companion researchers, theorized that
perhaps the exposure to frequent contact during captive rearing is causing the frogs to be
“over-stimulated” and thus less responsive to predator attacks resulting in decreased
survivorship post-release. His study tested two separate populations of captively-reared
Oregon Spotted frogs with a ball drop test to simulate a predator dropping on them and
gauged their response. Though the rearing conditions for both populations were the same,
Tidwell did find a significant difference in response times between them, which suggests
that there are factors affecting the Oregon Spotted frogs ability to respond to a predator
threat (Tidwell, Shepherdson, & Hayes, 2013). He suggests that “it is possible that
husbandry activities such as cleaning and feeding made the frogs progressively warier,”
(Tidwell et al., 2013). Tidwell’s results led scientists and husbandry personnel involved
in the captive rearing program to wonder if a mesocosm environment might not be better
suited than captivity for raising the Oregon Spotted frogs and hopefully keeping them
properly “stimulated’ and aware of predators.
Mesocosm History in Captive Rearing
The use of aquatic mesocosms to study amphibian ecology was explored
beginning with the need to move away from purely descriptive field studies and
correlative analyses and move towards more manipulative studies that would allow for
hypothesis testing (Dodd, 2010). “Studies using aquatic mesocosms are a compromise
between the variability of natural wetlands, confounding factors and the lack of control
that is common in field experiments,” (Dodd, 2010). Mesocosms rather than microcosms
are used in amphibian rearing as they are self-sustaining once they are established, which
allows all organisms kept in the mesocosm to complete the critical phases of their life
8
cycle (Dodd, 2010). The two most common forms of mesocosms for amphibian rearing
are (1) cages or floating mesh tanks that are placed directly in the stream/lake/water body
the eggs are taken from and (2) large containers (i.e. plastic cattle watering tanks)
mimicking wetland systems. Either of these options give the researcher the ability to
capture a pocket of the natural environment, while still being able to manipulate much of
the environmental processes within the mesocosm either for study or for the support of
endangered amphibian species. However, it is crucial that we understand the nutrient and
chemical cycles that should occur within the mesocosms as a part of the natural processes
as well as what cycles need to be avoided to protect the health of the animals. My
research was designed to test for the differences between water chemistry and quality of
traditional, mesocosm and natural wetland ecosystems used by Oregon Spotted Frogs.
Wetland Biochemistry in Pacific Northwest Wetlands
Wetland ecosystems as well as mesocosms mimicking wetlands have diurnal
cycles of oxygen. These cycles revolve around the ratio of photosynthesis to respiration
which in turn are affected by temperature and pH. Water solutions typically contain
dozens of dissolved solids, such that overall charges balance for electrical neutrality
(Aber et al., 2012). The degree to which the wetland biochemical cycles fluctuate varies
by wetland and in some cases can have a very wide fluctuation either daily or seasonally.
The chemistry of wetlands depends on many factors including the influence of
bedrock and soil, inflow and outflow of surface and groundwater, climate and vegetation,
characteristics of surrounding terrain, and human impacts (Aber et al., 2012). “Key
wetland elements include nitrogen, potassium, iron and manganese, sulfur, phosphorous
and carbon,” (Aber et al., 2012). The specific chemical status of these elements within the
9
wetland environment depends primarily on the presence of oxygen within the wetland
waters (Aber et al., 2012).
“Nitrogen is a major nutrient and is often a limiting factor in flooded wetlands or
peat lands. Once ammonium is formed, it may be used directly by plants or anaerobic
microbes, which convert it again into organic matter. In marshes with algal blooms, pH
may exceed 8, in which case ammonium is converted into ammonia (NH3) and released
into the atmosphere (Mitsch & Gosselink, 2007). Under aerobic conditions, ammonium
may be converted into nitrite (NO2-) and then nitrate (NO3-),” (Aber et al., 2012)
Phosphorous, like nitrogen, is a limiting factor in wetlands. It’s most bioavailable
under neutral to slightly acidic conditions (Aber et al., 2012). Phosphorous is considered
to be an essential element for life in the DNA molecule and adenosine triphosphate,
which stores chemical energy. It is also an essential nutrient for growth and development
of algae and other plants (Stewart & Howell, 2003).
Sulfur is generally abundant in the wetland environments and thus is not likely to
be considered a limiting factor for plant growth. In the anaerobic zone, sulfur and sulfate
are reduced into hydrogen sulfide. This can alter the pH and thus the acidity of the
wetland water chemistry.
My research will begin to address the gaps in our knowledge about water
chemistry both in wetlands and captive rearing environments.
10
CHAPTER 2
Introduction
The Oregon Spotted frog, Rana pretiosa is a medium-sized, aquatic amphibian
native to Canada, Washington, Oregon and California (Watson, McAllister, & Pierce,
2003). R. pretiosa was once distributed from northern California to southern British
Columbia. However, it has since been extirpated from 70-90% of its historic geographical
range (Pearl & Hayes, 2005). Currently its range is restricted to isolated sites in western
and south-central Washington and the east Cascades region of central and south-central
Oregon. R. pretiosa is listed as an Endangered Species both in Washington State and
federally under the Endangered Species Act of 1973 (WDFW 2013, Federal Registry
2014). R. pretiosa is distinguished from other Pacific Northwest ranids by its unique
spotting pattern, higher eye placement and increased webbing on the feet (USFWS 2014).
These characteristics are related to R. pretiosa’s life history and highly aquatic habits.
Due to its completely aquatic lifestyle, R. pretiosa not only requires specific
habitat characteristics, but is also more vulnerable to various threats—both biological and
chemical—throughout its lifetime. There are many theories suggested as reasons for R.
pretiosa population declines. Top theories currently include habitat alteration, which can
range from small hydrological changes to transforming wetlands, predation by invasive
fish and amphibians, and possible physiological impairments caused by exposure to
toxins in the water (Pearl & Hayes, 2005).
Habitat loss and alteration/degradation are considered one of the most likely
causes of the decline of R. pretiosa (McAllister & Leonard, 1997). The loss and
11
degradation of shallow breeding wetlands are particularly concerning as R. pretiosa is
reliant on inundated wetland habitat year-round. Watson et al. (2003) observed the
Oregon Spotted frog making use of a variety of different aquatic habitats—which varied
with the time of year—during an animal behavior study. For example, the frogs used
shallow pools with stable water levels for egg deposition and tadpole development during
mating season in the spring, deep pools for juveniles and adults in the dry seasons
(suitable for temperature regulation during the hotter months and for predator evasion),
and finally shallow water overlaying emergent vegetation during the winter rainy/icy
season (Watson et al., 2003). Due to their specific habitat requirements, spotted frogs are
susceptible to habitat changes and as a result population declines.
Pollution of groundwater by agrochemicals (chemical runoff from agricultural
practices (Hayes & Jennings, 1986) is a prevalent problem in wetland habitats. Even now
many people see wetlands only as “wastelands” and place development and conversion
priorities over the preservation of these important wetland habitats for the future (Aber,
Pavri, & Aber, 2012). As such, pollution that reaches the wetlands originates from many
different sources that aren’t controlled (i.e. agricultural run-off, storm water run-off, etc.).
R. pretiosa spends its whole life in the aquatic environment, and thus is vulnerable to
direct exposure of chemicals that are in the water. The eggs especially are extremely
susceptible to siltation and water pollution (Bugg & Trenham, 2003). Due to their highly
permeable skin, the transdermal movement of toxins—absorption of toxins through the
skin—can happen easily. Deterioration in water quality can therefore have potentially
lethal or sub-lethal effects on amphibians (Boyer & Grue, 1995).
12
Captive rearing programs have been successfully raising R. pretiosa for a number
of years with the goal of releasing them back into their native habitat to boost local
populations or establish new populations. As guided by recovery goals for this species,
collaborative efforts between government agencies (such as Washington Department for
Fish and Wildlife) and private organizations such as the Oregon, Pt. Defiance and
Woodland Park Zoos (found in Portland, Tacoma and Seattle, respectively) that foster
captive rearing projects for the rearing and reintroduction of the now endangered Rana
pretiosa. However, despite years of successful rearing and release, there has not been a
physical confirmation of increased populations in the form of egg masses which leads to
one question—what is happening to the frogs once they’re released (K. Tidwell, personal
communication)? Kyle Tidwell, a doctoral student at Portland State University, as well as
his companion researchers, theorized that perhaps the exposure to frequent contact during
captive rearing is causing the frogs to be “over-stimulated” and thus less responsive to
predator attacks resulting in decreased survivorship post-release (Tidwell, Shepherdson,
& Hayes, 2013). Due to Tidwell’s results, local ecologists involved in the R. pretiosa
recovery plan theorized that perhaps using mesocosms as the captive rearing environment
would have more positive benefits beyond the goal of increasing survival during
development.
Ecology is studied across varying geographic scales: from whole system scales to
mesocosms, to microcosms. Mesocosms are moderately sized (i.e., smaller than whole
ecosystem studies, bigger than microscopic ecosystem studies) human-made ecosystems
that are used as tools in ecological research—allowing a certain amount of control over
natural complexity through smaller scale, simplified studies. Scientists also use
13
mesocosms in applied research and educational development (Kangas & Adey, 1996).
They combine a technological component related to the form of container structures with
environmental management and control of boundary exchanges with living populations
(Kangas & Adey, 1996).
Despite the strong arguments in favor of rearing R. pretiosa in mesocosms, there
are still many facts we do not know about methodology to inform rearing protocols or
how individual frog will respond to the treatment. My research addressed the water
quality needs of rearing R. pretiosa in mesocosms. My primary research question was “is
there a difference in water quality parameters—pH, temperature, dissolved oxygen,
nutrient concentration—between a traditional captive rearing environment and a
mesocosm rearing environment. Secondarily, I explored the differences between two
captive rearing treatments and the natural prairie wetland environment.
Methods
Study Design
I collected water quality measurements and samples from three different rearing
habitats for R. pretiosa: 1) Traditional rearing habitat which is a regularly-cleaned tank
environment with no plant material, 2) Mesocosm rearing habitat which is a lab14
contained tank environment that
resembles the natural habitat through
the presence of wetland plants and 3)
Prairie wetland habitat which is a
natural habitat of R. pretiosa. The
traditional and mesocosm rearing
habitats were 300 gallon cattle tanks
Figure 1—the captive rearing tanks at Woodland
Park Zoo. Four tanks were Traditional rearing
environments (without any natural elements) and
four were Mesocosm rearing environments (which
included wetland plants collected from West Rocky
Prairie)
(see Figure 1) kept at Woodland Park
Zoo in Seattle, WA (hereafter referred
to as WPZ). The wetland habitat
sampled at West Rocky Prairie (hereafter referred to as WRP) consisted of four randomly
chosen points. There were 4 separate tanks of the Traditional and Mesocosm rearing
habitat, for a total of 8 tanks sampled each week at WPZ. To complete the sampling, 4
points were chosen at random in the WRP wetland to be the 4 replicates of the third
rearing environment; these same points were sampled during every sample collection.
Sample Collection
Samples were collected once per week for six weeks between July 26 and August
30, 2014. Woodland Park Zoo samples were collected every Saturday between 10:00am
and 12:00pm and West Rocky Prairie samples were collected every Sunday within the
sampling period between 10:00am and 12:00pm. During each sample collection the
following measurements were taken: pH, temperature, and Dissolved Oxygen.
Additionally 500ml water samples were collected in plastic (PP) sample lab bottles from
each replicate back to the lab for nutrient analysis. See Table 1 below.
15
Table 1. Sampling schedule for each of the three rearing environments for R. pretiosa. 500ml
samples were taken from each of the four replicates for each of the three treatments once per
week during the three weeks of sampling
Habitat Type
N (# of Replicates)
Times Sampled
Volume Sampled
Traditional
(Captive Rearing)
4
3
500ml
Mesocosm
(Captive Rearing)
4
3
500ml
Prairie
(Wild)
4
3
500ml
Lab Methods
All sample bottles were washed with hot, soapy water and rinsed three times with
DI (Deionized) water prior to field collection, to prevent sample contamination.
Additionally all lab equipment and glassware used for nutrient analysis (volumetric
glassware, vacuum filtration apparatus) were washed with hot soapy water and rinsed
with DI water prior to analysis. During the analysis process, the syringe to insert samples
into the Ion Chromatograph was washed with hot soapy water and rinsed with DI water
between every sample to prevent cross-contamination.
Materials List
For full materials list see Appendix A
Nutrient Analysis
The nutrient analysis portion of my research was conducted at The Evergreen
State College using a DIONEX IC25A ion chromatograph. I followed the EPA methods
300.1 (Plaff, 1993) as a guideline for testing nutrient concentrations in each sample.
Nutrient analyses were conducted during the same week samples were collected.
16
According to the EPA methods 300.1 (Plaff, 1993) the holding time for chloride and
sulfate is 28 days; for nitrate, nitrite and phosphate the holding time is 48 hours. Based on
these holding times, samples were stored at 4°C prior to analysis to maintain accurate
nutrient levels until analysis. Stock standards for most anions are stable for at least 6
months when stored at 4°C. Phosphate stock standards are only stable for 1 month
when stored at 4°C. Working standards for chlorine, nitrate and sulfate were prepared
once per month and the working standards for phosphate were prepared fresh on the day
of analysis in order to prevent instability of nutrient concentration.
Concentration/Dilution
McKibbin (2008) found these the nutrient concentrations in a local wetland during
sampling in British Columbia:
Nutrient
Chloride
Sulfate
Nitrate
Phosphorous
Concentration (mg/L)
1.03
0.5
0.5
0.04
There is a lack of published research on nutrient concentrations in water quality tests of
wetlands—thus I used these concentrations as a starting point for my analysis. These
determined the concentration of my High, Medium and Low Standards during the testing.
In the process of ion chromatography High, Medium and Low standards are created in an
effort to bracket the actual nutrient concentration in the sample being tested. These three
working standards give the ion chromatograph three calibration curves for each nutrient
17
being tested, giving it three points of reference when assessing the nutrient concentration
of the sample.
Preparing Standards
1. Stock Standards
a. Prior to creating stock solutions, each compound was placed in glass petri
dishes and put into an oven at 105°C for 24 hours. Compounds were then
transferred to a glass desiccator to cool and prevent moisture from reabsorbing into the solid compounds.
b. A pre-determined amount of each compound (see Table 2 below) was
added to a 1L volumetric flask, DI water was then added for a total
volume of 1L. Next, each stock standard was poured into a specific 1L
plastic bottle and mixed well by shaking.
Table 2-Amount (g) of each analyte added to deionized water to create stock standards at a concentration of
1000mg/L. These amounts were taken from the EPA methods 300.1 (Plaff, 1993) for detecting nutrient
concentrations in drinking water
Analyte Stock
Chloride
Sulfate
NO 3 --N
PO 4 3--P
Concentration
1000 mg/L
1000 mg/L
1000 mg/L
1000 mg/L
Weight of Compound (g)
Final Volume (L)
0.1649g Sodium Chloride
1L
0.1814g Potassium Sulfate
1L
0.606g Sodium Nitrate
1L
0.4394g Potassium
1L
Dihydrogenphosphate
c. Use Stock Standards to make mixed intermediate stock standards:
a. Mixed intermediate stock standards were made by adding 10mL
each of chloride, nitrate, phosphate and sulfate stock standard
solution to a 1L volumetric flask and bringing to a volume of 1L
18
with DI water. Intermediate stock standards were used to make
working standards because the stock standards, as instructed by
EPA methods 300.1 (Pfaff, 2003) were made at much higher
concentrations than what was being tested for and it made the
creation of the working standards more accurate.
d. Quality Control Stock Standards, Intermediate Standards and Working
Standards were made following the aforementioned methods, using
different compound stocks.
2. Working/External Standards
Use the following pre-determined amount of the mixed intermediate (chloride, nitrate,
sulfate and phosphate) stock standard solution to create the specified mixed Low,
Medium and High working standards. Add calculated volume of mixed intermediate
stock standard for each analyte to a 500 mL volumetric flask and bring to volume with DI
water. Then transfer to a clean 500 mL plastic bottle for use in analysis.
19
Analyte
Chloride
Sulfate
NO 3 --N
PO 4 3--P
Analyte
Chloride
Sulfate
NO 3 --N
PO 4 3--P
Analyte
Chloride
Sulfate
NO 3 --N
PO 4 3--P
MIXED LOW STANDARD
Concentration
Volume Needed
(mg/L)
from Int. Stock
(mL)
0.50
25
0.20
10
0.10
5
0.02
1
Total: 31mL
MIXED MEDIUM STANDARD
Concentration (mg/L)
Volume Needed
from Int. Stock
(mL)
1.0
50
0.7
35
0.5
25
0.06
3
Total: 113mL
MIXED HIGH STANDARD
Concentration (mg/L)
Volume Needed
from Int. Stock
(mL)
1.5
75
1.2
60
1.1
55
0.2
10
Total: 200mL
Final Volume(mL)
500mL
Final Volume(mL)
500mL
Final Volume(mL)
500mL
Running Samples through Ion Chromatograph
The ion chromatograph components (oven, eluent generator and chromatograph
unit) were turned on 2 hours prior to running samples to allow the various readings to
stabilize. Samples were also taken out 2 hours prior to running to allow them to reach
room temperature. This was an important step to ensure accuracy during analysis. The
20
syringe and filter apparatus was washed with hot, soapy water, rinsed with DI water and
then rinsed three times with the sample being run for “pre-contamination” This last step
was important to ensure that no one sample contaminated another sample reading.
Syringe with approximately 2 milliliters was then loaded and injected into the ion
chromatograph for analysis. While one sample was running, the next sample was
prepared for injection following the same protocols previously stated. See Appendix B
for full Run List
Statistical analysis was completed using non-parametric tests as the data set did
not pass preliminary tests for normality and spread.
Results
pH, Dissolved Oxygen, Temperature
All three parameters measured in real time with probe instruments demonstrated a
statistically significant difference between both the captive rearing treatments (Mesocosm
and Traditional) and the Prairie wetland habitat. For pH, the data were mostly consistent
in all three treatments across the full six weeks (p=0.0313, df=2), see Figure 2.
Though the dissolved oxygen and temperature results showed more variability
across the sampling period, both parameters showed a statistically significant difference
between the captive rearing treatments (Mesocosm and Traditional) and the Prairie
wetland habitat (DO: p=0.313, df=2; Temp: p=0.313, df=2). For dissolved oxygen, see
Figure 3, for temperature see Figure 4.
21
pH
8.3
pH
7.8
7.3
6.8
6.3
8/2/2014 8/9/2014 8/16/2014 8/23/2014 9/6/2014 9/13/2014
Sampling Date
Mesocosm pH
Traditional pH
Prairie pH
Figure 2—pH measurements for each rearing habitat (Traditional, Mesocosm and
Prairie) taken over the sampling period of 6 weeks. Mesocosm and Traditional pH
readings differed significantly from the Prairie habitat (p=0.313, df=2).
Dissolved Oxygen (mg/L)
Dissolved Oxygen
10.0
9.0
8.0
7.0
6.0
5.0
4.0
3.0
2.0
1.0
8/2/2014
8/9/2014
8/16/2014
8/23/2014
9/6/2014
9/13/2014
Sampling Date
Mesocosm DO
Traditional DO
Prairie DO
Figure 3—Dissolved oxygen measurements for the three rearing habitats across the
sampling period. Statistical analysis showed a significant difference between both
captive rearing habitats (Traditional and Mesocosm) and the natural Prairie habitat
(p=0.313, df=2)
22
Temperature
Temperature (C Degrees)
24
22
20
18
16
8/2/2014
8/9/2014
8/16/2014
8/23/2014
9/6/2014
14
9/13/2014
Sample Date
Mesocosm Temp
Traditional Temp
Prairie Temp
Figure 4—Temperature measurements across the sampling period. Statistical
analysis showed a significant difference between both captive rearing habitats and
the natural Prairie habitat (p=0.313, df=2)
Nutrient Analysis
All four nutrients measured showed significant differences between the treatments
during the three sampling weeks. Phosphorous showed the most significant difference
between the rearing environments across the sampling period. Week 1 of sampling
showed a significant difference between the Mesocosm and Traditional rearing
environments (p=0.029, df=2) as well as between the Mesocosm and Prairie rearing
environments (p=0.019, df=2). Week 2 of sampling showed a significant difference
between both captive rearing treatments (Mesocosm and Traditional) and the Prairie
habitat (p=0.023, df=2; p=0.025, df=2, respectively). See Figure 5.
23
Figure 5—Phosphorous concentrations across the three sampling weeks for each
rearing habitat. Here the data shows that there was a difference found between the
Prairie wetland habitat and the two captive rearing habitats, statistical analysis
determined that there was a statistically significant difference between both captive
rearing treatments and the Prairie habitat (p=0.026).
Seen in more detail in Table 3, we can see the distinct difference in phosphorous
concentration between the three rearing habitats for R. pretiosa.
Table 3—Data representing the weekly averages for the phosphorous concentration in each of the three
rearing habitats (Traditional, Mesocosm and Prairie wetland). Though no significant differences were found
during statistical analysis, the data to reveal a strong difference between the natural prairie habitat and the
two captive rearing habitats.
Treatment
Mesocosm
Traditional
Prairie
Average Week 1
0.51075
0.32725
8.1845
Average Week 2
1.87775
2.28425
6.484
Average Week 3
2.601
2.43075
6.74575
Nitrate concentrations found in the three rearing habitats were on average much
higher than all other nutrients measured, and showed significant differences between the
24
Mesocosm and Prairie rearing environment. Week 1, week 2 and week 3 sampling
revealed a significant difference between the Mesocosm captive rearing treatment and the
Prairie wetland habitat (Week 1: p=0.029, df=2; Week 2: p=0.028, df=2; Week 3:
p=0.029, df=2).
Figure 6—The nitrate concentrations across the three sampling weeks for the each of
the three rearing habitats (Traditional, Mesocosm and Prairie). Statistical analysis
revealed significant differences between the Mesocosm and Prairie environment for
all three weeks of sampling
Additionally the Traditional rearing habitat showed higher nutrient concentrations
over all three weeks of sampling when compared to the Mesocosm and Prairie habitat.
Nitrate concentrations also showed the most variation from week to week compared to
the other nutrients measured (see Figure 6, Table 4).
Table 4—Average data per sample week for each rearing habitat (Traditional, Mesocosm and Prairie). These
data, though statistical analysis did not reveal any significant difference, show what appears to be a trend that
may be discovered after additional sampling.
Treatment
Mesocosm
Average Week 1
7.188666667
Average Week 2
4.676333333
Average Week 3
3.103666667
25
Traditional
Prairie
2.698
0.55475
4.2545
1.2505
1.957
0.52
Chloride concentrations also showed significant differences between the captive
rearing treatments (Mesocosm and Traditional) and the Prairie habitat. In Week 2 of
sampling, chloride concentrations differed significantly between the Mesocosm captive
rearing treatment and the Prairie wetland (p=0.033, df=2) as well as between the
Traditional captive rearing treatment and the Prairie wetland (p=0.031, df=2). Similarly
in Week 3 of sampling, chloride concentrations significantly differed from both captive
rearing treatments and the Prairie wetland habitat (Mesocosm: p=0.026, df=2;
Traditional: p=0.026, df=2). See Figure 7, Table 5.
Chloride Concentration (mg/L)
Chloride
35
30
25
20
15
10
5
0
Average Week 1
Average Week 2
Average Week 3
Sampling Week
Mesocosm
Traditional
Prairie
Figure 7—Chloride concentrations for each of the three rearing habitats across the
three sampling weeks. This data shows that the Prairie wetland habitat had
consistently higher concentrations of chloride, despite statistical analysis not
revealing a significant difference between the mean values of each treatment.
26
Table 5—Average weekly values for the chloride concentrations in each of the three rearing habitats for R.
pretiosa. Though statistical analysis did not reveal any significant differences or relationships, the data indicate
that there are differences between the captive rearing environments and the natural Prairie environment.
Further investigation may reveal more statistically significant relationships.
Treatment
Mesocosm
Traditional
Prairie
Average Week 1
6.99625
5.03125
6.635
Average Week 2
10.6985
10.931
23.377
Average Week 3
9.898
10.0795
23.379
Sulfate concentrations showed significant differences in the rearing habitats only
one of the three sampling weeks. Week 1 of sampling showed a significant difference in
sulfate concentrations between the Mesocosm and Traditional rearing treatments
(p=0.046, df=2). This is the only nutrient concentration that did not show significant
difference between the two captive rearing treatments (Mesocosm and Traditional) and
the Prairie wetland habitat, however as seen in Figure 8 and Table 6, there appear to be
more trends in the data than those that were detected by statistical analysis.
Sulfate Concentration (mg/L)
Sulfate
3.5
3
2.5
2
1.5
1
0.5
0
Average Week 1
Average Week 2
Average Week 3
Sampling Week
Mesocosm
Traditional
Prairie
Figure 8—The concentrations of sulfate found in each of the three rearing
environments. Each rearing habitat had 4 replicates whose results were averaged for
analysis. Week 1 showed a significant difference in the sulfate concentrations of the
Mesocosm and Traditional captive rearing treatment (p=0.046, df=2)
27
Table 6—Average sulfate concentrations for each of the three rearing environments
across the three sampling weeks. *0 mg/L average was detected in the Mesocosm sample
from week 2, however this may only indicate a sulfate concentration below the Ion
Chromatograph’s detectable range.
Treatment
Mesocosm
Traditional
Prairie
Average Week 1
1.15075
0.427
0.068
Average Week 2
0*
0.654
1.954
Average Week 3
0.203
0.12875
2.404
Discussion
pH is reflective of the hydrogen ion concentration in water and is a key
characteristic in any aquatic habitat (Dodd, 2010). Acidic water affects survivability of
juvenile and adult frogs, specifically disrupting the ionic balance within cells during
developmental (Zug, 2001). Studies have reported a pH range of 6.0-7.5 to be neutral or
at least within a range that should not harm organisms living in that aquatic environment
(Dodd, 2010). The pH levels observed in this study, unsurprisingly, stayed right around
7.5 with slight fluctuation for both the Traditional and Mesocosm rearing treatments.
These levels were maintained by frequent water changes (approximately every 3 days)
which prevented organic build-up, which could have made the tanks more acidic. Low
pH (or high acidity) can adversely affect osmoregulation in larval amphibians by
substantially increasing rates of sodium loss through the skin (Wells, 2007); therefore, it
is critical to maintain healthy pH levels in future mesocosm tanks for amphibian rearing.
For example, the urea excreted by the frogs in the captive rearing tanks can make the
tanks highly alkaline if the water is never changed, therefore we need to find a balance of
a self-sustaining mini ecosystem and healthy water quality parameters for the frogs.
28
Given that the captive rearing tanks were kept within the pre-determined “pH neutral”
bracket, it is curious that the Prairie wetland habitat pH readings would differ
significantly from the both captive rearing treatments across the sampling period
(p=0.313, df=2). Could this indicate high adaptability by Rana pretiosa in both its native
and simulated environment, or are these reading reflective of the time of year during
which sampling occurred? More research is needed to fully explore the pH limits of R.
pretiosa and what those individuals will need during their captive rearing treatment,
especially considering that those in captive rearing have no ability to move to different
water conditions like those in the wild.
Aquatic environments can present less favorable conditions for oxygen uptake
and oxygen levels are often more variable than terrestrial environments (Wells, 2007).
Additionally, the oxygen content of closed bodies of water (i.e. captive rearing tanks) are
greatly affected by the respiration of aquatic organisms and even daily fluctuations in
respiration can result in major changes in the availability of oxygen (Wells, 2007).
Previous studies have indicated that under laboratory conditions, oxygen concentrations
below 4mg/L are deemed stressful to amphibian larvae and aquatic adults (Dodd, 2010).
However, each species is different and may have the ability to acclimate to varying
conditions depending on its niche in the local ecosystem (Dodd, 2010). In both the
captive rearing treatments—Traditional and Mesocosm—the dissolved oxygen hovered
between 8 and 9mg/L. The dissolved oxygen readings of both captive rearing treatments
differed significantly from the Prairie wetland habitat (p=0.323). The average dissolved
oxygen measurements for the Prairie wetland habitat had higher variability from week to
week; starting around 6mg/L and ending closer to 2mg/L (see Figure 3 on page 22). The
29
higher variation and on average lower dissolved oxygen values found in the Prairie
habitat are likely due to the increased decaying plant matter and lowering water levels.
Though the sampling period was only for six weeks during the summer, the Prairie
wetland habitat demonstrated a much higher variability than both the captive rearing
environments. These results could indicate that R. pretiosa has become adaptive to
varying dissolved oxygen concentrations as a result of leading an entirely aquatic
existence, or that perhaps all three treatments would show different dissolved oxygen
fluctuation patterns given a longer sampling period.
One thing to consider when considering the dissolved oxygen results is that
dissolved oxygen levels vary throughout the day depending on photosynthetic rates in
response to local weather patterns. As such, dissolved oxygen concentrations in ponds are
often lowest at dawn and increase throughout the day as the plants begin to
photosynthesize (Dodd, 2010). This indicates the possibility that though dissolved
oxygen readings appeared to be decreasing over the sampling period, it could have been
linked to the amount of photosynthetic activity prior to sampling as all data was collected
between 10am and 12noon. Regardless of fluctuations, though the captive rearing
habitats—Traditional and Mesocosm—differed significantly from the Prairie ecosystem,
they did support healthy growth throughout their captive rearing period. This indicates
that R. pretiosa seems well adapted to fluctuating dissolved oxygen concentrations in
captivity and in the wild, and therefore would not be oxygen-inhibited in a mesocosm
environment under these conditions.
It is crucial to consider, however, that in a true mesocosm environment (i.e. one
with less frequent water changes that more closely resembles a self-sustaining ecosystem)
30
oxygen levels may become lower due to the accumulation of organic matter. For
example, hypoxia can be fairly severe in smaller bodies of water due to the respiration of
organisms quickly depleting the available O 2 supply (Wells, 2007). As a result of
potentially hypoxic conditions, the metabolic rates of developing larvae or tadpoles could
decline as a result of decreasing oxygen availability, which could inhibit healthy growth
(Wells, 2007). Regardless of any possibility of an amphibian species ability to adapt to a
less oxygen-rich mesocosm environment, it will be important to monitor each species
during captivity to ensure their health is not declining as a result.
Temperature also is a key factor in providing healthy growing habitat for
amphibians in captive rearing environments. For ectotherms—such as frogs—
temperature may be the single most important physiological variable because all cellular
processes are temperature dependent (Zug 2001). Temperature influences the amphibians
directly as well as other biological processes in their environment such as pH and
dissolved oxygen concentration. In the case of this thesis research, temperature was not a
limiting factor in the captive rearing environments as it was consistently regulated to be
within a healthy range for the developing frogs. However, as indicated in the results
(Figure 4, page 23) we see that the Prairie ecosystem showed a decrease in temperature
across the sampling period. This is likely in response to the nighttime temperatures
dropping with the approach of fall. Though the frogs developing in the Prairie ecosystem
would have more freedom of movement than in captive rearing, this difference in
temperature between the three rearing habitats could indicate a wider range of
adaptability by R. pretiosa that could be explored in future captive rearing habitats.
31
Phosphorous is an essential plant nutrient but excessive amounts can cause water
quality to deteriorate (Stewart, B. A., & Howell, T. A., 2003). The results indicate that the
Prairie habitat had consistently higher concentrations of phosphorous than did either of
the captive rearing treatments. This difference was further supported by the statistically
significant difference found among the nutrient concentration data: there was a
statistically significant difference between both the captive rearing treatments and the
Prairie habitat all three weeks. Wetland environments, as the local weather gets warmer
and drier from spring through summer, show changes in nutrient concentration as well as
other water quality parameters such as dissolved oxygen and pH. These changes are a
result of increased water evaporation leaving little standing water by the end of the
summer months. This pattern was also observed during sampling at West Rocky Prairie
between August and September—very little water remained in the large wetland. It has
also been documented that in Pacific Northwest wetlands, concentrations of nutrients
such as nitrate and phosphorous are much higher in sediments and belowground biomass
than in overlying waters and aboveground plant tissues (Washington State Department of
Ecology, 1986). This may be an explanation behind the increased levels of phosphorous
in the Prairie environment as samples were taken at the end of the summer when there
was a significantly less amount of water in the wetlands which could have been more
influenced by nutrient concentrations in the sediments.
Nitrate concentrations were consistently highest in the Mesocosm rearing habitat.
The Prairie habitat having the lowest concentrations of nitrate across the sampling period
was unexpected due to it being a natural environment and more influenced by
groundwater runoff and biological accumulation. The nitrate concentrations of the
32
Mesocosm captive rearing treatment differed significantly from the Prairie wetland
habitat during all three sampling weeks, and showed the Mesocosm environment having
consistently higher nitrate concentrations over the Prairie wetland habitat. Nitrates are
readily absorbable and are quickly taken up by plant material. However the plants in the
Mesocosm environment were not adapting to their new environment very well and as
such were not performing (photosynthesizing, respiring, taking up nutrients) at maximum
capacity and therefore not absorbing the nitrates in the water very quickly. This combined
with the frogs excreting urea explains the increased nitrate levels in the Mesocosm
environment. Several studies have shown negative impacts of increased ammonium
levels (a nitrogen containing waste product excreted by most aquatic amphibians (Wells,
2007)) on amphibian species. For example a study conducted in Oregon by Adolfo
Marco, Consuelo Quilchano & Andrew R. Blaustein (1999) observed that prolonged
exposure to higher concentrations of nitrates proved fatal to R. pretiosa. Additionally, a
similar study from Spain demonstrated a reduction of growth rate as a typical direct effect
of ammonium nitrate on amphibian larvae (Ortiz-Santaliestra, et al. 2012). In contrast, the
results of this pilot study demonstrated that R. pretiosa can survive well in the captive
rearing treatments which had much higher nitrate concentrations than their natural prairie
environment showed. This could indicate, similar to the other nutrient concentrations, R
pretiosa’s ability to adapt the varying nutrient concentrations that the levels of nitrate
may have had a lower overall average across the entire captive rearing period. More
research needs to be conducted into the nitrate concentrations associated with captive
rearing mesocosms.
33
Chloride ions, along with sodium and potassium ions, are a part of aquatic
amphibian’s active ion uptake to maintain electroneutrality between themselves and their
environment (Wells, 2007). Ions are also reabsorbed within the kidneys to prevent large
ion loss and thus increase the need to actively intake these ions from their surrounding
environment. There is little research completed on specific needs of amphibians in
captive rearing in terms of ion concentrations, however like any organic element within
an ecosystem, moderate concentrations—not too low, not too high—will most likely be
the best suited for amphibian captive rearing. However, this also varies by species. The
chloride concentrations were consistently higher in the Prairie habitat across the sampling
period. A study conducted in British Columbia (McKibbin et al., 2008) examined water
quality as parameters for embryonic survivorship in three known R. pretiosa populations.
They measured chloride levels between .063 mg/L and 1.83 mg/L across three sampling
sites. In comparison, these rates seem rather low to the chloride levels detected during
this thesis research, however their research was conducted earlier in the year and only
one sample per year was collected for analysis. In addition, the chloride levels in the
Prairie wetland habitat were higher due to their natural presence accumulating as the
water levels dropped toward the end of the summer. There is a significant lack of
published information regarding the nutrient levels of wetland ecosystems and what one
can deem as “normal”; as such, it is difficult to interpret what my thesis results may
indicate.
Sulfate concentrations were highly variable for all three rearing habitats across the
sampling period. The Prairie environment showed the highest concentrations in the final
two weeks of sampling and the Mesocosm rearing treatment showed the highest sulfate
34
concentration during the first week of sampling. Though there was only on significant
difference detected during the sampling period—Week one showed a significant
difference in sulfate levels between the Mesocosm and Traditional captive rearing
treatments (p=0.046)—as we can see on Figure 8 on page 27 there appears to be a large
difference in sulfate concentration between the captive rearing habitats and the Prairie
habitat. Changes in concentrations of nutrients such as sulfate can vary dramatically over
the course of amphibian breeding and rearing season (Gerlanc & Kaufman, 2005). More
research is needed to determine what specific effects these changes might have on
amphibians in captive rearing environments.
This thesis research was part of a pilot study in partnership with Woodland Park
Zoo in Seattle, WA. The sampling period was not ideal in length, and as such the data
results may be skewed or not show an accurate account of the water quality parameters of
each of the rearing habitats. Additionally, the top priority during this captive rearing pilot
project was the health of the developing frogs. As such the caretakers at WPZ did weekly
tests and changes to the aquatic environments—both Traditional and Mesocosm
treatment—to ensure the continued health of the growing frogs. Tests for ammonia levels
were conducted once per week, as well as visual checks for algae accumulation which
may have inhibited the ability for the keepers to see the frogs clearly—to determine if
they were in distress. If ammonia levels were outside healthy standards or if an algal mat
had formed inhibiting visual scope of the animals, the water in the tank was changed. It
was the original goal of my thesis research to test the differences in water quality
between the Traditional treatment and the Mesocosm treatment, however frequent water
changes in both treatments may have skewed my ability to accurately describe the
35
differences between these two environments. For example, based on the log of water
changes supplied by Woodland Park Zoo, the water in both the Traditional tanks and
Mesocosm tanks was changed roughly every 3 days. This was done to maintain the health
of the animals during development, but may have inadvertently skewed the results. The
data gathered are still important in beginning to describe baseline conditions for future
mesocosm and captive rearing aquatic environments.
Captive Rearing Management Conclusions and Recommendations
The results of this pilot project were not as I had originally imagined, however
they provide an interesting baseline for gaining insight into the inner workings of using
mesocosms to captive rear amphibians and how closely those mesocosms need to
resemble the natural environment or how closely they can resemble traditional captive
rearing tanks as well. As we have seen from the results here, the water quality
parameters in the captive rearing treatments (Mesocosm and Traditional) differed from
the Prairie environment—some statistically significant, some not—and these differences
open up a wide array of new questions that need to be answered.
Do the differences in the captive rearing treatments and the Prairie ecosystem
indicate that the captive rearing environments could be more constant or less variable in
terms of water quality parameters? For example, West Rocky Prairie houses a known
Rana pretiosa population that is able to survive and reproduce every season in these
conditions that were identified during this research project. If they can survive in current
conditions, would it be more beneficial to expose captive-reared tadpoles to varying
36
conditions to closer simulate the natural environment? What is the balance of selfsustaining mesocosm environments and continued health of the growing frogs? These are
all questions that should be addressed with future research.
I set out on this project to begin to understand whether or not mesocosms were a
good idea for the captive rearing of R. pretiosa. Based on the data gathered, mesocosms
show amazing potential for captive-reared individuals of all amphibian species,
particularly those like R. pretiosa whose numbers are quickly declining. Currently,
however, more information must be gathered before we can take true strides forward in
adding mesocosms to the captive rearing plan for R. pretiosa. Based on the results of this
thesis research, there is still a lot left to interpretation in terms of explaining the
difference between the Prairie environment and the captive rearing treatments. There is a
lack of research describing what is “normal” in any given wetland, and as such it may be
difficult to pinpoint and replicate precisely what the frogs require while in captivity.
Suggestions for future research include looking into mesocosms as captive rearing
environments for threatened/endangered amphibian species in the Pacific Northwest,
specifically by examining biochemical cycles within a mesocosm environment across the
entire captive rearing period (roughly February through September). It is recommended
that future studies work to keep the water changes for mesocosms notably different from
the control rearing environment so as to observe a more clear difference for all
parameters being monitored. Mesocosms show amazing promise in the world of
amphibian captive rearing, given continued research to assess how captive rearing
environments can promote the rearing frogs that can survive and thrive once released into
natural environments.
37
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APPENDICES
Appendix A: Materials List
Materials List
Ion Chromatograph (Model: DIONEX IC25A), Plastic (PP) Sample Bottles (500
mL), Volumetric Pipettes (1, 1mL pipette, 2, 5mL pipettes, 1, 10mL pipette, 1, 15mL
pipette, 4, 25mL pipettes, Bulb), Millipore Filtration Apparatus (47mm, 500mL
capacity), 1 liter vacuum filtration flask, 1, 500 mL vacuum filtration flask, 70, 47mm
0.45 micropore filters, 1 rubber stopper with two openings, 2 vacuum rubber hoses, 2
ring stands, 2 ring clamps: large enough to hold 1L and 1 500mL vacuum filtration
flasks, 1 small rubber hose “pincher”, 0.2 micrometer syringe filters for the IC, Lab
Tape for labeling supplies, Glass Dessicator, 8-100mL beakers, 4 spatulas, 10 1L
plastic bottles, 4-1L volumetric flasks, 5-500mL volumetric flasks, 12, 1L plastic
bottles for individual stock standards, mixed intermediate standards and mixed
working standards, Analytical Balance for measuring salts, Chem-grade cooler for
cold transport of samples for preservation
Standard Preparation Ingredients:
Nutrient Testing For
Chloride
Ingredients Needed to Make Standard Soln. (EPA Method 300.1)
1000mg/L: 0.1649g sodium chloride (CASRN 7647-14-5)
44
Sulfate
NO 3 --N
PO 4 3--P
1000mg/L: 0.1814g potassium sulfate (CASRN 7778-80-5)
1000mg/L: 0.6068g sodium nitrate (CASRN 7631-99-4)
1000mg/L: 0.4394g potassium dihydrogenphosphate (CASRN 7778-770)
Appendix B: Full Run List for Ion Chromatography Analysis
Run List
1. Rinse Blank
2. Second Rinse Blank (must match first rinse blank before proceeding)
3. Standards (Low to High)
a. This generates the calibration curve
b. Check the R2 (EPA standard R2 value = 0.9998) to judge the fit of the line
4. Rinse Blank
5. Quality Control
a. +/- 5% Retention Time
b. +/- 15% Concentration
c. Compare these first quality control values to 1) medium working standard
retention times and 2) lab notebook concentrations for Quality Control
standards.
6. Up to 10 samples
7. Quality control
a. +/- 5% Retention Time
45
b. +/- 15% Concentration
c. **Compare these values to first quality control. If they do not match
within the approved interval the previous samples run are null**
8. Up to 10 more samples
9. Quality Control
a. +/- 5% Retention Time
b. +/- 15% Concentration
**Compare these values to first quality control. If they do not match within the approved
interval
46
