PHYSIOLOGICAL RESPONSES BY HALODULE WRIGHTII ASCHERS. (SHOAL
GRASS) TO THE INTERACTIVE EFFECTS OF HYPOSALINITY AND NITROGEN
prepared by
JOSEPH L. KOWALSKI
FEBRUARY, 2015
for
The Graduate Committee
Marine Biology Program
Department of Life Sciences
Texas A&M University-Corpus Christi
Corpus Christi, Texas
Approved: ____________________________________________
Dr. Kirk Cammarata, Co-Chairperson
____________________________________________
Dr. Paul Zimba, Co-Chairperson
____________________________________________
Dr. Paul Montagna, Member
____________________________________________
Dr. Hudson DeYoe, Member
____________________________________________
Dr. Warren Pulich, Member
____________________________________________
Dr. Joe Fox, Department Chairperson
Format: Estuaries and Coasts
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Project Summary
Halodule wrightii Ascherson (shoal grass) is a widely distributed tropical to subtropical
colonizing seagrass species that exhibits broad tolerance to salinity extremes. In
hypersaline environments salinity stress can affect its metabolism, physiology,
photosynthetic performance, and uptake and utilization of nitrogen. Most studies on
seagrass response to salinity stress have focused on hypersalinity effects. Few studies
have examined the effects of hyposalinity on H. wrightii, and none have assessed its
effects on metabolism and physiology of this species. The proposed research will test
how H. wrightii responds to and persists during periods of hyposalinity. Hyposalinity is
expected to decrease growth and induce stress responses. Respiration rates in leaf and
rhizome/root organs are expected to increase as a stress response during osmotic
adjustment, and tissue concentrations of principle ions and free amino acids are expected
to correlate to the degree of hyposalinity. Because of the energy expenditure for osmotic
adjustment, uptake rates of inorganic nitrogen by both stressed leaves and roots are
expected to be less than that of a control, and thus sediment amendments of inorganic
nitrogen under hyposalinity are expected to have no stimulatory effect on growth rates or
biomass of H. wrightii. Three laboratory experiments are proposed to test these
expectations at three lower salinities (20, 10, and 5) relative to that of a control (35).
Intellectual Merit
Morphometric leaf growth measurements, PAM fluorometry, and tissue
respiration rates will provide an integrated understanding of how H. wrightii responds to
hyposalinty stress, nutrient stress and a combination of the two. Furthermore, it utilizes
physiological assays of osmolytes (organic and inorganic) to assess plant response to
hypoosmotic stress. Collectively, these techniques will provide predictive capacity to
estimate the effects of freshwater discharge on H. wrightii populations under the
influence of stormwater discharge drainage channels. This study also opens a new avenue
of research into differences that may exist between the more dynamic water column
(increased flow) and the sediment porewater environment, and how these putative
differences may affect plant function and growth when they are not in equilibrium. Since
H. wrightii is a colonizing species, this study also lays a foundation for future
experiments that could provide insight into the autecology of the stenohaline seagrass
species, Syringodium filiforme (manatee grass) and Thalassia testudinum (turtle grass).
Broader Impacts
This study will provide me with opportunities for development as an independent
investigator and educator. It will also allow for direct and indirect benefits to students in
underserved South Texas through involvement with the proposed research and by
teaching and mentoring them as developing scientists through competitive research
projects in future efforts. Furthermore, this project will provide insight into application of
appropriate resource management strategies and/or actions to protect seagrasses, a critical
and protected resource for marine habitat. Accordingly, this proposed work will advance
habitat restoration and preservation of coastal resources.
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Background and Relevance
Plant stress has been broadly defined as factors that impede plant growth and
reproduction below the potential of the genotype (Osmond et al. 1987). Examples include
extremes of temperature (Smith et al. 1984; Li et al. 2014), herbivory (White 1984),
excess light (Demmig-Adams and Adams 1992), and exposure to heavy metals (Gratão et
al. 2005). Water-related stressors include drought conditions (Shinozaki and YamaguchiShinozaki 2007), which induces salt stress (Osmond et al. 1987), and flooding (Jackson
and Colmer 2005; Mommer and Visser 2005), which causes hypoxia. All plants endure
varying levels of stress, although each type may respond uniquely. When stress occurs,
physiology and metabolism are often affected, usually in proportion to the stress applied
(Osmond et al. 1987). Because it is uniquely tied to plant function, photosynthesis is
often compromised and is directly related to the activity of other environmental stressors.
For example, hypersaline conditions increased dark respiration rates up to 98% in the
seagrass Posidonia oceanica, but was also related to a decrease of 13 to 33% in
photosynthetic rates and decreased leaf growth rates (Marin-Guirao et al. 2011). Growth
and photochemical efficiency decreased, and expression of antioxidant enzymes
(catalase, superoxide dismutase, glutathione reductase), and total soluble proteins
increased under suboptimal saline conditions of 10 and 60 in Ulva prolifera (Luo and Liu
2011). Desiccation stress in resurrection plants (Selaginella lepidophylla) forms reactive
oxygen species which damages the photosynthetic apparatus (Dinakar et al. 2012) and is
associated with increase in metabolic (respiration) and physiological activity (Rejeb et al.
2014). Factors such as high concentrations of heavy metals, pathogens, and reactive
oxygen species can negatively affect photosynthetic performance and chlorophyll
fluorescence in many aquatic magnoliophyta species (Ferrat et al. 2003). In the brown
macroalgae, Ascophyllum nodosum and Fucus vesiculosus, increased rates of respiration
occurred immediately and were directly proportional to the degree of exposure to low
salinity (Tropin et al. 2003). Additionally, these plants experienced disruption and lysis
of several organelles and decreases in protein synthesis. Photosynthetic efficiency
decreased by 88% within 1.5 hr after exposure to a salinity of 10 in the macroalga
Caulerpa taxifolia (Theil et al. 2007).
Osmoregulation through adjustments in intracellular solute concentrations has
been documented across several marine plant divisions, from algae (Dickson et al. 1987;
Kirst 1989; Rani 2007), to emergent vascular species (Tuffers et al. 2001; Parida et al.
2004) to submersed species (Touchette 2007). Common organic osmolytes identified are
proline, glycine betaine and carbohydrates (Touchette 2007). Rani et al. (2007) found
concentrations of both proline and glycine betaine in direct proportion to salinities of 10,
35 and 65, respectively, in Cladophora vagabunda. Accumulation of organic solutes is
an osmoregulation mechanism documented in seagrasses (Parida and Das 2005;
Touchette 2007) and algae (Reed 1989). Posidonia oceanica and Cymodocea nodosa
each osmoregulate under hypersalinity by accumulating organic solutes, such as proline
and carbohydrates (Sandoval-Gil et al. 2014). Pulich (1986) found that proline was used
by H. wrightii, T. testudinum, and Ruppia maritima as an osmoticum while alanine was
also used in Halophila engelmanni under hypersalinity stress. He suggested the metabolic
pathways to produce these, and other amino acids, may be tied to control of high
concentrations of ammonium. Specifically, amino acids may be used in the synthesis of
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proline, thought to be important in osmoregulation. Similarly, Jiang et al. (2013)
suggested that nitrate metabolism, through regulation of nitrate reductase activity in
Thalassia hemprichii, may be responsible for regulation of leaf non-structural
carbohydrates concentrations under hyposaline conditions. As nitrate reductase activity
increased (under nirate-enriched conditions), non-structural carbohydrates were
consumed, decreasing osmolytic concentrations.
Transport of inorganic ions is also used by plants in response to environmental
changes in osmotic pressure (Parida et al. 2004). Tomato plants increased intracellular
concentrations of organic acids and Na+, K+, and Cl- as an osmoregulatory response to
increased soil salinity (Wang et al. 2011). Simulation of more frequent and prolonged
tidal flooding under greenhouse conditions were made on the upland ecotype of Spartina
densiflora and was compared to lowland plants which experience daily tidal flooding.
Upland plants had decreased concentrations of Cl-, increased leaf concentrations of K+,
increased root aerenchyma, as well as decreases in leaf mass, blade size and shape, and
decreases in chlorophyll fluorescence and quantum efficiency (Di Bella et al. 2014).
Results suggested that upland plants are not well-acclimated to increased frequency of
salinity perturbation through increased tidal flooding frequency. The mangrove,
Avicennia marina, had greater leaf concentrations of the inorganic ions, Na+, K+,
Ca2+,Mg2+, Cl-, at a salinity of 30 compared to a low salinity site (< 12), and lower
photosynthetic efficiency at the low salinity site (Tuffers et al. 2001). Jagels (1983) and
Babourina and Rengel (2010) reported the plasma membrane of epidermal cells of the
seagrasses R. maritima, T. testudinum and Zostera marina have abundant ATPases and
mitochondria and suggested that these are used for ion transport through a Na+ efflux/K+
uptake by way of a Na+/H+ antiporter where K+ is substituted for H+. They report that
such a system exists in freshwater plants. These studies document that inorganic ions play
a key role in plant physiology under a range of stressful conditions. Shabalaa and
Pottosina (2014) also describe how K+ can be considered a “master switch” in cell
physiology that works through selective transport channels in the plasma and vacuolar
membranes to maintain optimum plant performance.
Although there is a general positive correlation between salinity and cellullar
inorganic ion concentration, some studies do not completely follow this trend. Kirst
(1989) reported stimulation of photosynthesis in some macroalgal divisions, primarily in
the Phaeophyta, while photosynthesis was inhibited in several Chlorophyta, Phaeophyta,
and Rhodophyta species by both hypo-osmotic and hyper-osmotic conditions, and that
K+, Na+, and Cl- concentrations were positively affected. The same trends for selective
regulation of Na+ and K+ were also found for the seagrass, Z. marina (Rubio et al. 2011).
Mg2+ and Ca2+ concentrations were less affected, perhaps because they are used as
messengers. Hessini et al. (2009) found that sugars and other organic compounds
(proline) in Spartina alterniflora were more important to osmoregulation under water
stress than inorganic ions.
Aquaporins are a family of transmembrane channel proteins found in plasma and
vacuole membranes that are used for the passive movement of water down a water
potential gradient and operate in response to water stress (Kjellbom et al. 1999). Some
aquaporins are linked to transport of CO2 and metalloids, while others act in association
with ion gates and the regulation of ammonium uptake in terrestrial plants (Li et al.
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2014). Although the mechanisms are not yet understood, Ca2+ is believed to be an
important regulator of transport (Li et al. 2014). A plasma membrane aquaporin has been
identified in P. oceanica and is believed to be involved in maintenance of osmotic
balance under hypersaline stress (Serra et al. 2011).
Plants may also adjust cell wall elasticity in an attempt to maintain appropriate
turgor (Martínez et al. 2007), a process tied to physiological adjustments to regulate wall
composition and structure (Wei and Lintilhac 2003). Cell walls expand or contract until
osmotic equilibrium achieved. Adjustments of cell wall elasticity relative to osmotic
stress are not always clear. Spartina alterniflora leaf cell walls became less elastic under
water stressed conditions that may be related to the inability to maintain cellular turgor
(Hessini et al. 2009). However, seagrasses, like H. wrightii, with flexible cell walls, are
more tolerant to short-term fluctuations in salinity (Touchette 2007).
Molecular techniques that identify if stress genes are induced have been proposed
by Macreadie et al. (2014). This approach utilized real-time polymerase chain reaction to
measure the extent “housekeeping” genes responded to heat stress in the seagrass Z.
marina (Ransbotyn and Reusch 2006). Activation of stress response genes to stressors
has been used in terrestrial (Shinozaki and Yamaguchi-Shinozaki 2007) and aquatic
species (Massa et al. 2011) and is a promising avenue of research that can provide early
evidence of acute stress that can eventually lead to prediction of landscape-level
mortality. Transcriptomics can be used to identify candidate genes or groups of genes that
are switched on during a stress event. Identification of these genes can be used in tandem
with controlled laboratory experiments that, for example, assess increased respiration
rates in seagrasses and could be an additional tool to predict the trajectory of seagrass
stress and subsequent change in populations (Macreadie et al. 2014).
These studies demonstrate that the stress experienced by plants induces a
metabolic and physiological response that requires allocation of substantial energy
resources in an attempt to maintain a positive carbon balance (Collier et al. 2010;
Sandoval-Gil et al. 2014). Seagrasses like H. wrightii tend to be more resilient to
perturbations and suboptimal growth conditions than others (Pulich 1980; 1982). Yet they
still have carbon demands and increased respiration rates to compensate for the stress
response. However, there are also documented cases where colonizing seagrasses
(Halophila ovalis, Halodule uninervis, and Zostera muellari) have, under hyposaline
conditions, increased shoot density and maintained normal shoot production prior to
mortality (Collier 2014).
Growth response (leaves, rhizomes, and roots) of seagrasses to experimental
additions of nutrients, typically nitrogen, have been seen numerous times since Orth
(1977) pioneering work on the temperate species Z. marina. Early work proposed that
nitrogen limits growth in terrigenous sediments and phosphorus is limiting in carbonate
sediments (Short 1987; Short et al. 1990). Lee et al. (2007) provided a comprehensive
review across species and geography of seagrass growth as related to light, temperature
and nutrients. Lee and Dunton (2000) is the only study from the LLM to have
documented a positive response to sediment nitrogen additions on seagrass growth in T.
testudinum. Experimental studies that focused on H. wrightii response to addition of
nutrients are fewer (Table 1) with a focus on the eastern Gulf of Mexico (Powell et al.
1989). However, Pulich (1989) demonstrated growth response to nitrogen in H. wrightii
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in laboratory microcosms. Some researchers have examined the correlation of areal
seagrass distribution with freshwater sources to the estuary (Table 1), but few have
involved controlled laboratory experiments. van Katwijk et al. (1999) used a controlled
experiment to explain the distribution and ecology of Z. marina in the Dutch Wadden
Sea. Plants from two sites, a high nutrient-low salinity marine site, and an estuarine site,
were subjected to changes in salinity and nutrient load. At highest salinity, high nutrient
load had no effect on plants, but estuarine plants were adversely affected by high nutrient
load. Moderate salinities (23-26) in both had a positive growth effect.
Studies of nutrient uptake by seagrasses have also been performed dating back a
number of years. Early work demonstrated nitrogen uptake by leaves and roots in Z.
marina (Short and McRoy 1984). Subsequent work has demonstrated the relative
importance of leaves (50% or more) to acquisition of nitrogen (Table 1; Stapel et al.
1996; Lee and Dunton 1999) and phosphorus (Stapel et al. 1996; Gras et al. 2003). In
almost all studies the plants followed Michaelis-Menten kinetics. Uptake kinetics has
utilized the disappearance of substrate (Lee and Dunton 1999), but tracing the movement
of a nitrogen species using tissue incorporation of 15N-labeled ammonium and nitrate has
been implemented with greater frequency (Cornelisen and Thomas 2004; Nayar et al.
2010; Alexandre et al. 2011; La Nafie et al. 2014). Ammonium is more readily taken up
by both roots and leaves than nitrate in seagrasses (Lee and Dunton 2000; Nayar et al.
2010; La Nafie et al. 2014). Water column nitrate and ammonium concentrations in the
LLM are generally low (< 3 µM) (Kowalski et al. 1999). Porewater ammonium
concentrations in the LLM are likewise low across most of the basin (9-70 µM)
(Kowalski et al. 1999). Nitrate is found in the sediment porewater, but its concentration is
negligible and limited to the top few mm (µM) (Lee and Dunton 2000).
Halodule wrightii, like all other seagrasses, is typically distributed at the
convergence of the open ocean (e.g., tides and storms) and land (e.g., watersheds and
associated runoff) (Nixon 1995). Anthropogenic pressures influence each side of the
estuary. Storms (thunderstorms and tropical cyclones) have increased over the past
decade and more, associated with global climate change (Emanuel 2005; Knutson et al.
2010). The freshwater storm discharge which follows causes periodic hyposalinity events
within the estuary. The degree of metabolic stress to sessile estuarine species is often
proportional to the magnitude and duration of the hyposalinity event (Gavin and Durako
2014). Estuarine plants are adapted to the diurnal rhythm of estuarine salinity variation.
During periods of drought, they adjust their physiology and metabolism to cope with
gradual increases in salinity (weeks to years), and possess mechanisms to adjust to
osmotic stresses brought with hypersalinity. With episodes of hyposalinity,
environmental changes can occur relatively quickly (hours to days) and the subsequent
osmotic stress can in the case of submersed plants, cause mortality and substantial loss of
cover (Preen et al. 1995; Campbell and McKenzie 2004; Griffin and Durako 2012; H.
DeYoe, unpublished report to the Texas General Land Office 2013).
Superimposed on atmospheric climate change are the effects of engineered flood
diversion systems and coastal development, the impacts of which are yet to be completely
understood (Sklar and Browder 1998; Scavia et al. 2002). These systems divert
freshwater discharge, often laden with organic and inorganic nutrients, directly into the
estuary. Seagrass community diversity and species abundance are subsequently altered
6
(Montague and Ley 1993; Biber and Irlandi 2006). Whether through direct precipitation
or by channeled floodwater discharge, seagrasses are now more frequently exposed to
episodes of nutrient-rich freshwater discharge, periods of drought, and the associated
metabolic stresses which accompanies exposure, rapid change, and loss (Murphy et al.
2003; Parida and Das 2005; Orth et al. 2006; Gavin and Durako 2012).
Recent efforts to monitor and identify stress factors that point to early indications
of seagrass loss have focused on selected biological and water quality indices (Neckles et
al. 2012), but these indicators shift relatively slowly on timescales of months to years.
Increased frequency of storm events that introduce large pulsed volumes of freshwater
hinders the ability to forecast the extent of seagrass stress and subsequent loss. It is
possible to examine nitrogen metabolism, physiological tolerance thresholds, and
photosynthetic performance of seagrass species as mechanisms for coping with osmotic
stress, and the extent to which they can sequester and translocate nitrogen and survive
under the influence of hyposalinity. Of the seagrass species found in the Laguna Madre,
H. wrightii (shoal grass) has been documented to be the most euryhaline (McMillan and
Moseley 1967) and, consequently, a logical species to use to test the sub-lethal effects of
hyposalinity and nutrient uptake/utilization on metabolism, physiology, and growth.
There is an energetic cost associated for seagrasses maintaining osmotic
equilibrium by active transport of ions in hyposaline surroundings (Touchette 2007). As
estuarine plants attempt to osmoregulate, adjustments in their photosynthetic capacity are
expected to be compromised (Murphy et al. 2003; Koch et al. 2007). Under hyposalinity
stress, respiratory rates increase (Shafer et al. 2011) and there is subsequent accumulation
of reactive oxygen species (Parida and Das 2005; Gavin and Durako 2012), and decreases
in ion and free amino acid concentrations (Parida and Das 2005). Studies on the salinity
tolerance of numerous seagrass species have been made (McMillan and Moseley 1967;
McMahon 1968; Koch et al. 2007; Fernández-Torquemada and Sánchez-Lizaso 2011),
however, all have focused upon the effects on the leaf portion of the plant. There are no
studies which have addressed the effects of hyposalinity on the root/rhizome tissues. It is
likely that seagrass leaves, in the more dynamic water column, respond differently to
changes in salinity compared to rhizomes and roots. Rationale for this supposition is
grounded in limited studies on changing salinity gradients within the interstitial sediment
zone in estuarine and riverine-influenced zones that document a lag period in comparison
to that of the water column (Sanders et al. 1965; Chapman 1981; Pardo et al. 2011).
Rhizome/root response to salinity fluctuations is relevant to understanding
potential resiliency in seagrass communities, yet little is known of their respiratory
responses to environmental stress (Hemminga 1998). For example, the freshwater
discharge from Hurricane Alex in 2010 caused more than one-half of the Lower Laguna
Madre (LLM) to experience salinities between 5 and 10 PSU for more than a month (H.
DeYoe, 2011). This water was directed into the LLM by way of an engineered flood
system, which carries stormwater discharge with each precipitation event. Some H.
wrightii populations closest to the discharge source (Arroyo Colorado) endured zero
salinity for more than three weeks (H. DeYoe, 2013, unpublished report to the Texas
General Land Office). It is not known what stresses H. wrightii endured during this
period of hyposalinity. There was widespread leaf loss and how metabolically adapted
arises is not known. If the leaves were sloughed by the vertical short shoots because of
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extreme stress, or to eliminate a resource burden, but re-emerged once salinities
ameliorated, then the question shifts to how the root-rhizome tissues responded. It is
likely that the rhizome/root portion experienced low salinity stress effects out of phase
and amplitude with that of the leaves.
Purpose, Objectives and Hypotheses
Osmotic stress in plants induces metabolic and physiological responses that can
be measured to assess the magnitude of response, prior to the morphological appearance
of stress. Hypersaline conditions accrue gradually under drought conditions, while the
freshening of estuaries is typically caused by more rapid, pulsed deliveries of stormwater,
either through direct precipitation, runoff, or indirectly by way of alteration and diversion
of natural flow (Montagna et al. 2013). Most studies on seagrass response to salinity
stress have focused on hypersalinity effects (Table 1). While there are few, but increasing
numbers of experimental studies on the metabolic and physiological responses of
seagrasses to hyposalinity, many of these have focused on Halophila johnsonii and H.
decipiens, and R. maritima (Table 1). How does photosynthetic performance in H.
wrightii change as it devotes more resources to homeostasis? How do respiration rates
change as the plant attempts to meet its energy needs? As energy resources are allocated
to meet physiological needs, will this be reflected in changes in intracellular solute
concentrations (amino acids and simple metal ions) and inorganic nitrogen (ammonium
and nitrate)? The purpose of my proposed study is to gain an understanding of how H.
wrightii may cope with low salinities and how it responds and persists during periods of
hyposalinity. This study will measure how salinity stress affects the photosynthesis,
metabolism, physiology and uptake and utilization of nitrogen of H. wrightii.
Laboratory and aquaria experiments are proposed which focus on selected
indicator metabolic and physiological parameters (e.g., photosynthetic efficiency, leaf
elongation rates, photosynthetic and respiratory rates, stress response) measured under
decreasing salinity (20, 10, 5) compared to that of a control (35) with the following
objectives:
1.
2.
3.
4.
5.
6.
To measure leaf production, primary productivity rates, and photosynthetic
efficiency
To measure respiratory rates of all major organs
To assay and compare stress indicators (e.g., lipid peroxidation) in leaves and
rhizome/roots
To measure changes in inorganic ions (e.g., K+, Na+) and free amino acid
concentrations in leaves and rhizome/roots
To determine the degree to which salinity reduction and increased inorganic
nitrogen concentrations interact to produce a synergistic suppressive effect on leaf
growth, photosynthesis and respiration
To compare the extent to which H. wrightii incorporates nitrogen into its leaves
and rhizome/root tissues under hyposalinity stress
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The following hypotheses are identified.
1. To measure leaf growth rates, respiratory and photosynthetic rates of leaf and
rhizome/root organs in H. wrightii at selected salinity levels (35 - control, 20, 10, and, 5)
experiments are proposed to test the following hypothesis.
Ha: Decreases in salinity below an optimum will impair leaf production rates,
photosynthetic rates and efficiency, whereas respiration rates will proportionally
increase.
Ho: There will be no difference in leaf production, photosynthetic or respiration
rates, irrespective of salinity treatment.
2. To measure changes in tissue concentrations (leaves, rhizomes and roots) of principle
simple (metal) ions and free amino acid concentrations in H. wrightii in order to identify
possible mechanisms used by this species to osmoregulate and maintain homeostasis
under varying salinities (35 - control, 20, 10, and 5), as well as to assay lipid peroxidation
to indicate stress, experiments are proposed to test the following hypothesis.
Ha: Decreases in salinity below an optimum will decrease tissue concentrations of
principle ions and free amino acids, and lipid peroxidation will increase under
decreasing salinity, compared to a control.
Ho: There will be no difference in tissue concentrations of principle ions, or free
amino acid concentrations, and no difference in lipid peroxidation irrespective of
salinity treatment.
3. To measure leaf growth, photosynthetic efficiency, respiration rates, and tissue
nitrogen incorporation, under conditions of water column and sediment porewater
nitrogen enrichment and decreased salinity (35 - control, 20, 10, and 5) (compared to
controls), experiments are proposed to test the following hypothesis.
Ha: The interactive effects of decreased salinity and nitrogen enrichment of the
water column or sediment will lead to a proportional decrease in leaf growth,
photosynthetic efficiency, and nitrogen incorporation by leaves and roots, as well
as increased respiration rates in leaves and rhizome/roots in H. wrightii.
Ho: There will be no difference in leaf growth, photosynthetic efficiency,
respiration rates, and nitrogen incorporation by leaves and rhizome/roots in H.
wrightii compared to control plants.
Methods
Study Site and Culture Facilities
Plant collection and sampling
Plants for all experiments will be collected during spring and summer months. as
sods from a previously studied shallow (< 1 m) site in a uniformly dense monotypic H.
wrightii bed, LLM, Texas (Fig. 1). Sods will be 10 - 15 cm thick (deep) to ensure
inclusion of most rhizomes and roots and placed in 38 l (51 x 28 x 30.5 cm, 0.14 m2 area)
9
aquaria. Water column temperature, salinity, pH, and dissolved oxygen will be recorded
with a Hydrolab Quanta multiprobe at the time of collection. Low nutrient, low
chlorophyll seawater will be collected in 20 l carboys for use in the aquaria and to cover
the sods with sufficient seawater to keep plants saturated during transport. Plants will be
installed in aquaria within 8 hours of collection and placed in an environmentally
controlled room at the University of Texas - Pan American, Edinburg, Texas.
Culture facilities
Individual 38 l aquaria with sods, (hereafter, tanks) will be filled with seawater
from the collection site so that the leaves will be in 15-20 cm of water. Plants will be
allowed to acclimate for 10 days before initiation of experiments. Basic water column
physical parameters (temperature, salinity, pH, and dissolved oxygen) will be recorded
every other day to monitor and maintain environmental conditions. Salinity will be
adjusted by adding distilled water to an initial target salinity level (ca. 35). Water
temperature will be kept at 25 to 30 °C (±2 °C), dependent on time of year. The plants
will be exposed to low heat T5 high output fluorescent white lights on a 14:10 light-dark
cycle (ca. 300 µmol photons m-2 s-1) during the 10 day acclimation and experimental
periods. This value is near the saturation irradiance of 315 µmol photons m-2 s-1 estimated
for H. wrightii in Texas waters (Dunton 1996), and greater than 200 to 300 µmol photons
m-2 s-1 estimated for H. wrightii in the Indian River Lagoon, Florida (Rice et al. 1983).
Dunton (1994) cited a minimum of 2 hours of saturation irradiance necessary for plant
maintenance with 3 and 8 hours, respectively, needed to sustain spring and summer
growth.
For all experiments each tank will be treated as an individual replicate (N = 3) per
salinity treatment. Shoal grass shoot density in the LLM can vary between 5000 to 8000
shoots m-2 (Kowalski et al. 2009). This should translate to a minimum shoot density of
several hundred shoots (to more than 800) tank-1, even with some shoot mortality. Water
for each tank will be circulated using bottom lift aquarium pumps, one on each end of
each aquarium, using charcoal filters with a flow rate of more than 200 l hr-1. Three
replicate tanks will be assigned a salinity treatment of 5, 10, or 20, with 35 as the control.
Following the acclimation period, respective replicate salinity treatment tanks will have
their salinities adjusted in series to target levels by removal of seawater and addition of
distilled water. Adjustments to target salinities will be made in proportion to the salinity
of the experimental treatment so that all treatments reach target salinity at the same time.
Target salinities will be held for one day prior to the initiation of experiments. The
salinity adjustment rate will approximately mimic the maximum overall mean salinity
change rate of the water column during monitoring of freshwater inflow to the LLM
during the Hurricane Alex freshet of 2010 (ca. 5 d-1) (DeYoe and Kowalski, unpublished
data). This rate of change will require a greater percent salinity dilution as target salinities
are approached. Water column salinity in each tank will be recorded once each day for
the duration of the salinity adjustments and corresponding experimental period. Plants
will be at target salinity for two days prior to initiation of experiments. Adjustments be
made during the experiment to maintain the target salinity.
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Experiment 1: Measurement of respiration and photosynthetic rates, photosynthetic
efficiency, and determination of simple (metal) ion and free amino acid
concentrations (addresses hypotheses 1 and 2)
This experiment will use environmentally controlled chambers to effect stepped changes
in salinity (35-control, 20, 10, and 5) and then measure leaf growth rates, respiration and
photosynthesis rates, photosynthetic efficiency (PAM fluorometry), and simple (metal)
ion and free amino acid concentrations.
Respiration Measurements
Respiration measurements will be made in the laboratory using a FireSting
(PyroScience, Inc.) optical dissolved oxygen (DO) fixed oxygen minisensor (3 mm
diameter) probe and 4 channel meter connected to a notebook computer. The FireStingO2
uses REDFLASH dyes embedded in the probe which are excited at orange-red
wavelengths. Four probes will be inserted into individual 8 ml reaction chambers through
a rubber grommet fitted in a screw cap. Triplicate chambers will contain plant tissue and
one additional chamber will be used as a blank (control). One 2 cm long intact long shoot
(rhizome), with associated vertical (short) shoot and roots will be collected from each
treatment tank, thoroughly cleaned of sediment (with seawater of the respective
treatment) and placed in a reaction chamber. Temperatures will be maintained to within 2
°C for all runs. Before measurements begin, oxygen levels will be drawn down to ca.
50% saturation by bubbling in N2 gas. Circulation will be achieved by placing the
experimental apparatus on a tilt rotator. DO measurements will be made in the dark and
logged once every five minutes for thirty minutes. Respiration rates will be calculated as
the difference in DO values divided by time increment, minus the rate calculated in the
control blank. Tissue used for incubations will be dried at 80 °C for 48 h and weighed.
Results for incubations will be expressed as µg O2 (consumed) mg dw tissue-1 hr-1. It is
known that O2 will accumulate in aerenchyma tissue in submersed plants (Kemp et al.
1986). However, Herzka and Dunton (1997) used a six minute lag period for lacunal O2
concentations to equilibriate with the surrounding water column in the broad-leaved
seagrass T. testudinum.
Photosynthetic Measurements
Photosynthetic measurements will be made in the laboratory using the FireSting
(PyroScience, Inc.) optical dissolved oxygen (DO) fixed oxygen minisensor (3 mm
diameter) probes as described for the respiration measurements. Photosynthetic rates
along a leaf can vary from tip to sheath (Enriquez et al. 2002) and H. wrightii typically
has ca. 2 leaves per shoot (Kowalski et al. 2009; unpublished data). To account for these
differences, only the youngest fully greened and emerged leaf sections that attain ca. 5
cm length will be cut and used. The youngest leaves on one replicate shoot per replicate
tank per treatment will be collected from each treatment tank and placed in a reaction
chamber. It is acknowledged that cutting the plant may introduce flooding of arenchyma
tissue and possibly act as a confounding artifact. However, since the experiment was
designed to show an effect of treatment, and given the fact that the control will be
exposed to the same potential artifact(s) as the experimental variables, any differences in
results will be attributable to the treatment alone. Oxygen levels will be drawn down to
11
less than 50% saturation by bubbling in N2 gas. Circulation will be achieved by placing
the experimental apparatus on a tilt rotator. DO measurements will be made under
saturating irradiance (ca. 300 µmol m-2 s-1) provided by two 100 W cool, white light LED
bulbs positioned on opposite sides of the chambers with measurements logged once every
five minutes for 30 minutes. Photosynthetic rates will be calculated as the difference in
DO values divided by time increment, minus the rate calculated in the control blank.
Tissue used for incubations will be dried at 80 °C for 48 h and weighed. Results for net
photosynthetic rates will be determined by subtracting respiration rates from gross
photosynthetic rates divided by incubation time and chlorophyll concentrations and
expressed as µmol O2 mg-1 Chl a hr-1.
Blade Chlorophyll Concentrations
Fresh leaves (ca. 15 mg) from each treatment tank (N = 3) will be collected
weekly, finely minced, and extracted in the dark for 48 to 72 hours at 25 ºC in 5 ml of N,
N-dimethylformamide (DMF) solvent for spectrophotometric analysis of chlorophylls
(Chl) a and b (Porra et al. 1989). Samples will be analyzed on a Shimadzu 160 UV
spectrophotometer at 664 nm and 647 nm. Absorbance values will be read at 750 nm to
correct for turbidity. Tissue will be dried at 80 °C for 48 h and results expressed as mg
Chl a and b g-1 dw leaf tissue, and Chl a:b ratio.
Chlorophyll fluorescence measurements of photosynthetic efficiency
Maximum quantum efficiencies and rapid light curves (RLC) will be obtained
from plants in each replicate treatment using a portable pulse amplitude modulated
fluorometer (Junior-PAM, Walz, Germany). RLCs represent the relationship between
electron transport rate and irradiance. The youngest fully emerged 5 cm minimum length
leaves will be used to minimize differences in leaf age and photosynthetic performance.
Leaves will be gently scraped to remove epiphytes. Chlorophyll fluorescence
measurements will follow the protocol outlined in Gavin and Durako (2012). A dark leaf
clip will be used to hold the PAM fiber optic probe 3 mm from the surface of the leaf. All
measurements will be made on the same relative position on the leaf. Maximum quantum
efficiency measurements will be made before dawn (0530 - 0600 hrs) on measurement
days. Initial fluorescence (F0) will be measured first and the leaf then subjected to a pulse
of saturating light during which a second (maximum) fluorescence measurement (Fm)
will be taken. Maximum (dark-acclimated) quantum efficiencies estimate maximum
photochemical efficiency of photosystem II, summarized in the equation, (Fm - F0)/Fm =
Fv/Fm. RLC measurements will be made between 1000 and 1100 hours and using nine
light levels (0, 20, 49, 82, 136, 194, 298, 418, and 654 µE m-2 s-1, each at 10 s intervals.
Relative electron transport rates (rETR) will be estimated using the following equation,
rETR = (Fm' - Fs)/Fm' x PPFD x 0.5, where Fm' = light acclimated maximum fluorescence
(30 min), Fs = steady-state fluorescence yield in the light adapted state, PPFD = intensity
of photosynthetically available radiation (PAR) at the corresponding RLC irradiance step,
and 0.5 assumes one-half of the available PAR is absorbed by photosystem II. Mean
values of photosynthetic efficiency at subsaturating PAR (α), irradiance at onset of
saturation (Ek), and maximum relative electron transport rate (rETRmax) will be calculated
for each treatment from RLCs using a double exponential decay function described by
12
Ralph and Gademann (2005), but with the incident irradiance (Ii) replaced by irradiance
per quanta of light (photons) absorbed (Ia) by the photosynthetic pigments (Saroussi and
Beer 2007).
Leaf elongation rates
The leaf-clipping method of Kowalski et al. (2001) will be used to measure leaf
elongation (shoot production) because it is an integrator of production. Halodule wrightii
shoots in a haphazardly selected 5 cm diameter area from each replicate treatment will be
clipped 1-2 cm above the sediment surface. Leaves of clipped shoots will be allowed to
re-grow for 2 weeks and shoots re-clipped. Leaves from each clipped shoot will be
collected, gently scraped of epiphytes, measured for leaf length to the nearest mm, and
dried at 80 °C to a constant weight. Lengths of all leaves per shoot will be pooled to
calculate mean leaf elongation rates of re-grown leaves expressed as mm day-1, and shoot
production rates (mg shoot-1 day-1).
Lipid Peroxidation
Estimation of lipid oxidation will follow the method described in Hodges et al.
(1999) for the thiobarbituric acid-reactive substances (TBARS) assay where
malondialdehyde (MDA) is a secondary end product of the oxidation of polyunsaturated
fatty acids and is reacted with thiobarbituric acid (TBA). The TBARS assay is an index
of lipid peroxidation. Approximately 100 mg of tissue is homogenized by adding 0.5 ml
0.1 % (w/v) TCA. The homogenate is centrifuged for 10 min (15000 x g, 4.0 °C). The
supernatant is collected and 0.5 ml is mixed with 1.5 ml 0.5% ΤΒΑ diluted in 20% TCA
which is then incubated in a waterbath at 95 °C for 25 min and then incubated on ice.
Absorbance is measured at 532 and 600 nm.
Major Ion Concentrations and Free Amino Acid Analysis
Metal concentrations (Mg, Na, Ca, and K) will be measured from 500 mg (dry
weight) leaf and rhizome-root tissue taken from each of the replicate salinity treatment
tanks. Samples will be assayed by flame emission using an inductively coupled plasma
optical emission spectrometry (ICP-OES) (Khan et al. 1999) in the laboratory of Jason
Parsons, the University of Texas - Pan American. Leaf and rhizome-root tissue from each
experimental tank will be dried to a constant weight for at least 48 hr at 80 °C. Results
will be expressed as mg [ion] g-1 dw.
Free amino acids (FAA) will be extracted from leaf and rhizome-root tissue
(separately) salinity treatments using the methods outlined in Hacham et al. (2002).
Briefly, 150 mg of fresh tissue will be collected from each replicate tank and ground in
liquid nitrogen using a mortar and pestle in the presence of 600 µl of
water:chloroform:methanol (3:5:12 v/v) and then transferred to a 1.5 ml tube and
centrifuged at 5000 g for 2 minutes. The supernatant will be removed and saved and the
residue re-extracted with another 600 µl of water:chloroform:methanol extraction buffer
and centrifuged for 2 minutes. This supernatant will be combined and 300 µl of
chloroform and 450 µl of water added to the combined supernatants and centrifuged for 2
minutes. The upper water:methanol phase will be collected and transferred to a fresh tube
and placed in a speed vac to dry for about 3 hours. The pellet will be kept at -20 °C until
13
analyzed using fluorescence high performance liquid chromatography (HPLC) from a
modified protocol of Liu et al. (2013) in his laboratory (The University of Texas at
Austin, Marine Science Institute. FAAs will be separated on a 25 cm x 5 µm Alltech
Alltima C18 column with a flow rate of 1 ml min-1. A binary gradient of 0.05 M NaOAc
(pH 5.7) and 5% tetrahydrofuran (eluant A) and MeOH (eluant B) will be used, ramping
from 20% B to 50% B in 40 minutes, then to 100% B in 20 minutes. FAAs will be
detected by fluorescence and identified by retention time comparison with an authentic
standard mixture (Pierce) and γ-aminobutyric acid (GABA, Sigma) manually added to
the mixture. Concentration of FAAs on triplicate samples generally agree within 12 (±
8)%. Results will be expressed as µmol mg-1 wet wt tissue.
Statistical Design
Statistical analyses will be performed using a general linear model procedure
(Systat Software, Inc. 2013). Values will be reported as means ± standard error. Data will
be evaluated to ensure compliance with the assumptions of parametric statistics (Zar
1984). In the event that assumptions of normality and equal variance are violated, data
will be transformed and re-analyzed. Differences in photosynthetic rates, leaf elongation
rates, and Chl concentrations among salinity treatments (5, 10, 25, and 35) will be tested
with One-Way Analysis of Variance (ANOVA) where salinity is the two main effect.
Differences in respiration rates of leaves and rhizome/roots tissues will be tested with
Two-Way Analysis of Variance (ANOVA) where salinity treatments (5, 10, 20, and 35)
and tissue type are the two main effects. Differences in pulse amplitude-modulated
(PAM) fluorescence will be tested with Two-Way Analysis of Variance (ANOVA) where
salinity treatments (5, 10, 20, and 35) and sample date are the two main effects.
Differences in ion and free amino acid concentrations, and lipid peroxidation will be
tested by Three-Way Analysis of Variance (ANOVA) where salinity treatments (5, 10,
20, and 35), tissue type, and sampling dates are the two main effects. Where significant
differences (α = 0.05) for a main effect are detected, the means will be examined using
the post-hoc Holm-Sidak method multiple-comparison test to determine where
statistically significant differences among means occurred (Systat Software, Inc. 2013).
Experiment 2: Synergistic effects of water column nitrogen and hyposalinity in leaves of
Halodule wrightii under laboratory conditions.
This experiment will subject Halodule wrightii to the interactive effects of hyposalinity
(35-control, 20, 10, and 5) and increased water column inorganic nitrogen concentrations
(+”N”). Leaf growth rates, respiration and photosynthetic rates, and photosynthetic
efficiency (PAM fluorometry) will be measured and compared to controls obtained in the
absence of nitrogen amendments. (Addresses hypothesis 3)
This experiment is related to Experiment 1 in that plants under hyposalinity stress
with elevated water column nitrogen are not expected to demonstrate growth and
photosynthetic stimulation as a result of energy expenditure to maintain osmotic
homeostasis. Control conditions of ambient water column nitrogen concentrations at a
salinity of 35 will stimulate normal rates of leaf growth and photosynthesis. Respiration
14
is expected to be less than those of plants under hyposalinity conditions, regardless of
nitrogen addition.
Culture set-up and experimental conditions
Halodule wrightii will be cultured in 38 l (51 x 28 x 30.5 cm, 0.14 m2 area)
aquaria as in experiment 1 with the same preparations and method and timing to bring
tanks to the experimental salinity levels. An initial loading of 2 g of ammonium chloride
will be added to +”N” tanks for an initial ammonium load of 12 g N m-2 and a
concentration of ca. 125 µM. Conditional controls will include stepped reductions in
salinity, as in experiment 1, but without supplement of nitrogen. Absolute controls will
have no nitrogen supplement with salinity at 35. Concentrations of the ammonium
chloride placed in respective experimental treatment will be adjusted to initial target
salinities. Tissue nitrogen concentrations will be assessed at the beginning and end of the
experimental period. Water column samples for nitrite+nitrate and ammonium will be
assayed the first three days of the experiment to ascertain that porewater nitrogen is not
leaking sufficiently quickly into the water column so as to provide uncontrolled elevated
nitrogen concentrations. Tanks will be filled with seawater from the collection site so that
the leaves will be in 15-20 cm of water. Plants will be allowed to acclimate for one week
before initiation of experiments (as with other experiments) at ambient water column
nitrogen concentrations (ca. 2 µM). Nitrogen will be added the day of final salinity
adjustments and experiments begin. Inorganic nitrogen (nitrite+nitrate and ammonium)
concentrations will be assayed using standard colorometric techniques (Parsons et al.
1984).
Plant Growth Measurements - Leaf growth
Same as in experiment 1 above
Photosynthetic Measurements
Same as in experiment 1 above
Respiration Measurements
Same as in experiment 1 above
Blade Chlorophyll Concentrations
Same as in experiment 1 above
Changes in Nitrogen Tissue Concentrations
Leaf tissue will be assayed at the beginning and end of an experimental period.
Leaf tissue will be finely ground, dried and analyzed in a Carlo-Erba 2500 NC elemental
analyzer for nitrogen and carbon concentrations.
Statistical Design
Analysis of results will be performed using SigmaPlot 12.5 (Systat Software, Inc.
2013). Differences in leaf elongation rates will be tested using Two-Way Analysis of
Variance (ANOVA) where salinity treatments (5, 10, 25, and 35) and nitrogen
15
concentrations are the two main effects. Differences in photosynthetic efficiency, blade
chlorophyll concentrations, and leaf nitrogen concentrations will be tested using ThreeWay Analysis of Variance (ANOVA) where salinity treatments (5, 10, 25, and 35),
nitrogen concentrations, and sample date are the main effects. Where significant
differences (α = 0.05) for a main effect are detected, the means will be examined using
the post-hoc Holm-Sidak method multiple-comparison test to determine where
statistically significant differences among means occurred (Systat Software, Inc. 2013).
Experiment 3: Impact of nitrogen enriched sediments on Halodule wrightii leaf growth
under conditions of hyposalinity. (Addresses hypothesis 3)
This experiment is related to Experiment 2 in that plants under hyposalinity stress will be
expected to decrease nitrogen incorporation (ammonium and nitrate) in order to maintain
osmotic homeostasis. Under control (field) conditions when the plant is normally limited
by nitrogen, inorganic nitrogen (ammonium and nitrate) will be taken up and assimilated
into accumulating leaf growth and biomass (assuming no light or carbon limitation). This
experiment will test the effects of stepped reductions in salinity (35-control, 20, 10, and
5) on leaf growth and biomass allocation in H. wrightii under laboratory conditions to test
the interactive effects of nutrient amendments to the sediment under decreasing salinity.
It is expected that experimental treatments will not stimulate an increased growth and
biomass to the same extent as that of the control.
Culture set-up and conditions
Sediment nitrogen stimulation in H. wrightii under hyposaline water column and
sediment porewater conditions will be assessed by amending sediments with ammonium.
Because this study involves manipulation of sediment inorganic nitrogen concentrations a
sediment "sprinkler" system will be used (Pulich 1982). This system allows for direct
infusion of nutrient treatments without disturbing the sediment or water column. Each
sprinkler will consist of four horizontal PVC pipes, each drilled with paired 2 mm holes,
each pair separated every 6 mm. The ends of each vertical pipe will be closed by a cap.
Horizontal pipes will be attached to a vertical standpipe that will rise above the water
surface for routine sampling and addition to the sediment of a total of 35 g of ammonium
chloride crystals per tank. This loading will yield an initial ammonium load of 50 g N m-2
and a concentration of ca. 500 µM. Concentrations of the ammonium chloride placed in
respective experimental treatment solutions will be adjusted to initial target osmolalities.
Two sprinklers and standpipes each will be placed in individual 38 l tanks, covered by a
bottom layer of washed aquarium gravel overtopped with sods of plants collected from
the donor site. Tanks will be filled with seawater from the collection site so that the
leaves will be in 15-20 cm of water. Plants will be allowed to acclimate for one week
before initiation of experiments (as with other experiments) at ambient water column
nitrogen concentrations (ca. 2 µM). Leaf and rhizome/root issue nitrogen concentrations
will be assessed at the beginning and end of the experimental period. Water column
samples for nitrite+nitrate and ammonium will be assayed the first three days of the
experiment to ascertain that porewater nitrogen concentrations are not leaking sufficiently
16
quickly into the water column so as to provide uncontrolled elevated nitrogen
concentrations.
Plant Growth Measurements - Leaf elongation rates
Same as in experiment 1 above
Photosynthetic Measurements
Same as in experiment 1 above
Blade Chlorophyll Concentrations
Same as in experiment 1 above
Statistical Design
Statistical analyses will be performed using a general linear model procedure
(Systat Software, Inc. 2013). Values will be reported as means ± standard error. Data will
be evaluated to ensure compliance with the assumptions of parametric statistics (Zar
1984). In the event that assumptions of normality and equal variance are violated, data
will be transformed and re-analyzed. Differences in respiration and photosynthetic rates,
pulse amplitude-modulated (PAM) fluorescence, Chl concentrations, and leaf elongation
rates will be tested by Two-Way Analysis of Variance (ANOVA) where salinity
treatments (5, 10, 25, and 35) and sampling dates are the two main effects. Where
significant differences (α = 0.05) for a main effect are detected, the means will be
examined using the post-hoc Holm-Sidak method multiple-comparison test to determine
where statistically significant differences among means occurred (Systat Software, Inc.
2013).
17
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28
Figure 1. Map showing the location of the proposed donor site for Halodule wrightii and
locations of tidal passes and the Arroyo Colorado, the principal source of fresh water to
the Lower Laguna Madre, Texas.
29
Table 1. Studies on the influence of changes in salinity, water column enrichment effects,
nutrient uptake studies, tissue nutrient status (ratios), and sediment enrichment
experiments and the effects on selected plant performance parameters for various
seagrass species.
Species
Location
Independent
Dependent
Results
Source
(Experimental Variable(s)
Variable(s)
Setting)
unvegetated
Cape Cod,
Massachusetts
(USA)
(Field)
Sediment
salinity regime
relative to
position along
the Pocasset
River estuary
Top 2-3 cm
Salinity levels
(28-31) and 61
Salinity by
season and depth
Bottom interstitial
salinities more
stable at the lower
end of the river
(nearest Buzzard's
Bay)
Sanders et al.
(1965)
Thalassia
testudinum,
Syringodium
filiforme,
Halodule
wrightii,
Ruppia
maritima, and
Halophila
ovalis
Halodule
wrightii and
Syringodium
filiforme
Texas (USA)
(Microcosms)
Leaf growth
(height)
Chlorophyll a
Hierarchical
sequence of
salinity tolerance:
Halodule most
tolerant (over
Ruppia) and
Syringodium least
tolerant of all
McMillan and
Moseley (1967)
Texas (USA)
(Microcosms)
Salinity levels
(0-87.5) in fairly
equal increments
Leaf length and
qualitative
appearance
(color/condition)
McMahon
(1968)
unvegetated
British Columbia
(Canada)
(Field)
Salinity by
season and depth
Halophila
ovalis
Cape Peron,
(Australia)
(Cultures and
microcosms)
Sediment
salinity regime
relative to
position along
the Fraser River
estuary
(oligohaline to
polyhaline)
Top 6 cm
Marine and
estuarine
ecotypes
subjected to
salinity
treatments of 10,
25, and 35
Halodule most
euryhaline with
"growth" <3.5 psu
to ca. 70 psu
Syringodium
survived best at
salinity of 35 to
44
Vertical salinity
gradients present
(especially the
mesohaline zone)
and influenced
seasonally
High salinityadapted plants
intolerant of
hyposalinity (10
psu)
Estuarine plants
healthy at 20 and
10 psu, but
appeared stressed
at 4 weeks at 10
Benjamin et al.
(1999)
Ultrastructure
(qualitative),
Rhizome
branching, leaf
length, leaf
surface area
Chapman
(1981)
30
Thalassia
testudinum,
Syringodium
filiforme, and
Halodule
wrightii
Florida (USA)
(Field and
microcosms)
Field
observations of
distribution and
abundance and
associations of
seagrasses to
salinity regimes
Leaf growth and
biomass
Salinity
exposure
experiments (15
days / salinities
of 5 to 45
Ruppia
maritima
Florida Bay
(USA)
(Cultures)
0, 10, 20, 40
salinity
treatments
Total and nonvacuolar
osmolality
Soluble and total
carbohydrates
Photosynthetic
capacity (PAM)
Leaf senescence
accelerated, cell
surface area, leaf
and internode
length and
rhizome diameter
decreased with
prolonged
exposure
Chloroplasts
number decreased
All species found
in study area, but
Halodule
associated with
canal discharges.
Species-specific
susceptibility to
treatments
Thalassia grew
best at salinity of
30 to 40 Lowest
growth at salinity
extremes (5 and
45)
Syringodium grew
best at 25 and
growth dropped
above and below
25
Halodule was
most tolerant of
salinity extremes
and grew well at
all treatments
Osmolality
decreased at 0 psu
and increased at
40 psu
Total
carbohydrates
decreased 65%,
Soluble increased
by 34%, proline
levels increased at
salinity of 10 and
decreased at 0
Soluble
carbohydrates and
proline act as
osmolytes
Photosynthetic
Lirman and
Cropper (2003)
Murphy et al.
(2003)
31
Thalassia
testudinum,
Halodule
wrightii, and
Ruppia
maritima
Florida Bay
(USA)
Microcosms
(chambers) within
mesocosms
Two salinity
treatments pulsed high and
gradual high
(each at 8
incremental
levels)
Shoot decline
Net
photosynthesis
Photosynthetic
performance
(PAM)
Leaf growth for
Thalassia and
new shoot growth
(rhizometagging) for other
two species
Tissue osmolality
Zostera marina
Seden Strand,
Odense Fjord
(Denmark)
(Microcosms)
Temperature
treatments of 5,
10, 15, 20, 25,
27.5 and salinity
treatments of
2.5, 10, 15, 20,
25, 30, and 35
Shoot mortality,
shoot size (leaf
number and
biomass), leaf
growth and
photosynthetic
capacity
Cymodocea
nodosa and
Zostera noltii
Almadraba, Spain
(Microcosms)
Salinity levels (2
to 72) and pulsed
Leaf growth and
survival
Zostera
japonica
Washington and
Oregon, USA
(Laboratory)
Salinity (5, 20,
and 35)
Photosynthesis
and respiration
(O2 method)
efficiency lowest
at 0 and 40 psu
(best at 10 and 20
psu)
All species
tolerant of high
salinities at slow
rate of change. All
tolerated salinities
to 45, but above
this stress
responses
increased (C drain
associated with
osmoregulation
(synthesis of
solutes),
decreased
photosynthetic
performance,
increased
respiration)
Low salinity
treatments cause
increased
mortality in plants
along with
negative effects
on leaf elongation
and shoot
morphology
Best salinity range
10-25
Plants tolerated
salinity change
better when
salinity change
was gradual
Zostera noltii
tolerated
hyposalinty better
Pmax at 20 psu
(Oregon), but no
effect for
Washington
Salinity affects
respiration rates in
both, but no
differences
between
populations
Koch et al.
(2007)
Nejrup and
Pedersen
(2008)
FernándezTorquemada
and SánchezLizaso (2011)
Shafer et al.
(2011)
32
Zostera marina,
Halodule
wrightii, and
Ruppia martima
North Carolina
(USA)
(Mesocosms)
NO3- water
column
enrichment
Leaf
elongation
(Zostera) and
lateral short
shoot
production
(Zostera and
Ruppia)
Thalassia
testudinum
Texas (USA)
(Laboratory)
Nitrogen (NO3-,
NH4+)
concentrations
(leaves and roots)
seasonally
(temperature)
Leaf and root
NO3- + NH4+
uptake rates
(seasonal)
Zostera marina
Netherlands
(Laboratory
microcosms)
Nutrients (NO3-,
NH4+ and PO3-)
concentrations
and salinity (23,
26, 30)
Leaf metrics,
Chl a, tissue N
and P
Halodule
wrightii,
Syringodium
filiforme and
Thalassia
testudinum
Halodule wrightii
and Thalassia
testudinum
Florida (USA)
(Field)
In situ water
column
enrichment of N
and P (Osmocote)
14C
Florida (USA)
(Field)
Current velocity
(field flume)
Leaf NH4+
uptake rates
Syringodium
filiforme and
Thalassia
testudinum
Florida Bay
(USA)
(Field)
In situ sediment
fertilization (+N,
+P, and +N+P)
Nearshore and
offshore
Leaf length,
abovesediment
biomass, and
Thalassia leaf
growth
uptake by
leaves and
epiphytes
Change in leaf
biomass
Zostera growth
negatively
affected at
high NO3concentrations,
but growth in
Halodule
slightly and
Ruppia highly
stimulated
Leaves and
roots provide
ca. 50% each
total N
requirements,
but rates vary
seasonally
Higher salinity
adversely
affected plant
responses
Higher
salinityadapted plants
not affected by
high nutrients
All plants at
low salinities
stimulated by
higher
nutrients
Leaf
production
increased in
enriched plots,
but not leaf
biomass
Leaf uptake
rates
influenced by
water velocity
as affected by
leaf and
canopy
morphology
N limited in
offshore
carbonate
sediments
N+P in
nearshore
Burkholder et
al. (1994)
Lee and
Dunton
(1999)
van Katwijk
et al. (1999)
Wear et al.
(1999)
Thomas et al.
(2000)
Ferdie and
Fourqurean
(2004)
33
Halodule wrightii
Florida (USA)
(Field)
In situ water
column
enrichment of N
and P (Osmocote)
Halodule wrightii
and Thalassia
testudinum
Florida and
Alabama (USA)
(Field)
Examined spatial
trends of C, N,
and P in
seagrasses and
their epiphytes
Fish
abundance and
composition
with changes
in leaf and
epiphyte
biomass, leaf
growth and
Chl a
Spatial and
seasonal
differences
apparent in
nutrient ratios
Zostera marina
Málaga, Spain
Protein pumpmediated
electrochemical
gradients
Leaf and root
uptake of NH4+
and cellular
transport
Syringodium
filiforme
San Salvador,
Bahamas
(Field)
+N, +P, +N+P
enrichment to
carbonate
sediments
Halodule wrightii
and Thalassia
testudinum
Florida Bay
(USA)
(Field)
N and P nutrient
enrichment from
bird roosts
Biomass,
tissue nutrient
concentrations,
whole plant
growth,
nitrogen
fixation
Leaf biomass
and short shoot
density
Enhalus
acoroides,
Thalassia
hemprichii and
Cymodocea
rotundata
Cape Bolinao,
The Philippines
(Field)
+N+P enrichment
Biomass,
tissue nutrient
concentrations,
tissue Chl a,
photosynthetic
rates (O2),
shoot density
Decrease in
Halodule
biomass in
treated plots,
attributed to
increased
grazing on Nrich leaves
Heck et al.
(2006)
P limitation in
epiphytes and
seagrasses in
Alabama
(terrigenous
sediments)
High affinity
NH4+ transport
not dependent
on Na+
concentrations
Syringodium
growth was Plimited on
carbonate
sediments, and
increased Nfixation
Halodule
wrightii
has a higher
nutrient
demand than
Thalassia
testudinum and
can replace T.
testudinum
during
secondary
succession
under high
nutrient
concentrations
The nature and
extent of
nutrient
limitation
varied between
sites and
among species
Thalassia
hemprichii was
mainly P
Johnson et al.
(2006)
Rubio et al.
(2007)
Short et al.
(1990)
Fourqurean et
al. (1995)
Agawin et al.
(1996)
34
Posidonia
oceanica
Spain
(Field)
+N+P enrichment
Biomass,
tissue nutrient
concentrations,
Leaf growth
Halodule
uninervis and
Syringodium
isoetifolium
Green Island
(Australia)
(Field)
+N, +P, +N+P
enrichment to
carbonate
sediments
Biomass,
tissues nutrient
concentrations,
stable N
isotope, amino
acid
concentration
Thalassia
testudinum
Texas (USA)
(Field)
Nitrogen (NH4+)
enrichment of
terrigenous
sediments at two
separated sites
Leaf growth
rates of
Thalassia
Leaf
morphology,
Shoot density
Above- and
below
sediment
biomass (and
ratio)
deficient and
Enhalus
acoroides to
be mainly N
deficient
Spatial and
seasonal
variations in
nutrient
limitation
Increases in
the growth
rate, amino
acid
composition
and tissue
nutrient
content of both
species in
response to
elevated
sediment N,
but not P
Concentrations
of the N-rich
amino acids
asparagine and
glutainine
increased 3- to
100-fold in
seagrass
leaves from N
treatments
Stable N
isotope values
of leaves
decreased in
response to
additions of
nitrogen
High nutrient
site (sediment
and water
column) little
stimulation Oligotrophic
site showed
strong positive
response in
dependent
variables
Alcoverro et
al. (1997)
Udy (1999)
Lee and
Dunton
(2000)
35
Thalassia
testudinum and
Syringodium
filiforme
Florida Keys
(USA)
(Field)
+N, +P, +N+P
enrichment to
sediment
Halodule wrightii
Florida Keys
(Field)
N and P nutrient
enrichment
Leaf growth
rates of
Thalassia
Leaf length,
above-ground
biomass,
macroalgal,
epiphyte, and
sediment
microalgae
abundance
Offshore
communities
increase in
dependent
variables to
+N
Nearshore
communities
increase in
dependent
variables to
+N+P
Nutrient
addition did
not strongly
affect food
web structure
at a eutrophic
site.
Enrichment at
a nutrient-poor
site increased
the abundances
of crustacean
epiphyte
grazers, and
the diets of
these grazers
became more
varied.
Nutrient
addition
increased
grazing on
Halodule
wrightii
Ferdie and
Fourqurean
(2004)
Armitage and
Fourqurean
(2009)
36
Appendix 1. Proposed budget for dissertation research.
______________________________________________________________________
Budget Item
Cost ($)
______________________________________________________________________
Equipment
12 38 liter aquaria with 12 airlift pumps
4 Sun Blaze T5 HO fluorescent 48 lamps
4' long x 8 Lamp at $259.00 each
8 carboys (20 l) at $122.58 each
Expendables
200.00
1036.00
900.64
1 Multi-channel pipetter
1808.00
1 Vapor Pressure Osmometer
5000.00
TBARS assay kit (96 wells) 2 kits at
$185.00 each (Cayman Chemical Co.)
370.00
Pipette tips (1 case) (1000-5000 µl) (1000/box)
73.77
Pipette tips (1 case) (101-1000 µl) (500/box)
90.70
Glass threaded vials (1 package at $81.36 each)
71.32
1 bottle N-N’ Dimethylformamide (1 liter)
(Fisher Scientific)
175.30
Ion analysis (60 samples at $16/sample)
960.00
Free amino acid analysis (72 samples at $42/sample) 3024.00
Travel
Stable isotope analysis (C and N)
(72 samples at $12/sample including
operator time at $78.00)
942.00
2 trips to South Padre Island
(170 miles per trip @ $0.55/mile)
187.00
Boat
2 days (2 trips) at $130/day
260.00
_______________________________________________________________________
Total ($)
14098.73
37
Appendix 2. Timeline for implementation of the proposed research.
Date
Activity
July 2013
Completion of coursework
January 2015
Completion of Formal Comprehensive/Qualifying Examinations
c
Experimental Work
Late May 2015
Collection of plant material from the Lower Laguna Madre for
experiments 1 and 2
Late May 2015
Set-up culture tanks and seagrass incubation (one week
acclimation)
Early June 2015
Begin experiment one (three week duration) (data collection)
Late June 2015
Begin experiment two (one week duration) (data collection)
Late June 2015
Collection of plant material from the Lower Laguna Madre for
experiment 3
Late June 2015
Set-up culture tanks and seagrass incubation (one week
acclimation)
Early July 2015
Begin experiment three (three week duration) (data collection)
August 2015
Data analysis (all experiments)
January 2016
Writing of dissertation
December 2016
Submission of dissertation to committee
May 2016
Graduation
38