University of Florida Thesis Or Dissertation

GROWTH OF SOUTHERN NAIAD (Najas guadalupensis) IN DIFFERENT SUBSTRATES AND FERTILIZATION LEVELS

HEATHER HASANDRAS

A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2016

To my mother Ivy, whose life ended during my research. May I carry on, creating things in this world that will continue to live on past our years.

ACKNOWLEDGMENTS

I wish to thank my parents George and Ivy for raising a child that values education and encouraging me in my academic and personal pursuits. I thank my children, husband Denny and uncle John for attending my conferences and supporting me through this process of academic exploration. I thank my advisor Dr. Kimberly

Moore for being an academic pillar during my years at University of Florida and for being that force that pushed me to get things completed. I thank my committee members Dr. Lyn Gettys for promoting the value of aquatic research and Dr. Vendrame for having oversight. I thank my lab group (Luci, Mun Wye, Tina, Nancy, Samar,

Joanne, Sandy) and Andy during my studies for sharing friendship, good times, and guidance. You have illustrated that surrounding yourself with the right network of people does indeed help in life.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 6

LIST OF FIGURES ...... 7

LIST OF ABBREVIATIONS ...... 8

ABSTRACT ...... 9

CHAPTER

1 INTRODUCTION ...... 11

2 LITERATURE REVIEW ...... 13

Aquatic Plant Growth ...... 14 Invasive Aquatic Plant Control ...... 17 Invasive Aquatic Plants ...... 20 Southern Naiad ...... 26

3 GROWTH OF NAIAD (Najas guadalupensis) AND HYDRILLA (Hydrilla verticillata) USING CONTROLLED RELEASE FERTILIZER

Materials and Methods...... 29 Results and Discussion...... 30

4 GROWTH OF NAIAD IN SUBSTRATES WITH VARYING PERCENTAGES OF SAND AND CONTROLLED-RELEASE FERTILIZER ...... 36

Materials and Methods...... 37 Results and Discussion...... 39

5 CONCLUSIONS ...... 48

LIST OF REFERENCE ...... 50

BIOGRAPHICAL SKETCH ...... 55

LIST OF TABLES

Table page

3-1 Southern naiad (Najas guadalupensis) and hydrilla (Hydrilla verticillata) shoot and root dry weight of plants grown in sand and fertilized with 0, 1, 2, or 4 g of Osmocote 15N-4.05P- 9.96K...... 33

4-1 Naiad (Najas guadalupensis) shoot and root dry weight of plants grown in 100:0 sand:peat, 75:25 sand:peat, 50:50 sand:peat, 25:75 sand:peat or 0:100 sand:peat (by volume)...... 42

4-2 Naiad (Najas guadalupensis) shoot and root dry weight of plants fertilized with 0, 1, 2, or 4 g of Osmocote 15N-4.05P- 9.96K...... 43

4-3 Shoot concentrations of ammonia (NH3-N), phosphate (PO3-P) and potassium (K) in naiad (Najas guadalupensis) shoots harvested from the experiment 2 conducted in spring 2015...... 44

4-4 Water sample pH, electrical conductivity (EC), nitrate (NO3-N), phosphate (PO3-P) and potassium (K) of naiad (Najas guadalupensis) plants from the experiment 2 conducted fertilized with 0, 1, 2, or 4 g per pot of Osmocote 15N-4.05P- 9.96K...... 45

LIST OF FIGURES Figure page

3-1 This is an example of the 18 gallon plastic containers (mesocosms) and experimental layout. Each mesocosm contained three pots of either hydrilla or naiad...... 34

3-2 During experiment 1, a parapoynx moth (Parapoynx diminutalis) was found at week 7 of the experiment...... 35

4-1 Naiad (Najas guadalupensis) shoot dry weight of plants fertilized with 0, 1, 2, or 4 g of Osmocote 15N-4.05P- 9.96K and grown in 100:0, 75:25, 50:50, 25:75, or 0:100 sand:peat substrate (by volume) ...... 46

4-2 Final substrate electrical conductivity for plants fertilized with 0, 1, 2, or 4 g of Osmocote 15N-4.05P- 9.96K and grown in 100:0, 75:25, 50:50, 25:75,or 0:100 sand:peat substrate (by volume)...... 47

LIST OF ABBREVIATIONS

C Celsius

CRF Controlled Release Fertilizer

EC Electrical conductivity

F Farenheit g Grams g/kg Grams per kilogram

K Potassium mg/L Milligrams per Liter

N Nitrogen nm Nanometer

NH4-N Ammonia

NO3-N Nitrate

P Phosphorus

PO3-P Phosphate

RH Relative humidity

µg/L Micrograms per Liter

W·m2 Watts per meter square

Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for The Degree of Master of Science

GROWTH OF SOUTHERN NAIAD (Najas guadalupensis) IN DIFFERENT SUBSTRATES AND FERTILIZATION LEVELS

Heather Hasandras

May 2016

Chair: Kimberly Moore Major: Horticultural Sciences

The native aquatic plant southern naiad (Najas guadalupensis) is often confused with the invasive weed hydrilla (Hydrilla verticillata). A lot of information has been published on the growth of hydrilla but few studies have investigated the growth of naiad. In experiment 1, we compared the growth of naiad and hydrilla plants fertilized with 0, 1, 2, or 4 grams controlled release fertilizer (CRF) (Osmocote 15N-4.05P-9.96K) layered under the surface per kilogram of the sand prior to planting. We had three mesocosms per fertilizer rate with 3 containers of naiad and 3 containers of hydrilla in each mesocosm. Mesocosms were randomly placed in a greenhouse and filled with water to create a submersed growing environment. The experiment ran for eight weeks in spring 2014 and was repeated in summer 2014. Data from summer 2014 showed that naiad shoot dry weight was greater with 2 g CRF/kg sand than 0 or 4 g CRF/kg sand.

Hydrilla shoot dry weight was greater with 4 g CRF/kg sand than 0 g. In the Experiment

2, we investigated the growth of naiad grown in 100:0, 75:25, 50:50, 25:75, and 0:100 sand:peat substrate fertilized with 0, 1, 2, or 4 g of CRF/kg substrate. We had three mesocosms per fertilizer rate with 5 pots (1 for each substrate). The experiment ran for

8 weeks in August 2014, and was repeated in June 2015. Naiad shoot dry weights were greater in 100:0 than 0:100 sand:peat substrates. Naiad root dry weight was greater in

50:50 than 0:100 sand:peat. The 0:100 substrate had higher substrate electrical conductivity (EC) levels than the other substrates. Naiad shoot dry weights were also greater with 1 to 2 g of CRF/kg substrate than 0 or 4 g CRF/kg substrate. There was no difference in tissue ammonia nitrogen (NH4-N) levels among the different fertilizer rates but phosphate (PO3-P) and potassium (K) were highest in naiad shoots fertilized with 1 and 2 g CRF/kg substrate than 0 or 4 g CRF/kg. The water analysis showed no differences in pH, EC, or K due to fertilizer rates but both nitrate nitrogen (NO3-N) and

PO3-P were higher in mesocosms with the 4 g CRF/kg substrate than the other rates.

Substrate EC concentrations also were highest in all substrates fertilized with 4 g

CRF/kg substrate. Based on these results we would recommend growing naiad in substrates with 50% or more sand and fertilized with 1 to 2 g CRF/kg substrate.

CHAPTER 1 INTRODUCTION

In the United States $100 million each year is invested for control of aquatic invasive weeds (Office of Technology Assessment [OTA], 1993). The most damage from invasive aquatic weeds occurs to our natural ecosystems in the Southern and

Western portions of the country (Pimental et al., 2005). Invasive plants alter the natural interactions in an aquatic habitat. Invasive plants are often responsible for reduction in oxygen levels, increases in water temperatures, and internal nutrient loading (Madsen,

1998). Control of aquatic weeds includes chemical control, biological control, and mechanical control methods. Many of these invasive aquatic weeds are controlled through the use of various herbicides, and mechanical harvesting methods (Wittenberg and Cock, 2001). In the process of removing these weeds, a vacuum is often created in these aquatic habitats that needs to be filled with natives to restore aquatic ecosystem stabilization.

Conservation of aquatic areas is crucial to Florida to promote a balanced ecosystem of food and habitats for fish and birds. Florida has 2.5 million acres of freshwater resources (UF/IFAS CAIP, 2015a). In Florida, invasive aquatic plants, including hydrilla (Hydrilla verticillata), and water hyacinth (Eichhornia crassipes) are altering aquatic habitats in addition to choking waterways, changing nutrient cycles, and reducing recreational use of rivers and lakes (OTA 1993).

Past estimates have shown, $14.5 million dollars annually is spent in Florida on hydrilla control. At that time, the state of Florida was experiencing a recreational loss of

$10 million dollars a year from problems caused by hydrilla vegetation growth in two major lakes (Center et al., 1997). Since then, the losses have substantially increased.

Southern naiad (Najas guadalupensis) is often mistaken for the invasive weed hydrilla and grows in similar areas. Unlike hydrilla, naiad is a native aquatic submersed plant in Florida. Southern naiad may prove to be a good candidate to reestablish some aquatic plant communities in natural areas previously damaged from invasive plants

(Smart et al., 1996). There has been much research published on hydrilla growth in relation to differing substrates and fertilizer levels (Sutton, 1990). However, little research has been conducted on the growth of naiad.

My hypothesis was that naiad could be grown in containers using similar substrates and fertilizer rates as described in the published hydrilla studies. The goal of this research was to generate data on the growth of naiad using standard greenhouse production techniques. The first objective was to compare hydrilla and naiad growth in a sand substrate fertilized with 4 rates of a controlled release fertilizer. The second, objective was to compare naiad growth in 5 substrates and with 4 controlled release fertilizer rates.

CHAPTER 2 LITERATURE REVIEW

According to the Southwest Florida Water Management District (SWFWMD,

2013), Florida contains 20% of all the wetlands in the United States. In these wetland areas various plant and animal ecosystems exist that are interdependent on each other for survival. Wetland areas are important because they serve as barriers against strong winds and storms and protect us from devastating flooding. Birds and fish also depend on wetland areas for breeding (SWFWMD, 2013). According to the Florida Fish and

Wildlife Conservation Commission (FWC), since Florida became a state, total wetland area has decreased by approximately 44% (FWC 2015).

Diversity of native aquatic plant communities is important to ecosystems because plants provide many habitats for fish and wildlife. Madsen (1998) reported that native plants support the natural interactions in aquatic communities. Native plant species are defined by having adapted and evolved with the competing species, predators, and diseases of an area over many thousands of years (Morse et al.,1999). They are in ecological balance with their surrounding environment. Furthermore, native aquatic plants improve water quality by removing nutrients such as nitrogen (N) and phosphorus

(P) as well as filtering storm water runoff, helping control erosion, and contributing to the aesthetic beauty of water bodies (Main et al., 2006).

The introduction of exotic (non-native) aquatic plants into streams, ponds, lakes, drainage canals, and ditches have become problematic in the United States as well as in Florida and is creating issues by altering the natural “environmental regimes”

(Godfrey and Wooten, 1981). Many exotic (non-native) aquatic plants have been introduced into the environment through travel, intentional input or accidental transport.

This inclusion in an environment causes it to quickly spread, dominating areas, changing habitats, and out competing the native species previously there. Several of these exotic introductions have become invasive. The United States Department of

Agriculture (USDA) defines an invasive species as non-native, alien to an ecosystem, and a species that are likely to cause economic or environmental harm or harm to human health (USDA, 2015a). Some of the most notable invasive aquatic plants in the

United States and Florida are hydrilla, Eurasian watermilfoil (Myriophyllum spicatum), and water hyacinth (Eichhornia crassipes).

Invasive aquatic plants tend to reduce species diversity, displace native plants, and alter the natural aquatic animal and plant interactions. Invasive plants can form dense surface canopies that block light from reaching lower growing native aquatic plants. These lower growing native plants are where small forage fish hide and grow to provide a food source to larger prey fish. The excessive growth of invasive aquatic plants negatively affects the growth of these fish and inherently affects predator fish ultimately causing them to move to other areas and reducing diversity in wildlife species

(Dibble et al., 1996).

Aquatic Plant Growth

All plants need carbon dioxide, oxygen, nutrients, water, and light in order to grow. During photosynthesis, plants use energy from the light to create carbohydrates and in the process release oxygen into the environment. Depending on the depth of the water in an aquatic system, the light levels will vary, as will the types of plants growing in these zones. A lake is made up of three zones. They are the limnetic or pelagic zone, the littoral zone, and the benthic zone. The limnetic zone is the open water portion where light is not able to penetrate to the bottom. This zone is characterized by being at

a depth where there are no rooted plants that can grow because sunlight cannot reach them. The bottom of the lake, which is covered in mud, is called the benthic zone

(MacIntyre and Melack, 1995). The shallow areas where there is enough light that can reach aquatic plants for growth is called the littoral zone. The plants that grow in this zone are divided into three categories; based on light availability, water depth of the zone, and plant characteristics. The first category is emergent plants. These plants grow in the shallow area of the littoral zone. They have the most availability of light, are rooted in the sediment of the lake or river, and their leaves and stems extend above the water surface. They root in saturated soil or in water up to 2 feet deep (Gibbons et al.,

1994). Examples of the types of plants in this category would be cattail (Typha latifolia) and exotic iris (Iris pseudacorus). The second category is composed of the floating leaf plants. The leaves of these plants float on the surface of the water. Within this category are free floating and rooted floating leaved aquatic plants. Some familiar free-floating aquatic plants are coontail (Ceratophyllum demersum), water hyacinth, water lettuce

(Pistia stratiotes) and duckweed (Lemna minor). While coontail does not have roots, some plants in this category have roots that hang freely beneath the plant but are not attached to the bottom sediment. Rooted floating leaved aquatic plants typically grow in depths of 1.5 to 10 feet and are rooted in the sediment with the leaves floating on the surface. An example of this is the water lily family (Nymphaeaceae).

The third category of plants, in the littoral zone is rooted submersed aquatic plants. These plants typically grow in the deepest parts of the zone where light is able to penetrate the water allowing the plants to grow (Barko and Smart, 1986). Rooted submersed plants among many species tend to grow with their stems and leaves

underwater. They often have long thin and flexible stems and show a variation of leaf forms. Leaf forms can vary from long and ribbon like such as with curly pondweed to feathery type whorls such as Eurasian watermilfoil. Many submersed plants send shoots to the surface of the water and produce flowers. Examples of submersed plants are southern naiad and hydrilla. Generally, submersed plants are found growing in depths of 6 feet to 30 feet. However, submersed aquatic mosses (Drepanocladus sp. and Fontinalis sp.) have been located growing in Crater Lake in Oregon at depths of

200 feet (Madsen, 2009). This variation is due to the variation of trophic state of the water.

The trophic state of the water refers to the nutrition in the water body and its ability to support plants, fish, and wildlife (Florida Lakewatch, 2002). When discussing growth of aquatic plants, it is vital to clarify that growth often is directly related to the trophic state of the water body. For example, floating-leaved and emergent growth forms generally produce more biomass than submersed growth forms and have proportionately greater demands for nutrients (Ott et al., 2012).

There are four different types of trophic states based on chlorophyll, phosphorus

(P), nitrogen (N), and water clarity. An oligotrophic state has the lowest level of biological productivity and tends to be clear with few aquatic plants, few fish, limited wildlife, and sandy bottoms. Chlorophyll is less than 3 ᶙg/L, P is less than 15 ᶙg/L, N is less than 400 ᶙg/L, and water clarity is greater than 13 feet. A mesotrophic state has a moderate level of biological productivity. Mesotrophic waterbodies have a medium level of aquatic plants and are moderately clear, between 8 and 15 feet. The chlorophyll is between 3 and 7 ᶙg/L, P is between 15 and 25 ᶙg/L, and N is between 400 and 600

ᶙg/L. A eutrophic waterbody has a high level of biological productivity. It has a lot of potential to support many fish and wildlife. The clarity is between 3 and 8 feet.

Chlorophyll is between 7 and 40 ᶙg/L. Phosphorus is between 25 and 100 ᶙg/L.

Nitrogen is between 600 and 1500 ᶙg/L. The fourth trophic state is a hypereutrophic waterbody. This has the highest level of biological productivity. The water clarity in this type of is less than 3 feet. It has the largest amount of aquatic plants, fish, and wildlife.

The chlorophyll criteria was greater than 40 ᶙg/L. Phosphorus will be greater than

100ᶙg/L. Total N is greater than 1500 ᶙg/L (Florida Lakewatch, 2002).

Invasive Aquatic Plant Control

There are several methods to control and manage aquatic invasive weeds.

Cultural controls can educate the public about prevention methods like inspecting boats for the presence of invasive plants before launching in a new area. Physical control methods are characterized by non-motorized action plans to control the spread of the aquatic weed such as hand pulling, water-level drawdowns, and benthic barriers. In water-level-draw downs, the water level is lowered in an attempt to expose the target weeds to atmospheric conditions that will kill it. Benthic barriers are a method to control aquatic weeds through blocking sunlight for a few months to kill the submersed plants.

This method is most commonly used in small areas because it can negatively affect the fish and wildlife in the water body over an extended period of time (Gibbons et al.,

1994). It is also expensive, non-specific, and requires maintenance.

Other methods of control and management include mechanical harvesting and biological control methods. Mechanical harvesting methods use a motorized machine that has a spinning mechanism that aims to rake, cut, and draw up the weeds at a certain depth. These machines range from cutter boats, to aquatic rototillers, to more

customized types of design. Mechanical harvesting tends to be a very expensive means of control that removes invasive weeds for a short term. In addition to being costly, many invasive aquatic weeds spread through fragmentation. Furthermore, desirable vegetation often is ripped out in the process, and fish are removed (Haller et al., 1980).

The clearing tends to be temporary, with weeds growing back 2 and 7 months later

(Netherland and Jones, 2012).

Biocontrol methods involve the action of using plant-eating animals such as grass carp (Ctenopharyngodon idella), weevils (Curculionoidea), or parapoynx moths

(Parapoynx diminutalis) to suppress the excessive growth of invasive weeds. To be a true bio-control, the animal must be plant specific. For example, these methods have had success in reducing growth of invasive alligatorweed (Alternanthera philoxeroides) and melaleuca (Melaleuca quinquenervia), but less success in controlling waterhyacinth, Eurasian watermilfoil, or hydrilla (Harley and Forno, 1992).

The last type of control method most often used is two types of herbicides: contact herbicides and systemic herbicides. Contact herbicides are immovable herbicides that cause death to the plant tissue that is comes in contact. Some of the more common contact herbicides used for aquatic weed control are Diquat, Endothall, and Carfentrazone. Contact herbicides are often used to control waterlettuce, duckweed, and Salvinia. Contact herbicides used on submersed plants remain in the water around the target plant. Systemic herbicides work by being absorbed by the plant tissue and its uptake systems taking weeks to actually kill the plant. Systemic herbicides move through the plant tissue and aim to inhibit enzymes that plants need to produce proteins for growth. Some systemic herbicides are triclopyr and 2,4-D and they are used

to control submersed, floating, and emergent aquatics. Fluridone is also a type of systemic herbicide that is often used for floating, and submersed plants to manage whole lake areas (Netherland, 2009).

The application methods are the same for both contact and systemic herbicides for emergent plants. They are applied directly to the foliage and used to control floating and emergent aquatic plants. Submersed plant herbicides are applied as a concentrated liquid, granule, or pellet. The volume of the water has to be calculated before application to ensure even application relevant to the volume of the water body area being treated.

When determining the best method for control and management of invasive aquatic weeds, there are some important factors to be considered. It has been assumed that following an herbicide application, nutrients from the decaying weeds are put back into the ecosystem and can contribute to regrowth of new weeds, or more regrowth of the targeted weed (Sutton and Portier, 1983). Therefore, a plant manager needs to consider the nutrient capacity of the plant being controlled. If the plant is a lower nutrient plant it may be more cost effective to use herbicidal control methods versus the more expensive mechanical harvesting methods. However, the benefit of mechanical harvesting is being able to eliminate the nutrient loading in the ecosystem from aquatic weeds with high nutrient contents (Sutton and Portier, 1983). A drawback of mechanical harvesting is that it can be quite expensive.

Naturally, there is more concern about invasive aquatic plants threatening eutrophic and hypereutrophic water bodies because they have the highest levels of biological productivity, and can cause greater disturbance to their ecosystem. For

example, growth of Eurasian watermilfoil, an invasive aquatic species in a lake in upstate New York, was excessive and reduced the total number of species in the research plot from 21 to 9 over a three-year period (Madsen, 1998).

Invasive Aquatic Plants in the United States

Eurasian watermilfoil is an invasive submersed plant that is rooted in the sediment and grows completely underwater, but forms a canopy at the water surface. It reproduces mostly by fragmentation when the stems are broken through efforts to mechanically harvest it. Currently, it is controlled most effectively through the use of contact herbicides such as diquat, and fluridone (Smith and Barko, 1990).

The invasive aquatic curlyleaf pondweed is a submersed plant that grows in lakes, and rivers and out competes other submersed natives. This plant is most problematic in the northern states, and is controlled most effectively by early season treatment with the herbicides such as diquat, endothall, fluridone, penoxsulam, and imazamox, (Netherland 2009).

Egeria densa (Brazilian elodea) is a rooted submersed weed aquatic plant that is believed to have originated from South America in the 1890’s. It is similar to Eurasian watermilfoil in that it can withstand low light and low water temperatures. Egeria is similar to other submersed invasive plants in that it can reproduce through stem fragments, and therefore boat traffic can inadvertently spread it. However, it differs like eurasian watermilfoil in that it does not produce tubers and has poor heat tolerance in warm water above 90° C (Pennington, 2009). Aquarium trade is believed to be responsible for it being so widespread across the US. It is a monocot species that grows in a variety of fresh water and flowing water (USDA, 2015b).

Water hyacinth is a floating aquatic plant native to the Amazon River in South

America. It is a floating flowering monocot that can grow as an annual or perennial depending on the region. It was first introduced to North America in 1884 in New

Orleans, and then spread to other areas of the southeastern United States, California,

Hawaii and the Virgin Islands. It is currently classified as an invasive noxious plant

(FLEPPC). It requires temperatures of greater than 50 F for growth. Water hyacinth reproduces by seedlings and rosettes, which are plants attached to the main plant by a floating runner. Humans spread it indirectly through inclusion in ponds, or waterways. It is best controlled using weevils, in conjunction with foliar contact spray herbicides such as diquat and endothall (Gopal, 1987).

Water chestnut is a floating leaved plant believed to be introduced to the United

States from Asia in the 1870s. Some varieties are grown as an agricultural food crop in parts of India and China. It has submersed leaves that are 4 inches long and attached to the stem in a whorl. It reproduces by seeds and it has been believed to be spread by human transport. It produces a dense canopy in waterways that has the ability to reduce and obstruct sunlight from reaching the other submersed aquatic vegetation. This can cause a reduction in the habitat from the lack of sunlight. The preferred method of control is the use of 2,4-D and triclopyr herbicides (Netherland, 2009).

Hydrilla is an invasive submersed monocot aquatic weed introduced from Asia into the state of Florida in 1950-51. It hinders navigation and drainage of canals, rivers, lakes, and waterways in Florida (Haller et al, 1976; Gordon and Thomas, 1997; Schmitz et al.,1991; Schardt and Nall, 1983). Hydrilla is in the Hydrocharitaceae family. The specific epithet verticillata designates the characteristic of the plant leaves, which grow

in whorls of four to eight around the stems. Hydrilla is either monoecious, with both female and male flowers on the same plant, or dioiceous (male and female flowers on separate plants). It is believed that the dioiceous type was introduced into Florida through the aquarium trade business much like Egeria (Kay, 1992).

Hydrilla is a slender green long plant that grows up to 25 feet in length. It is a multibranched plant with stems that are long and flexible often growing at a rapid rates and forming dense canopies at the surface of the water that block out light from other lower growing plants. The leaves of hydrilla are attached to the stem of the plant.

Hydrilla does not produce seeds in Florida because it is only dioiceous, but easily spreads by vegetative fragmentation (Ramey, 2001). Boat traffic can easily spread this invasive weed between areas. Hydrilla, unlike Egeria, produces turions and tubers.

Hydrilla is identifiable by digging up the roots and seeing if there are tubers and turions among them.

The female flowers of hydrilla are tiny and white. Male flowers are tiny and greenish in color and pollinate the female flowers at the surface of the water by bumping into them. Hydrilla can grow in a few inches of water or in more than 20 feet of water. It does not need much light penetration and can grow in 1% full sunlight (Ramey, 2001).

Hydrilla has an optimum growth temperature of 20-27 o C (68-81 o F). It also has a maximum temperature, 30o C (86o F) (Kasselmann 1995). Hydrilla has often been confused with native aquatic plants such as southern naiad.

Control methods for hydrilla are very costly. Most hydrilla control management programs use mechanical harvesting, biological control methods such as Grass Carp, and/or herbicides. Mechanical harvesting methods are short lasting. Herbicidal methods

such as diquat and slower systemic herbicides such as fluridone have proven to be effective controls for hydrilla (Madiera et al, 2000). Hydrilla is listed as a Category 1 invasive plant on the Florida Exotic Pest Plant Council’s 2015 list of invasive plant species. A category 1 invasive means that it alters native plant communities by displacing native plant species and changes ecological functions. This designation is given based on the damage it causes. The Florida Exotic Pest Plant Council revises their list every two years based on the accounts of professional botanists, and other industry professionals (FLEPPC, 2015). Hydrilla is distributed throughout the state and is classified as a noxious weed that is a prohibited plant by the Florida Department of

Agriculture and Consumer services (FDACS, 2015).

Because hydrilla is one of the most troublesome invasive aquatic weeds in the state of Florida, much research on growth of hydrilla has been conducted in order to develop better control methods. For example, research to identify seasonality determined that hydrilla is a perennial plant and that the best seasonal period for control is more closely related to annual plant management, which would be in the early fall, after the tubers have germinated, and before new tubers begin (Haller et al., 1976).

There also have been various studies associated with the nutrients in hydrilla taken from several heavily infested areas throughout the state of Florida (Sutton and

Portier, 1983). Sutton and Portier (1983) compared the contents of N, P, and potassium

(K) to identify if hydrilla was a high nutrient weed or a low nutrient weed. The study found that in hydrilla taken from Lake Okeechobee had higher concentrations of N than hydrilla collected from the other areas in Florida sampled. It was determined that the sediments in the sample areas varied and that there may be a relationship between the

sediment of the sampled area and the nutrients in the plants. This led to more research on the sediments of the hydrosoils in which the invasive aquatic weed was rooted.

Steward (1984) grew hydrilla in hydrosoils of different compositions. It was determined that the tissue N, P, and K contents were related to the N, P, and K contents in the soil.

Phosphorus and N were more dependent on the fertility of the hydrosoil than the nutrients in the water around the plant. Barko et al. (1986) compared the growth of

Eurasial watermilfoil and dioecious hydrilla in relation to the organic sediment matter content. They found that the weight of the Eurasian watermilfoil and hydrilla decreased more rapidly as organic matter content in the sediment increased from 0% to 20%. This happened because higher absorption. However, other studies (Kiorboe, 1980; Spencer et al., 1992; Sondergaard and Sand-Jensen 1979) reported increased growth of submersed plants such as watermilfoil and hydrilla with increased sediment organic matter content. There also have been studies using emergent aquatic plants (Sharp and

Keddy, 1985; Lee et al., 1986) that support increased growth with increased organic sediment matter content in the aquatic plants studied. Spencer et al. (1992) investigated the growth of monoecious hydrilla on different soils amended with peat or barley straw up to 20%. They measured the weight and tuber growth of the hydrilla and found that the growth of hydrilla was influenced by the soil type as well as the amendment of organic matter. Their study showed that the un-amended sand substrate had the lowest growth.

Gunnison and Barko, (1989) conducted a study that suggested that altering the organic matter content of the sediments may be a potential method for managing growth of rooted aquatic plants. There was also a later study about the organic

sedimentation that is produced from hydrilla management. It found that less tubers and organic sediment were produced when hydrilla was treated with herbicides 2/3 of the way to the surface (Joyce et al., 1992).

Sutton (1985) grew hydrilla in a muck sand soil mixture, sand alone, and sand amended with 3 commercially available fertilizers at 3 different rates (low medium and high g/L). Tuber production was independent of the fertilizer rates. Water temperature did influence the dry weight of the plant and that dry weight was higher in the amended sand with fertilizer than in the muck sand soil mixture, or sand alone substrate. Only P in both the roots and shoots was dependent on the level of fertilizer in the root zone.

Sutton’s (1985) findings support the idea that the growth of hydrilla was controlled by the nutrients in the root zone. The take away message from this experiment was to use sand amended with different levels of fertilizers as a way to simulate the fertility levels of sediment in aquatic sites that supported hydrilla growth. He reported that the use of resin-coated fertilizer pellets was a good method to control the amount of nutrients in the root zone in mesocosms.

Resin-coated fertilizer products like Osmocote (Scotts Miracle-Grow, Everris

Dublin, OH) are classified as controlled-release fertilizers. There are different types of controlled release fertilizer products based on the method of fertilizer release and include plastic encapsulated (resin) fertilizers, slowly soluble fertilizers (i.e. lime), urea aldehydes, sulfur coated fertilizers, and chelated micronutrients (Nelson, 1998). The plastic encapsulated (resin) fertilizers release nutrients when water vapor moves into the pellet and dissolves the fertilizer. As more water moves into the capsule, the dissolved fertilizer is “squeezed” out” (Warncke and Krauskopf, 1983). The availability of

the fertilizer is regulated by pinholes in the coating. Moisture enters the encapsulated fertilizer pellet through the pinholes and dissolves the fertilizer salts. Upon the entrance of water, the solution pressure rises and disrupts the coating. The solution then flows out by gravity (Tisdale et al., 1985). Release rate can be controlled by the thickness of the coating as well as temperature and varies with the manufacturer of the product.

Products with multiple layers of resin tend to have a thicker coating that provides slower release of nutrients as compared to thinner coatings with more rapid release of nutrients. Studies have shown that when the temperature is raised that the rate of release can be doubled (Tisdale et al., 1985).

Southern Naiad

Southern naiad is a Florida native submersed aquatic plant that is often confused with hydrilla. Southern naiad is an annual plant, but a perennial in Florida that is in the family Najadaceae. It is a monocotyledonous submersed aquatic plant of the genus

Najas having narrow leaves and small flowers (UF/IFAS CAIP, 2015b). There are four species of Najas but the most common are N. guadalupensis, and N. minor (Missouri

Department of Conservation [MDC], 2015. It is rooted in the sediment. The leaves are less than 1/2 inch long and 1/8 inch wide and have fine-toothed edges and are in pairs or whorls. There are tiny greenish flowers that grow at the leaf bases (MDC, 2015). It produces seeds that are eaten by wildlife. The seeds of the plant are the means by which it reproduces as well as by fragmentation. Single seeds are found encased in the leaf sheath (Texas A & M Extension [TAMU], 2015). Naiad has been found growing in lakes, ponds, and other slow moving waterways (MDC 2015).

Southern naiad is distributed widely throughout the United States (Fernald, 1923) and has adapted to local environments and seasonal variations in Florida (Main et al.

2006). This makes it a good candidate for ecosystem restoration activities because it requires low fertilizer inputs to become established (Main et al., 2006). It is an important part of the aquatic ecosystem as it provides a habitat to wildlife and food for fish, waterfowl, and other birds (TAMU, 2015). Barnett and Schneider (1974) mentioned southern naiad as being beneficial to fish in plant aquatic communities in Florida.

Buddington (1979) reported that southern naiad was a preferred food in the digestion tracks of tilapia fish (Oreochromis).

Most of the research on southern naiad has been in reference to fish communities rather than on its growth habits. There was one study conducted on the control of southern naiad in 1962 done in Florida irrigation channels (Blackburn and

Weldon, 1962). However, past research has not determined if southern naiad is a high nutrient plant or how it might add to the sediment in an aquatic environment. Research on the growth of southern naiad will help expand its use in natural aquatic ecosystems.

CHAPTER 3 GROWTH OF NAIAD (NAJAS GUADALUPENSIS) AND HYDRILLA (HYDRILLA VERTICILLATA) USING CONTROLLED RELEASE FERTILIZER

The University of Florida Center for Aquatic and Invasive Plants reported that

Florida has 2.5 million acres of aquatic freshwater resources (UF/IFAS CIAP, 2015a).

Conservation of these areas is crucial to the state of Florida because they promote a balanced ecosystem in the form of food and habitats for fish and birds. Non-native plants tend to alter the natural interactions in an aquatic habitat (Madsen 1998). Non- native plants are often responsible for reduction in oxygen exchange, depletion of dissolved oxygen, increases in water temperatures, and internal nutrient loading

(Madsen, 1998).

A nuisance in Florida waters is the invasive aquatic weed, hydrilla (Hydrilla verticillata) that was introduced into the state in 1950-51. It hinders navigation and drainage of canals, rivers, lakes, and waterways in Florida (Gordon and Thomas, 1997).

It is essential to keep waterways clear as they provide a mechanism to reduce flooding of many land areas in Florida including farms and urban areas. The invasion of hydrilla also has decreased biodiversity, and threatens to push out native species. Hydrilla can be found in a variety of water body types and is known to easily spread by fragmentation (Ramey, 2001). The spread of hydrilla has shown substantial increase over the years (Schmitzt et al.,1991; Schardt and Nall, 1983).

The native aquatic plant southen naiad (Najas guadalupensis) is often confused with the invasive weed hydrilla. Southern naiad is a desirable native submersed aquatic plant in Florida. It has dark green leaves and shoots that grow long and thin like a ribbon. It reproduces by seeds and fragmentation and provides a habitat for invertebrates, food for fish and wildlife, and helps to remove some nutrients from the

water (TAMU, 2015). Hydrilla is a submersed aquatic plant that has thin green stems that can grow up to 25 feet long, and small leaves that grow in whorls of four to eight around the stem with branching occurring near the water surface (Ramey, 2001).

Most of the research on southern naiad has been in reference to fish communities rather than on its growth habits (Buddington, 1979; Barnett and Schneider,

1974). However, there has been much research on hydrilla growth in relation to differing fertilizer levels (Sutton and Portier, 1983; Steward, 1984; Sutton, 1985). Sutton (1985) used sand amended with different levels of resin-coated fertilizers as a way to simulate the fertility levels of sediment in aquatic sites that supported hydrilla growth. He reported that the use of resin-coated fertilizer pellets was a good method to control the amount of nutrients in the root zone in mesocosms. The objective of this experiment was to compare hydrilla and naiad growth in sand amended with four controlled-release fertilizer rates.

Materials and Methods

Cuttings approximately 4-6 inches long of hydrilla and naiad were obtained from the aquatic tanks located at the University of Florida Fort Lauderdale Research and

Education Center (FLREC), Davie, FL. Ten cuttings of each species were placed into 8- inch diameter round azalea pots, lined with plastic bags and filled with coarse builder’s sand. Prior to transplanting the cuttings, a controlled release fertilizer (Osmocote 15N-

4.05P- 9.96K, Scotts Miracle-Grow, Everris Dublin, OH) was layered under the surface of the sand at 0, 1, 2 or 10. 4 g of fertilizer per kg of substrate.

A randomized block design was utilized for this experiment with 24 mesocosms

(12 with naiad and 12 with hydrilla) (Fig 3-1). Each mesocosm was an 18-gallon plastic rectangular tub with three pots (30 cuttings per mesocosm). Each mesocosm contained

one fertilizer rate (group). The mesocosms were divided into 3 blocks of 4 groups. Each group represented a different fertilizer level (0, 1, 2, and 4 g of Osmocote CRF/kg of sand). The mesocosms were filled with rainwater collected from the surface ponds at the center to create a submersed growing environment. Mesocosms were arranged in an open-sided greenhouse at the University of Florida (FLREC) and exposed to ambient air temperatures and light levels (average air temperature 25 to 27 ºC, 76 to 80% RH, and average solar radiation in the greenhouse 174 W·m2)

The experiment was conducted for 8 weeks in spring 2014 and was repeated in summer 2014. After 8 weeks, shoots and roots were harvested from each pot in the mesocosm and combined to create one shoot and one root sample per mesocosm.

Plant tissue was washed clean of debris using a screen for straining. Shoots and roots were dried in a forced air oven set at 90 ºC until constant weight was achieved.

Dry weights were subjected to analysis of variance and least significance separation of means to identify the differences using ANOVA in R statistical program, version 3.2.0. The data from the two experiments was analyzed separately. Significant means were identified at the 0.05 significance level and separated by the Duncan

Multiple Range Test.

Results and Discussion

For experiment 1, there was no difference in shoot or root dry weight between naiad and hydilla. Naiad and hydrilla shoot dry weight was significantly greater in pots with 2 g CRF/kg sand, than with 0, 1, or 4 g CRF/kg sand (Table 3.1). Naiad and hydrilla root dry weight was greatest in containers with 1 g CRF/kg sand. Previous research with hydrilla suggested that higher nutrient levels in sediments resulted in greater growth of hydrilla, and that diminished growth was related to limited nutrients

(Barko, 1982). Our results may have been influenced by an infestation of parapoynx moths (Parapoynx diminutalis) decimating shoots a week prior to harvest (Fig 3-2).

Hydrilla shoot and root growth has characteristically been shown to increase with the addition of fertilizer (Sutton and Portier,1983; Sutton 1985).

In experiment 2, no insect pest problems were observed. Overall, hydrilla shoot dry weight was greater than naiad shoot dry weight (Table 3-1). Naiad shoot dry weight in pots with 1 g and 2 g CRF/kg sand was greater than in pots with 0 or 4 g CRF/kg sand. Naiad root dry weight was greater in 2 g CRF/kg sand than 0 g CRF/kg sand, but not different from 1 g or 4 g CRF/kg sand. Hydrilla shoot dry weight was greater in 4 g

CRF/kg sand than 0 g, but not different from 1 or 2 g CRF/kg sand. Hydrilla root dry weight was greater in 2 g and 4 g CRF/kg sand, than 0 g (Table 3-1). Barko (1982) found that there was a relationship in biomass growth of hydrilla to the differing fertilizer levels in the substrate with higher fertilizer rates resulting in more growth. Steward

(1984) grew hydrilla in hydrosoils of different compositions. It was determined that the tissue nitrogen (N), phosphorus (P), and potassium (K) contents were related to the N,

P, and K contents in the soil. Phosphorus and N were more dependent on the fertility of the hydrosoil than the nutrients in the water around the plant. Sutton (1985) grew hydrilla in a muck sand soil mixture, sand alone, and sand amended with 3 commercially available fertilizers at 3 different rates (low, medium, and high). Tuber production was independent of the fertilizer rates. Water temperature did not influence the dry weight of the plant. However, hydrilla shoot dry weight was higher in the amended sand with fertilizer than in the muck sand soil mixture, or sand alone

substrate. Only P in both the roots and shoots was dependent on the level of fertilizer in the root zone.

Southern naiad is distributed widely throughout the United States (Fernald, 1923) and has adapted to local environments and seasonal variations in Florida (Main et al.

2006). This makes it a good candidate for ecosystem restoration activities because it requires low fertilizer inputs to become established (Main et al., 2006). Large concentrations of naiad tend to be found in the central and southern parts of Florida where soils are mostly sandy loam and low in nutrients (Novak et al, 2009). This may explain why naiad performed better at lower fertilization rates compared to hydrilla.

Although these plants may look similar, it is clear from this study that their nutritional requirements are different.

Table 3-1. Southern naiad (Najas guadalupensis) and hydrilla (Hydrilla verticillata) shoot and root dry weight of plants grown in sand and fertilized with 0, 1, 2, or 4 g CRF/kg sand (Osmocote 15N-4.05P- 9.96K). Experiment 1 was conducted in spring 2014 and experiment 2 was conducted in summer 2014 at the University of Florida, Fort Lauderdale Research and Education Center. Means followed by different letters are significantly different at 0.05 level. Fertilizer Naiad Hydrilla (g/pot) Shoot dry Root dry Shoot dry Root dry weight (g) weight (g) weight (g) weight (g) Experiment 1 (Spring 2014)

0 26.9B 23.5B 26.8B 23.9B 1 27.8B 26.5A 27.9B 28.5A 2 34.1A 24.6B 32.3A 23.4B 4 30.2B 24.3B 29.5B 22.0B

Experiment 2 (Summer 2014)

0 14.6B 6.6B 12.2B 8.4B 1 19.1A 8.7AB 24.7AB 9.3AB 2 18.9A 12.2A 25.1AB 10.6A 4 13.4B 10.1AB 29.1A 10.9A

Figure 3-1. This is an example of the 18-gallon plastic containers (mesocosms) and experimental layout. Each mesocosm contained three pots of either hydrilla or naiad. The containers were lined with plastic bags and filled with sand. Each mesocosm contained only 1 or 4 fertilizer rates (0, 1, 2, or 4 g per pot). Three mesocosms of each fertilizer rate for each species were arranged in an open sided greenhouse at the University of Florida Fort Lauderdale Research and Education Center. Photo credit: H. Hasandras

Figure 3-2. During experiment 1, a parapoynx moth (Parapoynx diminutalis) was found at week 7 of the experiment. By the time it was found, it had damaged the shoots of several hydrilla and naiad plants in the experiment. Photo credit: H. Hasandras

CHAPTER 4 GROWTH OF NAIAD IN SUBSTRATES WITH VARYING PERCENTAGES OF SAND AND CONTROLLED-RELEASE FERTILIZER

Conservation of aquatic areas is crucial to Florida to promote a balanced ecosystem of food and habitats for fish and birds. Florida has 2.5 million acres of freshwater resources (UF/IFAS CAIP, 2015a). In Florida, invasive aquatic plants, including hydrilla (Hydrilla verticillata), water hyacinth (Eichhornia crassipes), and water lettuce (Pistia stratiotes), are altering aquatic habitats in addition to choking waterways, changing nutrient cycles, and reducing recreational use of rivers and lakes

(OTA 1993).

Southern naiad (Najas guadalupensis) is often mistaken for the invasive weed hydrilla and grows in similar areas. Unlike hydrilla, naiad is a native submersed aquatic plant in Florida. Southern naiad may prove to be a good candidate to reestablish some aquatic plant communities in natural areas previously damaged from invasive plants

(Smart et al., 1996). There has been much research published on hydrilla growth in relation to differing substrates and fertilizer levels (Sutton, 1990; Mony et al., 2007).

However, little research has been conducted on the growth of naiad.

Most of the research on Southern naiad has been in reference to fish communities rather than on its growth habits. There was one study conducted on the control of southern naiad in 1962 done in Florida irrigation channels (Blackburn and

Weldon, 1962). When aquatic plants die they add their nutrients to the water. However, past research has not determined if Southern naiad is a high nutrient plant or how it might add to the sediment in an aquatic environment. Research on the growth of

Southern naiad could help expand its use in natural aquatic ecosystem. The objective of

this study was to compare growth of Southern naiad in substrates with different percentages of sand and fertilized with four levels of a controlled-release fertilizer.

Materials and Methods

Cuttings approximately 4-6 inches long of naiad were obtained from the aquatic tanks located at the University of Florida Fort Lauderdale Research and Education

Center (FLREC), Davie, FL. Ten cuttings were placed into 8-inch azalea pots and filled with 100:0, 75:25, 50:50, 25:75 or 0:100 coarse builders sand:peat (by volume). Prior to transplanting the cuttings, a controlled release fertilizer (CRF) (Osmocote 15N-4.05P-

9.96K, Scotts Miracle-Gro, Everris Dublin, OH) was top layered into the substrates at 0,

1, 2 or 4 g CRF/kg of substrate per container.

A randomized block design was utilized for this experiment with 12 mesocosms

(3 blocks per fertilizer rate 0, 1, 2, and 4 g CRF/kg substrate). Each mesocosms contained one container of each substrate for that fertilizer rate.

The mesocosms were plastic 18-gallon rectangular tub filled with rainwater from surface ponds at the center to create a submersed growing environment. Mesocosms were arranged in an open-sided greenhouse at the FLREC and exposed to ambient conditions (average air temperature 25 to 27 ºC, 76 to 80% RH, and average solar radiation in the greenhouse 174 W·m2). In the first experiment the containers with

50:50, 25:75, and 0:100 sand:peat substrate were topped with small rocks to prevent floating once submersed in the mesocosms, then the cuttings were added between the pebbles into the substrate. In the second run of this experiment, the containers with

50:50, 25:75, and 0:100 sand:peat were topped with weed cloth over the substrate.

Holes were made in the weed cloth to insert the cuttings into the substrate, and then the

container was top dressed with small pebble rocks to prevent flotation.

The experiment ran for 8 weeks in August 2014 and was repeated in June 2015.

After 8 weeks, shoots and roots were harvested from each container in the mesocosm for both experiments. Plant tissue was washed clean of debris using a screen for straining. Shoots and roots were dried in a forced air oven set at 90 ºC until a constant weight was achieved.

In the June 2015 experiment, water samples were collected to determine pH electrical conductivity (EC), nitrate (NO3-N), phosphate (PO3-P) and potassium. Soil samples also were collected to determine final pH and EC using the 1:2 extraction method. Water and substrate pH and EC were determined using a Hanna Combo pH and EC waterproof meter # H198129 (Hanna Instruments, Woonsocket, RI). Water

NO3-N, and K were determined using an Accumet XL250 combination ion selective electrode (Fisher Scientific, Waltham, MA). Phosphate was determined by the ascorbic acid method using a Spectronic 20D+ spectrophotometer (Thermo Electron

Corporation, Waltham, MA) measuring absorbance at 440 nm. The standard curve measured between 0 and 1 mg/g. Sample preparation involved dilution in a 5 mL vial to an appropriate concentration for interpolation on the standard curve.

Shoots from the second experiment also were ground using a Wiley Mill (Thomas

Scientific, Swedesboro, NJ) to pass through a 40-mesh screen to ensure homogeneity.

The shoots (0.5 g) were placed into a 30 mL porcelain crucible and dry ashed to determine NH3-N, PO3-P and K using a Thermolyne Thermo Scientific muffle furnace

(Thermo Fischer Scientific, Waltham, MA) capable of 500 ºC. The ash was dissolved with 10mL 1.0N HCl solution. The contents of the crucible for each sample were then

transferred into a 50 mL volumetric flask, and diluted with deionized water to volume, capped, and inverted 3 times. Samples were filtered through Whatman 41 filter papers into 50mL plastic bottles. This digestate was then used for nutrient analysis of PO3-P,

NH3-N, and K. Samples were prepared for analysis by dilution in 5mL vials so that concentration could be interpolated on the standard curve. Analysis for NH3-N and

PO3-P were performed by colorimetry with a Seal Autoanalyzer 3. Analysis for K was performed by atomic absorption spectrometry with a PerkinElmer AAnalyst 400.

Dry weights, water analysis, substrate leachate analysis, and shoot nutrient analysis were subjected to analysis of variance and least significance separation of means to identify the differences using ANOVA in R statistical program, version 3.2.0.

The data from the two experiments were analyzed separately. Significant means were identified at the 0.05 significance level and separated by the Duncan Multiple Range

Test.

Results and Discussion

For both experiment 1 and 2, there were no significant interactions for shoot and root dry weight between substrate and fertilizer rate. Overall, in both experiment 1 and

2, the shoot dry weight was greater in the 100:0 sand:peat substrate than 0:100 mixture

(Table 4-1). In experiment 1, there was no difference in shoot dry weight among 100:0,

75:25, 50:50, or 25:75 sand:peat substrates. In experiment 2, shoot dry weight was also similar among 100:0, 75:25 and 50:50 substrates. The root dry weight was greater in both experiments 1 and 2 in the 50:50 sand:peat than 0:100 sand:peat, but not different from 100:0, 75:25, or 25:75. (Table 4-1). Southern naiad is distributed widely throughout the United States (Fernald, 1923) and has adapted to local environments and seasonal variations in Florida (Main et al., 2006). This makes it a good candidate

for ecosystem restoration activities because it requires low fertilizer inputs to become established (Main et al., 2006). Large concentrations of naiad tend to be found in the central and southern parts of Florida where soils are mostly sandy loam and low in nutrients (Novak et al, 2009). Similarly, hydrilla dry weight was higher in the amended sand with fertilizer than in the muck sand soil mixture, or sand alone substrate

(Sutton,1985). Only P in both the roots and shoots was dependent on the level of fertilizer in the root zone.

In both experiment 1 and 2, the shoot dry weight at fertilizer rates of 1 and 2 g

CRF/kg substrate were greater than 0 g and 4 g CRF/kg substrate (Table 4-2, Figure 4-

1). Similarly, in both experiments root dry weight was greatest with 2 g CRF/kg substrate than 0, 1, or 4 g CRF/kg substrate (Table 4-2). There were no significant substrate x fertilizer interaction or significant differences in shoot NH3-N, PO3-P, or K due to substrate. There also were no differences in shoot NH3-N due to fertilization

(Table 4-3). However, shoot PO3-P and K was greater in pots fertilized with 1 and 2 g

CRF/kg substrate than 0 or 4 g CRF/kg substrate. In previous research on another aquatic plant, Steward (1984) determined that the tissue N, P, and K contents in hydrilla were related to the N, P, and K contents in the soil. Phosphorus and N were more dependent on the fertility of the hydrosoil than the nutrients in the water around the plant. In our experiment, the water analysis showed no differences in pH, EC, or K due to fertilizer rates but both NO3-N and PO4-P were higher in mesocosms with the 4 g

CRF/kg substrate fertilizer rate than the other rates (Table 4-4).

Bunt (1974) examined the physical and chemical characteristics of various peat:sand and peat:vermiculite substrate on the release of nutrients from controlled-

release fertilizers and reported that leaching losses of N, P, and K were less in the peat:vermiculite substrate than in the peat:sand substrate. Substrate pH was 5.9 to 6.3 and was not different due to percentage of sand or fertilizer rate. However, EC levels were highest in the 0:100 substrate with no difference among the other substrates.

Substrate EC was highest in all of the substrates was at 4g CRF/kg substrate (Figure 4-

2). It is possible that the 0:100 substrate retained more fertilizer leading to higher soluble salt levels in the substrate and reduced naiad growth. Reduced growth and reduced shoot nutrient concentrations were observed in all of the plants fertilized with 4 g CRF/kg substrate.

One question we wanted to try to answer was if Southern naiad was a high nutrient plant like hydrilla and how it might add to the sediment in an aquatic environment. Sutton (1985) grew hydrilla in sand amended with 0 to 35 g of Osmocote and reported that maximum growth occurred at 25 to 35 g of Osmocote. He reported high levels of P in hydrilla shoots associated with high concentrations of Osmocote

(range of 384 to 648 ᶙg/g P). However, N in hydrilla shoots was the same in sand without fertilizer compared to plants grown with fertilizer (1.7 to 2.4 %N) and K in shoots was the same regardless the amount of fertilizer (20.8 to 22 mg/g K) (Sutton, 1985). In our study naiad growth was less in substrates with 4 g CRF/kg substrate, suggesting that it is not a high nutrient plant like hydrilla. Based on the results from this experiment, we would suggest growing southern naiad in substrates with 100:0 or 75:25 sand:peat and fertilized with 1 to 2 g CRF/kg substrate.

Table 4-1. Naiad (Najas guadalupensis) shoot and root dry weight of plants grown in 100:0 sand:peat, 75:25 sand:peat, 50:50 sand:peat, 25:75 sand:peat or 0:100 sand:peat (by volume). Experiment 1 was conducted August 2014 and experiment 2 was conducted June 2015 at the University of Florida Fort Lauderdale Research and Education Center, Davie FL. Means followed by a different letter are significantly different at 0.05 level. Sand:Peat Shoot dry weight (g) Root dry weight (g) Experiment 1 (August 2014)

100:0 8.0A 15.2AB 75:25 7.9A 18.4AB 50:50 7.8A 23.7A 25:75 7.8A 16.5AB 0:100 6.9B 8.2B

Experiment 2 (June 2015)

100:0 17.6A 18.2AB 75:25 17.1AB 18.2AB 50:50 17.4AB 21.4A 25:75 16.8BC 19.5AB 0:100 16.2C 17.3B

Table 4-2. Naiad (Najas guadalupensis) shoot and root dry weight of plants fertilized with 0, 1, 2, or 4 g CRF/kg substrate (Osmocote 15N-4.05P- 9.96K). Experiment 1 was conducted in August 2014 and experiment 2 was conducted in June 2015 at the University of Florida, Fort Lauderdale Research and Education Center. Means followed by different letters are significantly different at 0.05 level. Fertilizer (g/kg) Shoot dry weight (g) Root dry weight (g) Experiment 1 (August 2014)

0 17.1B 14.5B 1 18.2A 14.3B 2 18.4A 20.8A 4 17.2B 14.3B

Experiment 2 (June 2015)

0 16.6B 16.8B 1 18.1A 16.8B 2 17.5A 20.8A 4 15.9B 16.3B

Table 4-3. Shoot concentrations of ammonia (NH3-N), phosphate (PO3-P) and potassium (K) in naiad (Najas guadalupensis) shoots harvested from the experiment 2 conducted in June 2015. Plants were fertilized with 0, 1, 2, or 4 g CRF/ kg substrate (Osmocote 15N-4.05P- 9.96K). Means followed by different letters are significantly different at 0.05 level. Fertilizer (g/kg) NH3-N (mg/g) PO3-P (mg/g) K (mg/g) 0 2.05A 17.5B 58.6B 1 1.19A 35.8A 134.5A 2 1.22A 38.8A 247.6A 4 1.18A 18.5B 67.2B

Table 4-4. Water sample pH, electrical conductivity (EC), nitrate (NO3-N), phosphate (PO3-P) and potassium (K) of naiad (Najas guadalupensis) plants from the experiment 2 conducted in June 2015 and fertilized with 0, 1, 2, or 4 g per kg substrate (Osmocote 15N-4.05P- 9.96K). Means followed by different letters are significantly different at 0.05 level. Fertilizer pH EC (dS/m) NO3-N PO3-P K (mg/g) (g/kg) (mg/g) (mg/g) 0 8.6A 0.398A 0.353B 0.257B 0.90A 1 8.8A 0.357A 0.316B 0.293B 1.11A 2 8.6A 0.354A 0.394B 0.294B 0.84A 4 8.9A 0.359A 0.754A 0.406A 0.85A

Experiment 1 10 9 8 7 100 6 5 75 4 50 3 25 Shoot dryShootweight (g) 2 1 0 0 0 1 2 4 Fertilizer rate (g/kg)

Experiment 2 25

20 100 15 75 10 50 25 Shoot dryShootweight (g) 5 0 0 0 1 2 4 Fertilizer rate (g/kg)

Figure 4-1. Naiad (Najas guadalupensis) shoot dry weight of plants fertilized with 0, 1, 2, or 4 g/kg substrate (Osmocote 15N-4.05P- 9.96K) and grown in 100:0, 75:25, 50:50, 25:75, or 0:100 sand:peat substrate (by volume). Experiment 1 was conducted in August 2014 and experiment 2 was conducted in June 2015 at the University of Florida, Fort Lauderdale Research and Education Center. Bars represent standard error of the mean.

1 0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2

Electrical Electrical conductivity(dS/m) 0.1 0 100 75 50 25 0 Substrate (% Sand)

0 1 2 4

Figure 4-2. Final substrate electrical conductivity for plants fertilized with 0, 1, 2, or 4 g CRF/kg of substrate (Osmocote 15N-4.05P- 9.96K) and grown in 100:0, 75:25, 50:50, 25:75, or 0:100 sand:peat substrate (by volume). Experiment 2 was conducted in June 2015 at the University of Florida, Fort Lauderdale Research and Education Center. Bars represent standard error of the mean.

CHAPTER 5 CONCLUSIONS

My hypothesis was that naiad could be grown in containers using similar substrates and fertilizer rates as described in the published hydrilla studies. The first objective was to compare hydrilla and naiad growth in a sand substrate fertilized with 4 rates of a controlled release fertilizer. For the spring 2014 run of experiment 1, there was no difference in shoot or root dry weight between naiad and hydilla. Naiad and hydrilla shoot dry weights were both significantly greater in pots with 2 g CRF/kg sand, than with 0, 1, or 4 g CRF/kg sand. In the summer 2014 run of experiment 1, overall, hydrilla shoot dry weight was greater than naiad shoot dry weight. Barko (1982) found that there was a relationship in biomass growth of hydrilla to the differing fertilizer levels in the substrate with higher fertilizer rates resulting in more growth. Naiad shoot dry weight in pots with 1 g and 2 g CRF/kg sand was greater than 0 or 4 g CRF/kg sand.

Naiad root dry weight was greater in 2 g CRF/kg sand than 0 g CRF/kg sand, but not different from 1 g or 4 g CRF/kg sand. Hydrilla shoot dry weight was greater in 4 g

CRF/kg sand than 0 g, but not different from 1 g and 2 g CRF/kg sand. Hydrilla root dry weight was greater in 2 g and 4 g CRF/kg sand, than 0 g (Table 3-1). Naiad has adapted to local environments and seasonal variations in Florida (Main et al., 2006).

There are reported concentrations of naiad in the central and southern parts of Florida where soils are mostly sandy loam and low in nutrients (Novak et al, 2009). I suspect that this is why naiad performed better at lower fertilization rates compared to hydrilla.

The second objective was to compare naiad growth in 5 substrates and with 4 controlled release fertilizer rates. In both runs of experiment 2, shoot dry weight was greater in the 100:0 sand:peat substrates than in the 0:100 sand:peat substrates (Table

4-1). Furthermore, the shoot dry weights in both runs were greater with fertilizer rates of

1 and 2 g CRF/kg substrate than 0 and 4 g CRF/kg substrate. Root dry weight in both runs was greater in substrates with 2 g CRF/kg substrate than 0, 1, or 4 g CRF/kg substrate (Table 4-2). Bunt (1974) examined the physical and chemical characteristics of various peat:sand and peat:vermiculite substrate on the release of nutrients from controlled-release fertilizers and reported that leaching losses of N, P, and K were less in the peat:vermiculite substrate than in the peat:sand substrate. Electrical conductivity levels were highest in the 0:100 substrate with no difference among the other substrates. I suspect that the 0:100 substrate retained more fertilizer leading to higher soluble salt levels in the substrate and reduced naiad growth.

There is an increasing demand for native plants in our natural landscapes as well as interest in how to cultivate these type plants. It is my hope that this research will aid in providing evidence on naiad growth requirements for future use. Based on my experiments, I would recommend growing naiad in substrates containing 75 to 100% sand and fertilized with 1 to 2 g CRF/kg substrate.

LIST OF REFERENCE

Barko, J. W. 1982 Influence of potassium source (sediments vs. open water) and sediment composition on the growth and nutrition of a submersed freshwater macrophyte (Hydrilla verticillata (L.f.), Aquat. Bot. 12:157-172.

Barko, J. W., M. S. Adams, and N. S. Clesceri. 1986. Environmental factors and their consideration in the management of submersed aquatic vegetation: A review. J. Aquat Plant Manage 24:1–10.

Barko, J.W. and R.M. Smart. 1986. Sediment-related mechanisms of growth limitation in submersed macrophytes. Ecology 67:1328-1340.

Barnett, B.S. and R.W. Schneider. 1974. Fish populations in dense submersed plant communities. Hyacinth Control J 12:12-14.

Blackburn, R.D. and L.W. Weldon. 1962. Control of southern naiad (Najas guadalupensis) and other submersed weeds in south Florida irrigation and drainage channels. Proc. SWC. 15:254-255.

Buddington, R.K. 1979, Digestion of an aquatic macrophyte by Tilapia zillii (Gervais). J. Fish. Biol. 15:449-455.

Bunt, A.C., 1974 Some physical and chemical characteristics of loamless pot-plant substrates and their relation to plant growth. Acta Hort. 37:1954-1965.

Center T.D., J.H. Frank and F.A. Dray. 1997. Biological control. pp 245-266 In: D. Simberloff, D.C. Schmitz, and T.C. Brown, eds. Strangers in Paradise. Island Press, Washington, DC.

Dibble, E.D., K.J. Killgore, and S.L. Harrel. 1996. Assessment of fish-plant interactions. Amer. Fish. Soc. Symp. 16:357-372.

Fernald, M.L. 1923. Notes on the distribution of Najas in northeastern America. Rhodora 25:105-109.

Florida Exotic Plant and Pest Council (FLEPPC). 2015. Invasive Plant List. Definitions of Categories. Viewed June 1, 2015. http://www.fleppc.org/list/2015FLEPPCLIST-LARGEFORMAT-FINAL.pdf

Florida Fish and Wildlife Conservation Commission (FWC). 2015. Wetland Habitat Conservation in Florida. Florida Wetlands. Viewed June 1,2015. http://myfwc.com/conservation/freshwater/wetland-habitat/

Gettys, L.A., W.T. Haller, and M. Bellaud. (eds). 2009. Biology and Control of Aquatic Plants: A Best Management Practices Handbook. Aquatic Ecosystem Restoration Foundation, Marietta GA.

Gibbons, M., H. Gibbons, and M. Sytsma. 1994, A Citizens Manual for Developing Integrated Aquatic Vegetation Management Plans. Washington State Department of Ecology. Viewed June 1, 2015. https://fortress.wa.gov/ecy/publications/publications/93093.pdf

Godfrey, R.K., and J.W. Wooten. 1981. Aquatic and wetland plants of southeastern United States: dicotyledons (Vol. 2). University of Georgia Press, Athens, GA.

Gopal, B. 1987. Water hyacinth. Elsevier Science Publishers. Amsterdam, The Netherlands.

Gordon D.R. and K.P. Thomas, 1997. Impact and management of nonindigenous species in Florida. pp. 21-37 In: D. Simberloff, D.C. Schmitz, and T.C. Brown, eds. Strangers in Paradise. Island Press, Washington, DC

Gunnison, D. and J.W., Barko. 1989. The rhizoshpere ecology of submersed macrophytes. J. Amer. Water Res. Ass. 25 (1):193-201.

Haller, W.T., J.V. Shiremana, and D.F. Duranta. 1980. Fish harvest resulting from mechanical control of hydrilla. Trans. Amer. Fish. Soc. 109(5): 517-520.

Haller, W.T., J.L., Miller, and L.A., Garrard. 1976. Seasonal production and germination of hydrilla vegetative propagules. J. Aquat. Plant Manage. 14:26-29.

Harley, K.L.S. and I.W. Forno. 1992. Biological control of weeds: a handbook for practitioners and students. Inkata Press, Melbourne, Australia.

Joyce, J.C., K.A., Langeland, T.K., Van, and V.V. Vandiver. 1992. Organic sedimentation associated with hydrilla management. J. Aquat. Plant Manage. 30:20-23.

Kasselmann C. 1995. Aquarienpflanzen. Egen Ulmer GMBH & Co., Stuttgart.

Kay, S.H. 1992. Hydrilla: A rapidly spreading aquatic weed problem in North Carolina. Agriculture Bulletin AG-449. North Carolina Cooperative Extension Service, NC State University, Raleigh, NC., Viewed June 15, 2015. www.weedscience.ncsu.edu/aquaticweeds/hydrilla.PDF

Kiorboe, T. 1980. Distribution and production of submersed macrophytes in Tipper Grund (Ringkobing Fjord, Denmark), and the impact of waterfowl grazing. J. Applied Ecology 17:675-687.

Lee, M.L., W.R. West, P.A. Smith, and G.M. Booth. 1986. Determination and genotoxicity of nitrogen heterocycles in a sediment from the Black River. Env. Toxic. and Chem. 5(6):511-519.

MacIntyre, S. and J.M. Melack. 1995. Vertical and horizontal transport in lakes: linking littoral, benthic, and pelagic habitats. J North Amer. Benthol. Soc. 14:599-615.

Madeira, P.T., C.C., Jacono, and T.K., Van. 2000. Monitoring hydrilla using two RAPD procedures and the nonindigenous aquatic species database. J. Aquat. Plant Manage 38:33-40.

Madsen, J. D. 1998. Predicting invasion success of Eurasian watermilfoil. J. Aquat. Plant Manage. 36: 28-32.

Madsen, J.D., R.M., Stewart, K.D., Getsinger, R.L., Johnson, and R.M., Wersal, R.M., 2008. Aquatic plant communities in Waneta Lake and Lamoka Lake, New York. Northeastern Naturalist 15(1):97-110.

Main, M.B., G.M., Allen and K.A., Langeland. 2006. Creating Wildlife Habitat with Native Florida Freshwater Wetland Plants1. University of Florida, Cir, 912, pp.1- 11.

Missouri Department of Conservation, 2015. Naiads (Water Nymphs; Waterweeds); Viewed June 1, 2015. http://mdc.mo.gov/discover-nature/field-guide/naiads- water-nymphs-waterweeds

Mony, C., Koschnick, T.J., Haller, W.T. and Muller, S., 2007. Competition between two invasive Hydrocharitaceae (Hydrilla verticillata (Lf)(Royle) and Egeria densa (Planch) as influenced by sediment fertility and season. Aquatic Bot. 86(3):236- 242.

Morse, L.E., J.M., Swearingen, and J.M., Randall, J.M., 1999. Defining What is Native: What is a Native Plant. Roadside Use of Native Plants. Federal Highway Administration, Washington, DC, pp.12-14.

Neslon, P.V., 1998. Greenhouse operation and management. Prentice Hall, Upper Saddle River, New Jersey.

Netherland, M.D. and K.D. Jones. 2012. Registered herbicides and improving their efficacy on aquatic weeds. Aquatics 34 (3):12– 15.

Netherland, M.D. 2009. Chemical Control of Aquatic Weeds. pp. 65-77 In: Biology and Control of Aquatic Plants: A Best Management Handbook, L.A. Gettys, W.T. Haller, & M. Bellaud (eds.) Aquatic Ecosystem Restoration Foundation, Marietta, GA.

Novak, J.M., W.J. Busscher, D.L. Laird, M. Ahmedna, D.W. Watts, M.A.S. Niandu, 2009. Impact of Biochar Amendment on Fertility of a Southeastern Coastal Plain Soil. Soil Sci. 174(2):105-112.

Office of Technology Assessment [OTA]. 1993. Harmful non-Indigenous Species in the United States. Washington (DC): Office of Technology Assessment, US Congress

Ott R.A. Jr., C.C. Bonds, R.M. Smart, G.O. Dick, and L. Dodd. 2012. Propagation and Establishment of Native Aquatic Plants in Reservoirs. Texas Parks and Wildlife Department, Inland Fisheries Division.

Pennington, T. 2009. Egeria, pp. 129-133. In: Biology and Control of Aquatic Plants: A Best Management Handbook, L.A. Gettys, W.T. Haller, & M. Bellaud (eds.) Aquatic Ecosystem Restoration Foundation, Marietta, GA

Pimentel, D., R. Zuniga, and D. Morrison. 2005. Update on the environmental and economic costs associated with alien-invasive species in the United States. Ecological Econ 52(3): 273-288.

Ramey, V. 2001, Hydrilla verticillata, University of Florida Center for Aquatic and Invasive Plants, Viewed December 1, 2015. http://plants.ifas.ufl.edu/plant- directory/hydrilla-verticillata/#2

Schardt, J.D. and LE Nall. 1983. Aquatic Flora of Florida Survey Report, Florida Department of Environmental Regulation, Tallahassee, FL.

Schmitz, D. C., B.V. Nelson, L.E., Nall, and J.D. Schardt, 1991. Exotic aquatic plants in Florida: A historical perspective and review of the present aquatic plant regulation program. In: Proceedings of the Symposium on Exotic Pest Plants. US Natl. Park Serv. Tech. Rep. NPS/NREVER/NRTR-91/06 (pp. 303-323).

Sharp, M.J. and P.A., Keddy. 1985. Biomass accumulation by Rhexia virginica and Triadenum fraseri along two lakeshore gradients: a field experiment. Canadian J. Bo.t 63(10):1806-1810.

Smart, R. M., R. Doyle, J.D. Madsen, G.O. Dick. 1996. Establishing Native Submersed Aquatic Plant Communities in Southern Reservoirs. No. WES/TR/A-96-2. Army Engineer Waterways Experiment Station Vicksburg MS Environmental Lab. Viewed June 1, 2015. http://el.erdc.usace.army.mil/elpubs/pdf/tra96-2.pdf

Smith, C.S. and J.W. Barko. 1990. Ecology of Eurasian watermilfoil. J. Aquat. Plant Manage. 28(2):55-64.

Sondergaard, M. and K.A.J., Sand-Jensen. 1979. Distribution and quantitative development of aquatic macrophytes in relation to sediment characteristics in oligotrophic Lake Kalgaard, Denmark. J. Freshwater Bio. 9(1):1-11.

Southwest Florida Water Management District, Water (SWFWMD) Web Newsletter, 2013. Florida's Wetlands. pg. 2, Viewed June 1, 2015. http://www.swfwmd.state.fl.us/publications/files/waterweb_wetlands.pdf

Spencer, D.F., G.G. Ksander, and S.R., Bissell. 1992. Growth of monoecious hydrilla on different soils amended with peat or barley straw. Soil and Water 10:73-78.

Steward, K.K., 1984. Growth of hydrilla (Hydrilla verticillata) in hydrosoils of different composition. Weed Sci. 32: 371-375.

Sutton, D.L. and K.H. Portier. 1983. Variation of nitrogen, phosphorus, and potassium contents of Hydrilla in south Florida. J. of Aquat. Plant Manage.21(2):87-92.

Sutton, D.L. 1985. Culture of hydrilla (Hydrilla verticillata) in sand root media amended with three fertilizers. Weed Sci. 32:371-375.

Sutton, D.L. 1990. Comparison of two methods of Evaluating Growth of Hydrilla in Sediments Collected from Lake Okeechobee. J. Aquat. Plant Manage 28:80-83

Tisdale, S.L., Nelson, W.L. and Beaton, J.D., 1985. Soil fertility and fertilizers. Macmillan Ltd Publishers, New York, NY.

Texas A & M Agrilife Extension (TAMU) Department of Wildlife and Fisheries Sciences, Service. 2015. Description of Southern Naiad. Viewed June 15, 2015. http://aquaplant.tamu.edu/plant-identification/alphabetical-index/southern-naiad/>

Trophic 2002, Florida Lakewatch, November, PM6.5, Viewed June 15, 2015. http://lakewatch.ifas.ufl.edu/circpdffolder/trophic2.pdf

United States Department of Agriculture (USDA). 2015a. National Invasive species information Center. Viewed June 15, 2015. http://www.invasivespeciesinfo.gov/whatis.shtml

United States Department of Agriculture (USDA). 2015b. Plant profile for Egeria. viewed June 15, 2015. http://plants.usda.gov/core/profile?symbol=EGDE

University of Florida Institute of Food and Agriculture Science Center for Aquatic and Invasive Plants (UF/IFAS CAIP). 2015a. “Florida Waters: Ours to Protect”. Viewed June 15, 2015. http://plants.ifas.ufl.edu/wp- content/uploads/files/caip/pdfs/Florida_Waters_Ours_to_Protect.pdf

University of Florida Institute of Food and Agriculture Science, Center for Aquatic and Invasive Plants (UF/IFAS CAIP). 2015b. Southern Naiad. Viewed June 15, 2015. http://plants.ifas.ufl.edu/node/183#1

Warcke, D D. and D. M. Krauskopf. 1983. Greenhouse Growth Media: Testing & Nutrition Guidelines. Extension Bulletin E-1736. Michigan State University. Department of Crop & Soil Sciences.

Wittenberg, R., and M.J.W. Cock (eds.) 2001. Invasive Alien Species: A Toolkit of Best Prevention and Management Practices. CAB International, Wallingford, Oxon, UK, xvii - 228.

BIOGRAPHICAL SKETCH

Heather Hasandras is a student of environmental horticulture working on her

Master of Science degree at the University of Florida. She is originally a New York City kid who moved to Florida after graduation from Fordham University. She developed a love for horticulture through bringing her daughter to various 4H events in Florida. This led to her being trained by Dr. Mike Ofanedes at the Broward County Cooperative

Extension Service through a 4H demonstration garden for the public. Heather has continued to assist at University of Florida Fort Lauderdale Research and Education

Center events like the open house and trial garden. She is certified through the agriculture in the classroom program, which is a Florida program that aims at bringing agriculture back into the classroom through incorporating science and math curriculum.

She has taught multiple summer, fall, and spring workshops in farm to fork type curriculum, as well as the Smithsonian institute derived agricultural world perspective curriculum for children through her non-profit. She continues to serve as an appointed

Landscape Advisory board member for the city of Pembroke Pines where she offers landscaping guidance as far as the use of the right plant, in the right place, and Florida friendly initiatives to help the community. She is a member of the National Wildlife

Foundation, and the American Society for Horticulture Science. She continues her work with horticulture industry professionals, Florida Farm Bureau, and Broward schools to reach the next generation of children to educate students on the importance of agriculture to our communities and world environment.