Mitigating Cyanobacterial Harmful Algae Blooms: the Role of Plant Humics

AN ABSTRACT OF THE THESIS OF

Susan A.F. Theis for the degree of Master of Science in Botany and Plant Pathology presented on December 8, 2014.

Title: Mitigating Cyanobacterial Harmful Algae Blooms: The Role of Plant Humics

Abstract approved:

______Allen J Milligan

Abstract: Cyanobacterial harmful algae blooms (cyanoHABs) are a growing concern worldwide due to damage of ecosystems and threats to human health. Previous research indicates that plant humics from aquatic and wetland vascular plants are effective inhibitors of cyanobacterial metabolism and growth and may be useful as control agents for mitigating cyanoHABs. Using lake-side incubations, we exposed natural phytoplankton assemblages to plant humics. Three plant species, Hordeum vulgare

(barley straw), Typha latifolia (cattail), and the submerged, aquatic invasive

Myriophyllum spicatum (Eurasian watermilfoil) were tested for inhibitory effects against cyanobacteria in mesocosm experiments. Trials were conducted on a eutrophied

Northern Michigan lake against five cyanobacterial genera (Microcystis, Anabaena,

Lyngbya, Oscillatoria, and Arthrospira) and one Chlorophyta (Ulothrix). Lakeside mesocosm experiments were conducted in 3.4L jars containing lake water at continuous oxygen saturation. Treatment effectiveness was assessed via microscopic analysis of cyanobacterial and Chlorophyte biomass. Total phosphorus, soluble reactive phosphorus, and total nitrogen changes were analyzed with each treatment. All treatments significantly inhibited Microcystis growth rates. Anabaena was significantly inhibited by

M. spicatum at all doses. Lyngbya was significantly inhibited in over 65% of all treatments. Oscillatoria growth rates were unaffected in all but two treatments, and

Ulothrix was unaffected in all but one treatment. Results indicate that plant humic substances are effective in suppressing cyanobacterial growth and that local aquatic plants introduce less phosphorus while exhibiting similar inhibition rates as barley straw.

This work suggest that fringing wetlands and submerged aquatic plants may play an important role in regulating harmful cyanobacterial blooms by providing inhibitory humics and that this role should be considered when determining wetland value.

Mitigating Cyanobacterial Harmful Algae Blooms: The Role of Plant Humics

by Susan A.F. Theis

A THESIS

submitted to

Oregon State University

in partial fulfillment of the requirement for the degree of

Master of Science

Presented December 8, 2014 Commencement June 2015

Master of Science thesis of Susan A.F. Theis presented on December 8, 2014.

APPROVED:

______Major Professor, representing Botany and Plant Pathology

______Head of the Department of Botany and Plant Pathology

______Dean of the Graduate School

I understand that my thesis will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my thesis to any reader upon request.

______Susan A.F. Theis, Author

TABLE OF CONTENTS

Page

1. Mitigating Cyanobacterial Harmful Algae Blooms ...... 1

1.1 Introduction ...... 3

1.2 Study Site ...... 8

1.3 Materials and Methods ...... 8

1.3.1 Plant material ...... 9

1.3.2 Mesocosms ...... 9

1.3.3 Algal composition ...... 10

1.3.4 Time and Dosage ...... 10

1.3.5 Data Analysis ...... 11

1.3.6 Statistics ...... 12

1.4 Results ...... 14

1.4.1 Hordeum vulgare ...... 14

1.4.2 Myriophyllum spicatum ...... 15

1.4.3 Typha latifolia ...... 16

1.4.4 Community Results ...... 18

1.5 Discussion ...... 21

1.5.1 Mechanisms ...... 21

1.5.1.1 Reactive Oxygen Species ...... 21

1.5.1.2 HS Components and direct toxicity ...... 23

TABLE OF CONTENTS (Continued)

Page 1.5.2 Mesocosm Conditions ...... 24

1.5.2.1 Light Attenuation ...... 25

1.5.2.2 Microbial Role ...... 26

1.5.3 Experimental Effectiveness ...... 26

1.5.3.1 Nutrient Dynamics ...... 27

1.5.3.2 Hordeum vulgare ...... 28

1.5.3.3 Myriophyllum spicatum ...... 29

1.5.3.4 Typha latifolia ...... 30

1.5.4 General Phytoplankton Trends ...... 33

1.5.5 Management Implications ...... …35

1.5.5.1 Wetland Valuation ...... 37

1.5.5.2 Active Management Strategies ...... 39

1.6 Conclusion ...... 40

Bibliography ...... 65

LIST OF FIGURES

Figure Page

1. H. vulgare (barley straw) treatment effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira ……………………...41

2. Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Microcystis...... ……………………….………………....42

3. Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Lyngbya……………………………………………………………………...42

4. Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Oscillatoria………………………………………………………………….43

5. Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Arthrospira…………………………………………………………..43

6. Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira………………….44

7. H. vulgare (barley straw) treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen………………………………………..45

8. Myriophyllum spicatum treatment effects on the growth rates of Microcystis, Anabaena, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira…………46

9. Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Microcystis ………………………………………………...... 47

10. Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Ulothrix……………………………………………………………………..47

11. Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Lyngbya……………………………………………………………..48

12. Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Arthrospira………………………………………………………….48

13. Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira………………..49

LIST OF FIGURES (Continued)

Page 14. M. spicatum treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen ………………………………………………..50

15. Typha latifolia (cattail) green shoot treatments effect on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira….………..51

16. T. latifolia green shoot treatment impacts on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen…..……………..………..52

17. Typha latifolia (cattail) brown shoot treatment effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira….…...... 53

18. Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Microcystis ……………..…………………………...... 54

19. Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Lyngbya…………………………………………………54

20. Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira………………………………………………………………………...55

21. T. latifolia brown shoot treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen…….…………….…………………...56

22. Typha latifolia (cattail) root treatments effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira………….……..57

23. T. latifolia root treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen………………………………………………..58

24. Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Microcystis…………………………………………...... 59

25. Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Lyngbya………………………………………………60

26. Increases in total phosphorus (µg/L) from controls in treatments inhibiting Oscillatoria………………………………..……………………….61

LIST OF FIGURES (Continued)

Page

27. Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Arthrospira………………….………………………...62

LIST OF TABLES

Table Page

1. Dates, temperature, pH, and dose levels.………………..………………………..50

2. Tukey’s HSD p-values of experimental treatments...………………………….....51

Mitigating Cyanobacterial Harmful Algae Blooms: The Role of Plant Humics

Susan Theis¹, Allen Milligan²*

¹, ² Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR

97331

*Corresponding author email: [email protected] Abbreviated title: Mitigating Harmful Cyanobacterial Blooms: Plant Humics

(barley straw), Typha latifolia (cattail), and the submerged, aquatic invasive

Myriophyllum spicatum (Eurasian watermilfoil) were tested for inhibitory effects against cyanobacteria in mesocosm experiments. Trials were conducted on a eutrophied

Northern Michigan lake against five cyanobacterial genera (Microcystis, Anabaena,

M. spicatum at all doses. Lyngbya was significantly inhibited in over 65% of all

treatments. Oscillatoria growth rates were unaffected in all but two treatments, and

Key words: cyanoHAB, mitigation, cyanobacteria, Microcystis, Anabaena, Lyngbya, Oscillatoria, barley, Typha, Myriophyllum, wetlands, humics, peroxide

Introduction

Cyanobacterial Harmful Algal Blooms (cyanoHABs) are unresolved world-wide problems with negative effects that cover the range from individuals to ecosystems.

They inflict damages upon ecosystem health, water quality, and recreational water use through unpleasant scums, the creation of anoxic aquatic zones (creating toxic ammonia concentrations and fish kills), trophic web alterations, and positive feedback loops of nutrient loading (Paerl et al 2001; Bury 1995). In addition, many cyanobacteria create toxins that cause liver, neural, digestive, and kidney damage, tumor promotion, contact dermatitis, and even death in humans and wildlife (Paerl and Huisman 2009; Paerl and

Otten 2013).

Toxic cyanobacteria poisonings have been reported in animals such as birds, cattle, and sheep (Carmichael 2001) and have caused over 350 cases of suspected or confirmed poisonings or deaths in the U.S. between the 1920s and 2012 (Backer et al

2013). Bioaccumulations of toxins have been observed in Daphnia, bi-valves, and crayfish. Toxins have also been found to negatively impact multiple species of fish, causing liver, heart, kidney, and gill damage (Zurawell et al 2005).

Detrimental effects upon water supplies – for drinking, recreation, and irrigation – are reported with increasing frequency. According to the EPA’s 2011 National Lakes

Assessment, 27% of U.S. lakes (using WHO thresholds) offer a moderate to high risk of algal toxin exposure (Holdsworth 2011). In the summer of 2014, a state of emergency was declared in Toledo, Ohio, due to cyanobacterial toxins (microcystins) found in the

city’s drinking water supply. Toxins also pose a risk for agricultural water, as they have been shown to accumulate in plants and reduce plant yield (Codd et al 1999; McElhiney et al 2001; Dao et al 2014).

CyanoHABs originate with contributions from a variety of factors, primarily increased anthropogenic phosphorus inputs, eco-system disturbances, and trophic imbalances (Perovich et al 2008; Paerl and Otten 2013; Schrage and Downing, 2004;

Benndorf et al 2002). Climate change may exacerbate bloom frequency due to increased water temperatures and lake stratification (Paerl et al 2011; Posch et al 2012). Increases in water temperature by 1.8 – 3.3°C have been shown to double and triple Microcystis populations (Jöhnk et al 2008; Beaulieu et al 2013). Increased water column stability would limit mixing and give both filamentous and planktonic cyanobacteria a competitive advantage (Taranu et al 2012).

A primary objective for abatement of cyanoHABs is a reduction in nutrient loading (Piehler 2008; Paerl and Otten 2013; Jeppesen 2007). Other management techniques commonly include mechanical methods such as vertical mixing, oxygenation, and dredging as well as chemical methods. Copper has been used as an algaecide, for example (Lehman et al 2004), and alum is often added to eutrophic lakes to remove available phosphorous by forming insoluble aluminophosphates (Galvez-Cloutier et al

2012).

A large number of lakes that are afflicted by cyanoHABs are reservoirs with minimal plant growth and lakes that have lost substantial wetland and riparian plant

contact. In addition to plants’ roles in reducing nutrient loading, they are also a primary source of humic substances (HS). Various plant humic substances (HS, macromolecules including phenols, catechols, and quinones) have been shown to selectively suppress cyanobacterial growth (Prokhotskaya and Steinberg 2007; Bährs et al 2013; Gross et al

2006).

Humics from barley straw (Hordeum vulgare) have been shown to be an effective method of inhibiting various cyanobacteria (Ó hUallacháin and Fenton 2010; Everall and

Lees 1996; Iredale et al 2012). Native (and nuisance) plants can also supply humics to aquatic ecosystems (Steinberg et al 2006).

Typha spp. (cattail) are ubiquitous wetland plants and have been used for constructed wetlands and floating islands (Nakai et al 2008, 2010; Hubbard et al 2004).

To date, one study has examined the impact of Typha root exudates on cyanobacteria

(Nakai et al 2010) and found that M. aeruginosa growth was inhibited. Successful inhibition of M. aeruginosa has been demonstrated by application of T. orientalis (Chen et al 2012) and inhibition of Aphanizomenon flos-aqaue blooms by a mixture of Scirpus and Typha latifolia (Haggard et al 2013).

Myriophyllum spicatum (Eurasian Watermilfoil) is a common nuisance/invasive submerged aquatic, the irradication of which receives considerable attention from aquatic managers (Newman 2004). Tellimagrandin II (a hydrolysable polyphenol isolated from

M.spicatum) has been shown to inhibit cyanobacteria (Steinberg et al 2006). Studies using purified Tellimagrandin II found inhibition of both extracellular enzymes (e.g,

alkaline phosphatases) in Trichormus (Gross et al. 1996) and photosynthetic oxygen evolution in Anabaena PCC 7120. Inhibition of photosystem II (PS II) was confirmed in eukaryotes (Leu et al 2002).

There are likely multiple mechanisms by which HS inhibit cyanobacteria

(Steinberg et al 2006). One important mechanism is thought to be through photooxidation of colored HS (Waybright et al 2009, Prokhotskaya and Steinberg 2007). Photooxidation of HS has been shown to create reactive oxygen species (ROS) such as hydrogen peroxide, superoxide, and hydroxyl radicals (Scully et al 2006; Iredale et al 2012; Cooper and Lean 1989; Farjalla et al 2001). A growing body of research indicates that successful cyanobacterial inhibition by plant products depends on adequate UV exposure (Iredale et al 2012; Haggard et al 2013; Lesser et al 2008), moderate to high levels of dissolved oxygen (DO) (Lindell et al 2000; Appel 1993; Waybright et al 2009), and, in some cases, partial microbial decomposition of plant materials (Iredale et al 2012; Ball et al 2001;

Murray et al 2010). Inhibition of cyanobacteria by barley is usually unsuccessful when applications are attempted in low DO conditions (Boylan and Morris 2003 and Rajabi et al 2010) or experiments limit exposure time and surface area, precluding decomposition

(Kelly and Smith 1996; Boylan and Morris 2003; Rajabi et al, 2010). Exposure time, pH

(Bährs et al 2013) and temperature (Iredale et al 2012; Scully et al 2006) also affect the degree to which growth rates are inhibited.

7 Many lake and pond scale studies that have tested the impact of plant derived HS have shown a range of impacts from no inhibition to enhancement of cyanobacterial growth rates (Everall and Lees 1997; Barrett et al 1999; Welch et al 1990). Often these

disparate results cannot be evaluated due to lack of proper controls and limited assessments of environmental variables.

This study examines the effects of barley straw and two local plant species

(Hordeum vulgare, Typha latifolia, and Myriophyllum spicatum) on the growth rates of native phytoplankton populations via lake-side mesocosm experiments. Our mesocosms were designed to compare the well-known model source of HS, barley straw, with species found in situ. Emphasis is placed on inhibiting cyanobacteria while both limiting phosphorus additions and shifting community composition towards native microalgal species that promote aquatic ecosystem health. Mesocosm analyses focus on a single lake as a model, but results may be applicable for current and future restoration, protection, and adaptive management strategies.

Study site:

Viking Lake is a small (~62 acres), shallow (Zmax=35’), dimictic lake in the Au

Sable River headwaters area of northern Michigan (44º53’43”N 84º37’12”W) at an elevation of 376.7 m (1236 ft). The lake perimeter is minimally developed (16 part time residents), has no surface water inputs, and is free of Zebra mussels (Dreissena polymorpha). While the lake was historically oligotrophic with bog characteristics, recreational management techniques (liming and fish stocking) led to dense harmful cyanobacterial blooms (HCBs) with epilimnetic pH values exceeding 10.5. A complete lake profile is available upon request. During summer months, Microcystis aeruginosa and Anabaena sp. dominate the water column while large populations of Oscillatoria and

Lyngbya spp. form extensive, semi-benthic mats. Microcystin production has been confirmed in this lake system.

Materials and methods:

Mesocosms assays were conducted from July 30th to September 16th, 2013. Five experiments were conducted; each was temporally separated and conducted in three- to five-day increments. Each experiment was composed of three distinct treatments and a control (in quadruplicate) in randomized block design. All plant treatments were dried, cut into 1-2” pieces, and applied directly to mesocosm water.

Plant material: Typha latifolia (cattail) fresh leaves/shoots (referred to as ‘green shoots’) and the previous growing season’s partially decomposed leaf litter (‘brown shoots’) were harvested from the perimeter of Viking Lake and dried for 7 days in the sun. T. latifolia material was harvested in early July for experiments 1, early August for experiment 3, and in early September for experiment 5. All materials were weighed to

10mg accuracy for dry weight (DW) or wet weight (WW). T. latifolia roots and rhizomes were unearthed, thoroughly rinsed, and cut while wet. Myriophyllum spicatum

(Eurasian Watermilfoil) was harvested from Cedarville Bay in Lake Huron, MI on July

20. Hordeum vulgare (barley straw) was obtained from Oregon State University Hyslop research farm in Corvallis, OR, and was not fertilized during its cultivation. This variety of barley may have high levels of antimicrobial compounds due to its cultivation in an area subject to high disease pressure (Castro and Fontes 2005), thus increasing its efficacy in cyanobacterial inhibition.

Mesocosms: Mesocosm water was incubated in individual 3.4 L glass containers inside of a large-volume, circular tub placed near the lake. Incubation tub temperature was designed to loosely match lake temperature fluctuations by periodic manual adjustments. Dissolved oxygen concentrations were maintained at 100% saturation using micro-bubble air diffusers powered by aquarium pumps. Air bubblers were weighted to ensure constant contact with the bottom of mesocosm jars to provide continual mixing.

In addition, jars were capped and inverted once daily.

Effects of zooplankton grazing were precluded by the application 0.33 mL/L of the insecticide Sevin (Relyea 2006); effects were confirmed microscopically. These concentrations have been shown effective against cladocerans, yet are well below lowest observed effect concentrations (LOEC) observed against M. aeruginosa (Ma et al 2006).

Algal composition: Cyanobacterial biovolume and growth rates were monitored for Microcystis (M. aeruginosa and Microcystis sp.), Anabaena sp., Oscillatoria,

Lyngbya, and Arthrospyra sp. as well as for the Chlorophyta Ulothrix sp. A more extensive list of community assemblage is available.

Unusually high water levels and low summer temperatures resulted in a less severe summer bloom, and thus mesocosms were supplemented with additional algal biomass and detritus harvested from the lake. Floating masses of algae and particulate detritus were collected using 153 μm plankton netting; 30 ml of algal mat material was added to pelagic lake water and mixed before even distribution into experimental vessels.

Time and Dosage: Treatments were conducted in 3-5 day segments from July

30th – September 16th, 2013. Based upon both previous research and feasibility of potential in-situ application, treatment doses ranged from 1-7 g/L DW (Table 1).

Temperature, pH, oxidation reduction potential (ORP), and dissolved oxygen were measured daily with a Hydrolab Surveyor (Hach Environmental, Loveland, CO). ORP measurements were converted to Eh (mV) by the adjustment of +248. Biomass samples were taken daily, preserved using a Lugol’s iodine solution (1% final volume), and stored

in the dark until analyzed. Nutrient samples were stored in polypropylene bottles that were immediately wrapped in foil and frozen until analyzed.

Data analysis: Utermöhl settling chambers were used for microscopic enumeration and biovolume estimates. Water samples (10 ml) were settled for approximately 8 hours, (condensed to 2 ml) and viewed using an inverted microscope

(Axiovert S100) fitted with a digital camera. Specimens were counted using 4 photographs per slide (or more, until 200 filaments were counted) of randomly selected fields of view. The optimal number of filaments to count was determined by evaluating the 95% confidence interval (CI) of means that would yield a CI of less than 20%.

Lengths of filaments and area outlines of colonies were manually traced using calibrated

ImagePro software. Length measurements and software calculated areas were then exported to Microsoft Excel, and picture totals were averaged for each slide. Biovolume was calculated as follows for filaments:

where Ltot is the sum of all lengths measured (averaged over ≥ 4 pictures), WA is the filament cross-section area calculated from the measured width (WA = Wπr ²), and V is the mL of sample processed. After colonial area was measured, colonial volume was calculated as follows:

where Atot = the sum of all areas measured (averaged over ≥ 4 pictures), SV = the spherical volume calculated from the measured from total circular colony area, and V is the mL of sample processed. Growth rates (µ/d) were determined from the equation:

Nutrient analyses were conducted by spectrophotometry in 2ml cuvettes. Soluble reactive phosphorous (SRP) was determined by the ascorbic acid colorimetric method

(Murphy and Riley 1962). Total phosphorus (TP) samples were first oxidized using the persulfate method (Valderrama 1981) and analyzed for SRP. Nitrate analysis was conducted with spongy cadmium as described by Jones (1984). We found that increasing the reaction time with cadmium to 150 min improved replication between samples (data not shown).

Statistics: 95% confidence intervals were calculated for all data shown. Growth rate variance was analyzed across treatments with ANOVA and post-hoc Tukey’s HSD tests (α = 0.05). In some cases, assumptions of equal variance were not met (denoted by italics in Table 2), in which case Holm Bonferroni sequentially adjusted Welch’s t-tests were conducted in place of Tukey’s HSD (Holm 1979). Growth rate statistical analyses

were conducted using R (Rx64 3.1.2) software (R Core Team 2014). Regression analyses were conducted by ANOVA using Microsoft Excel.

Results

The inhibition of cyanobacterial growth rates was significant (p<0.05) in 57% of all treatments (Table 2). Microcystis was significantly inhibited by all but one treatment.

Anabaena was present only in experiment 4, where it was entirely inhibited. Lyngbya was significantly inhibited in 85% of barley, M. spicatum, and Typha green and brown shoot treatments, but was unaffected by Typha root treatments. Oscillatoria was largely unaffected by plant treatments, but was significantly inhibited in experiment 1 by both brown shoots and roots. Individual treatment results are listed below by plant category.

Hordeum vulgare – Both 3 and 7 g/L barley straw treatments significantly inhibited

Microcystis growth rates; applications of 7g/L were nearly twice as effective as 3 and 5 g/L treatments (Fig 1). Due to low data resolution, the 5 g/L barley dose was the only treatment out of all five experiments that did not significantly inhibit Microcystis.

Increases in barley straw doses (g/L) were linearly and negatively correlated with

Microcystis growth rates (Fig. 2). Barley treatments did not significantly inhibit Ulothrix or Oscillatoria. Lyngbya and Arthrospira growth rates were incrementally and significantly inhibited at all dose levels. Linear correlations between barley treatment doses and Lyngbya (Fig. 3), Oscillatoria (Fig. 4), Arthrospira (Fig. 5), and all four cyanobacteria (Fig. 6) also show negative, linear correlations.

Phosphorus loadings exhibited in barley treatments increased from controls by 1-

2.6 µg/L per DW gram of straw added (3-18.4 µg/L per dose), with 7 g/L treatments increasing TP 7 times that of the control (Fig. 7). Soluble reactive phosphorus constituted, on average, 83.7% of TP. Total nitrogen levels decreased in 3 and 5 g/L additions by 0.40 - 0.88 mg/L, but increased by 0.25 mg/L with the 7 g/L treatment.

Myriophyllum spicatum – All M. spicatum treatments significantly reduced Microcystis growth rates (Fig. 8). As increases in treatment dosage beyond 12 g/L WW did not increase Microcystis inhibition, a linear dose-response correlation explains just over a third of the observed effect (Fig. 9). Dose-response correlations between M. spicatum treatments and Ulothrix (Fig. 10), Lyngbya (Fig. 11), Arthrospira (Fig. 12), and all four cyanobacterial growth rates (Fig. 13) were negative and significant.

As Anabaena was not observed in any of the treatment analyses, growth rates were reduced to zero in all Myriophyllum treatments. Ulothrix was unaffected at the lowest dose of 12 g/L WW and significantly reduced at the highest dose of 24 g/L, but high variability in the mid-level dose of 18 g/L precluded a clear resolution of effect (p =

0.083). Lyngbya growth rates were significantly inhibited with 12 g/L and 24 g/L WW doses, but variability in growth rates at the mid-level dose made resolving an effect impossible (p=0.08). Oscillatoria was unaffected by all treatments. Arthrospira was unaffected at low and mid doses but was significantly inhibited at 7 g/L.

Total phosphorus increased from the controls with the addition of M. spicatum by

1-2.3 µg/L per DW gram (1.4-6.3 µg/L per dose) (Fig. 14). The largest M. spicatum dose increased TP 3 times that of the control and added half as much TP as the highest barley treatment. On average, SRP constituted 90.2% of TP additions. TN increased slightly by

0.05-0.20 mg/L per DW dose across all M. spicatum treatments.

Typha latifolia -

Green shoots: Typha green shoot (2.5 g/L DW) treatments significantly inhibited

Microcystis, Lyngbya, and Arthrospira growth rates (Fig. 15). Ulothrix and Oscillatoria were not affected.

Relative to the control, Typha green shoot TP additions increased by 2.9 µg/L per

DW gram of shoot added (7.3 µg/L per dose) (Fig. 16). SRP was 76.8% of TP, and total nitrogen increased by 0.15 mg/L per gram of green shoot added.

Brown shoots: All Typha brown shoot treatments significantly inhibited Microcystis over three independent experiments (Fig. 17). Dose-response correlations were negative and significant between experiment 5 and Microcystis growth rates (Fig. 18). Dose- response correlations between brown shoot treatments and Lyngbya (Fig. 19) and all four cyanobacterial growth rates (Fig. 20) were negative and significant.

Ulothrix was not affected by brown shoot treatments. Lyngbya growth rates were significantly inhibited by all treatments except the lowest treatment dose of 1 g/L in experiment 5.

Oscillatoria growth rates were inhibited by exposure to 2.5 g/L brown shoots in experiment 1 that were collected in early July, but were not significantly affected by shoots collected in early August and September for experiments 3 and 5. Similarly,

Arthrospira was significantly inhibited by brown shoots collected for experiment 1, but was unaffected by shoots in experiment 3 and 5. High variability of Arthrospira inhibition in experiment 3 (Fig 17) was due to an outlier showing near complete inhibition; this was not removed for data analysis. In addition, the presence and microscopic observation of Cladosporium fungal hyphae contributed to diminished data resolution.

Total phosphorus loadings from Typha brown shoots varied according to the time of year they were collected, ranging from 0 – 0.5 µg/L per gram DW of brown shoot added (0 – 1.26 µg/L per dose) (Fig. 21), with an average of 57.8% of TP in the form of

SRP. Total nitrogen increased from the control by 0.068 mg/L in experiment 1, was unaffected in experiment 3, and decreased by 0.43-1.19 mg/L per DW gram of brown shoot added in experiment 5.

Typha Roots: Microcystis growth rates declined significantly with both 10 g/L and 16.7 g/L treatments (Fig. 22). Both Ulothrix and Lyngbya growth rates were not significantly affected by addition of Typha roots. Oscillatoria was significantly inhibited

18 by 16.7 g/L Typha root treatments, while 10 g/L treatments did not show statistically significance (p = 0.069). Arthrospira growth rates were significantly inhibited by larger doses and stimulated by smaller doses.

The combined Typha root and shoot treatment significantly decreased

Microcystis, and did not exhibit a synergistic effect, as other genera were largely unaffected (data not shown).

Total phosphorus loadings from Typha root additions ranged from (0.09-0.19

µg/L per gram WW of root (1.6 - 1.9 µg/L per dose), with an average of 52.2% of TP in the form of SRP (Fig. 23). Total nitrogen in the 16.7 g/L treatment was 0.5 mg/L higher than the control, but 0.14 mg/L lower than Day 0.

Community Results

Despite an overall decline of total biovolume in every experiment, two treatments showed a significant stimulation of two non-cyanoHAB genera. M. spicatum (24 g/L

WW) significantly increased the growth rate of Ulothrix, and T. latifolia root treatments

(10 g/L) significantly increased Arthrospira growth rates. However, the majority of treatments had little effect on Ulothrix.

Decreases in average community growth rates were observed in all treatments across all experiments, despite increased available phosphorus in all treatments. Total phosphorus additions in M. spicatum, T. latifolia, and H. vulgare treatments ranged from

19 0 – 2.9 µg/L per gram of plant addition (0 – 18.4 µg/L per treatment), with an average of

73% of TP in form of SRP.

While the amount of total phosphorus addition per gram of plant added is somewhat comparable across plants, TP additions vary substantially when considering the amount of plant necessary to elicit a significant inhibitory response. For example, a selected range of treatments that significantly inhibit Microcystis (Fig. 24) show similar effectiveness (90-97% biovolume reduction), yet barley treatments add 9-20 times as much total phosphorus as Typha and M. spicatum treatments. While variations in TP additions from statistically significant treatments inhibiting Lyngbya (Fig. 25) are not as obvious as in Microcystis treatments, Typha and M. spicatum offer greater levels of inhibition per gram of phosphorus added to the system. Typha root and brown shoot treatments reduced Oscillatoria biovolume by 92% (Fig. 26), while 7 g/L barley straw treatments introduced 10 times the phosphorus with only 62% biovolume reduction.

Variations in Arthrospyra biovolume inhibition versus total phosphorus additions indicate that M. spicatum offers the greatest inhibition per gram of total phosphorus addition (Fig 27).

In all experiments, controls exhibited positive growth rates when averaged across all species considered (average 0.21 µ/d, range 0.7-0.36 µ/d). All controls exhibited an increase in both Microcystis and Ulothrix biovolume; however, control growth rates of

Oscillatoria, Lyngbya, and Arthrospira varied between experiments. These three genera experienced either a stasis or negative growth rates in almost all Typha experiments and a stasis or positive growth rates in experiment 2 (barley straw). In experiment 4 (M.

20 spicatum), Arthrospira and Lyngbya growth rates were positive, while Oscillatoria control growth rates were negative.

Controls and low dose treatments were observed to have relatively large clumps of particulate debris intertwined with Oscillatoria and Lyngbya filaments, causing issues in data resolution. An elimination of particulate debris masses was consistently observed with increased treatment dose, indicating an underestimation of treatment effectiveness.

Discussion

Mesocosm application of plant derived HS significantly inhibited the growth rates of several prominent cyanoHAB genera across five independent experiments.

Discussed below are two primary mechanisms of HS mediated cyanobacterial inhibition, environmental conditions affecting HS toxicity and cyanobacterial susceptibility, potential reasons for inter- and intra-study similarities and variability, and management implications applicable for a wide variety of aquatic systems.

Mechanisms

Plants contain a diverse array of humic substances – primarily phenols – that are exuded during growth or released during the decomposition of lignin. Humic substance inhibition of cyanobacteria occurs through two general mechanisms: 1) via the photochemical production of reactive oxygen species (ROS) and 2) via the toxicity of plant-specific compounds.

Reactive Oxygen Species: ROS can be extremely toxic to cyanobacteria, in part because it may interfere with gram negative bacterial extracellular enzymes and with photosynthetic electron transport chains (Steinberg et al 2006) by directly quenching electrons (Pflugmacher et al 2006). Application of plant HS have shown to impact cyanobacteria through increased extra- and intra-cellular ROS in conjunction with

22 decreased electron transport rates, decreased ATP production, and cell lysis (Shao et al

2010).

Cyanobacteria have been shown to be particularly sensitive to extracellular peroxide, most likely due to their simplified cellular structure (Prokhotskaya and Steinberg 2007).

For photochemical processes to be an effective driver of plant derived HS toxicity on cyanobacteria, two environmental conditions must be met: 1) Oxygen levels must be adequate to accept electrons from photoactivated organic compounds and 2) light of appropriate wavelength (300 – 500 nm) must penetrate into the water column. This exposure is necessary for chemical excitation as described in the equations below; proper light exposure has been demonstrated to be a limiting factor cyanobacterial inhibition

(Iredale et al 2012). For example, plant treatments with high UV exposure exhibited inhibition rates of A. flos-aquae that were three times stronger than low UV treatments

(Haggard et al 2013).

Linear, predictive relationships have been demonstrated between colored HS and

ROS (Scully et al 1996; Shao et al 1994; Cooper et al 1988); however, production seems to vary by plant genera (Farjalla et al 2001). Photooxidation of colored, organic HS is a complex series of reactions that produce reactive oxygen species; these reactions can be generalized as:

1 l Ground state (0 DOC) is excited by UVR to a singlet state (1 DOC*) and through

3 intersystem crossing (ISC) is transformed to the excited triplet state (1 DOC*) (Eq. 1).

-· This long-lived triplet state may react with molecular oxygen to form superoxide (O2 ) or its conjugate acid HO2 (Eq. 2) which reacts with itself to give H2O2 and O2 (Eq. 3).

-· - Finally, the interaction of HO2 and O2 results in the formation of OH and additional

H2O2 and O2 (Eq. 4) (Cooper et al 1994).

HS components and direct toxicity: While the production of ROS is thought to be the main mechanism of HS inhibition of cyanobacteria, some plants also produce specific HS that have been shown to impart direct toxicity on cyanobacteria. Extraction and analysis of chemicals released from various macrophytes reveal primarily polyphenols, low molecular weight acids, and esters (Waybright et al 2009; Choe and

Jung 2002; Bährs et al 2013; Wetzel et al 1995; Nakai et al 2001), with some polyphenols more effective than others. Polyphenols such as Tellimagrandin II have been shown in laboratory studies to impact PS II function and inhibit algal exoenzymes in eukaryotes

(Leu et al 2002). Polyphenolics from macrophyte HS have also been shown to complex with proteins such as the extracellular cyanobacterial enzyme alkaline phosphatase. This enzyme catalyzes the dephosphorylation of organically bound phosphate (Gross et al

24 1996). Thus, HS may limit the ability of cyanobacteria to access P in low nutrient conditions.

Mesocosm Conditions

The pH of mesocosms was near neutral for all treatments (Table 1) and may have increased the toxicity of HS. Plant polyphenols have been found to be significantly more effective in laboratory assays at pH 7 when compared to pH 11 (Bährs et al 2013).

Protonation of HS induces conformational changes that reduce HS electronegativity and reactivity and can affect the longevity of radicals formed through photochemistry (Paul et al 2006).

Throughout the course of mescosm experiments, oxygen was maintained at saturation using bubbling; this may have increased the toxicity of HS in comparison with other studies. Field applications of barley in low dissolved oxygen conditions are often unsuccessful in inhibiting cyanobacteria (Boylan and Morris 2003; Rajabi et al 2010). In laboratory conditions, photooxidation of HS has been shown to drop below measurable levels at O2 saturation levels below 25% (Lindell et al 2000).

Temperature variations in natural lake systems can strongly influence the inhibitory effectiveness of HS. These experiments may have underestimated in situ inhibition rates, as average mesocosm temperatures (21-22˚C) were lower than would be expected for many lake systems experiencing cyanobacterial blooms. Despite the fact that cyanobacterial growth rates are favored with increased temperatures, evidence

supports a positive correlation of increased temperature with increases in HS mediated cyanobacterial inhibition. In one laboratory assay, barley straw inhibition of M. aeruginosa was successful with a dose of 0.5g/L at 27°C, but the same experiment run at

20°C required ten times the dose (5 g/L) and a doubling of time to affect significant inhibition (Iredale et al 2012). This temperature dependent response may be a result of slower secondary reaction rates of superoxide to H2O2 (Scully et al 1996) and/or increased cyanobacterial metabolism rather than increased leaching of HS. Assays of

Typha root on M. aeruginosa showed a significant increase in treatment effectiveness at

25°C compared to 15°C, despite only a 10% increase in exudates DOC (Nakai et al

2010).

Light attenuation

The leaching of plant HS discolors water and may have reduced light levels during incubations; Prokhotskaya and Steinberg (2007) suggest that cyanobacteria may be unable efficiently absorb red light.

Continual mixing and regular inversion of mesocosms limited effects of light attenuation by HS. In similar mesocosm assays examining the effects of barley straw on

Aphanizomenon flos-aquae, it was found that when light was reduced by 80% (equivalent to shading provided by 5g/L barley), there was not a significant reduction in growth rates

(Haggard et al 2013). In fact, an 80% reduction in light results in a midday light level of roughly 200 mol photons m-2 s-1 at the bottom of the incubation jar and an average light level of 750 mol photons m-2 s-1, well above saturation for most natural phytoplankton

assemblages (Kirk 2010). Additionally, two genera in our experiments – Lyngbya and

Oscillatoria – are encouraged by low light conditions (Scheffer et al 1997). If light attenuation was affecting growth rates in this study, the expected response would be an increase in the growth of these two genera. However, Lyngbya and Oscillatoria growth rates did not increase in any treatment; rather, they were significantly inhibited in 67% and 13% of all treatments, respectively.

Microbial Role

The addition of plant material to aquatic systems increases the dissolved organic carbon concentration, resulting in increased biomass and growth rates of heterotrophic bacterial population (Wetzel et al 1995). It has been hypothesized that this increased bacterial activity is responsible for a draw-down of macronutrients (mainly P) and out- competing the phytoplankton for limited nutrients (Hessen 1992). While this mechanism may play a role in mitigating cyanobacterial growth over the long term, phosphorus levels in the short-term mesocosm experiments of this study increased from the control by 1.1 – 2.4 µg/L per gram (DW) of plant material added. Thus, nutrient draw-down by microbes is not responsible for cyanobacterial inhibition observed in this study.

Experimental effectiveness

Mesocosm applications of H. vulgare, M. spicatum, and T. latifolia revealed a significant inhibition of Microcystis in 93% of treatments, of Anabaena in 100% of

treatments, and Lyngbya in 67% of all treatments. Arthrospira was inhibited in 40% of treatments, and Oscillatoria in 13% of treatments. Ulothrix was inhibited in only one treatment, which is consistent with findings that Chlorophyta are less sensitive to HS than cyanobacteria (Prokhotskaya and Steinberg 2007; Bährs et al 2013; Gross et al 1996).

Masses of particulate debris mixing with filamentous cyanobacteria were observed in controls and precluded the microscopic observation of all cyanobacteria present, most likely as the result of these genera re-conglomerating after initial measurements. This may explain why some experiments showed negative growth rates in the controls for the mat-forming genera Oscillatoria, Lyngbya, and Arthrospira; thus, the results of these experiments most likely underestimate HS toxicity.

Nutrient dynamics

Effective barley straw treatments added up to 20 times more total phosphorus than other effective plant treatments. Barley straw used in these experiments did not receive fertilizer at any point during its cultivation; fertilized varieties may carry higher phosphorus loads. Variations in phosphorus additions may be due to TP release as the result of cell lysis and the resulting reduction phosphorus uptake. However, initial and control population biovolumes between experiments were relatively consistent and similar biovolume reductions resulted in greatly varying phosphorus loading (Fig. 24-27).

Increases in total nitrogen levels – as seen in exp. 1 – may be the result of plant leaching

or cell lysis. Reductions in TN – as seen in exp. 5 and low to mid doses of exp. 2 – may be attributed to reduced nitrogen fixation (either from cell lysis or cellular damage) or uptake of nitrogen by unaffected organisms.

H. vulgare: At nearly all applied doses, significantly inhibited the growth rate of

Microcystis, Lyngbya, and Arthrospira. All treatments showed no effect on Oscillatoria and Ulothrix.

The observation of Microcystis growth rate inhibition by barley straw application is consistent with other studies. Suppression of M. aeruginosa has been observed using either sterile liquor or direct straw application in both laboratory and field assays

(Waybright et al. 2009; Ferrier et al 2005).

Previous research indicates that the response of Oscillatoria to barley HS is variable. Oscillatoria was unaffected by barley straw applications in this study, which is consistent with findings of no effect on O. animalis (Martin and Ridge 1999). However, decreases were seen in O. tenuis and O. redekei (Everall and Lees 1997; Martin and

Ridge 1999), and growth rates increased in O. agardhii, O. lutea var. contorta, and O. lutea var. lutea (Molversmyr 2002; Ferrier et al 2005). Three of these four accounts measured Oscillatoria biomass using chl-a fluorescence, which may have reflected a shift in chl-a contained per filament rather than an increase in overall biomass, as light- induced

adjustments in chl-a content can be dramatic, particularly in laboratory settings

(MacIntyre et al 2002).

Previous research shows susceptibility of Ulothrix trentonense to barley straw application (Gibson et al 1990), while U. fimbriata and U. subtilissima show resistance

(Ferrier et al 2005). Several studies have observed an overall biomass decrease with barley application (Everall and Less 1996; Barrett et al 1999) in waters that include filamentous and planktonic chlorophyta (Gibson et al 1990).

M. spicatum: Most Myriophyllum treatments significantly inhibited the growth rates of Microcystis, Anabaena, and Lyngbya. Arthrospira and Ulothrix were moderately inhibited, and no effect was seen on the growth rates of Oscillatoria.

Microcystis and Anabaena growth rates decreased significantly (with a biovolume decrease of 93-99%) in all M.spicatum treatments. Our findings are consistent with laboratory assays against M. aeruginosa (Nakai et al 2000; Nam et al 2008) and

Anabaena (Saito et al 1989). In experimental assays against Anabaena PCC 7120 where

M.spicatum was harvested monthly (May-September) for treatments, it was found that growth rates were significantly reduced only by plants collected during August. Hilt et al

(2006) attributes this result to a shift in total phenolic compound composition during mid to late summer, which may be consistent with our findings of Anabaena inhibition by

Myriophyllum collected in late July (Hilt et al 2006).

Previous reports on the impact of M. spicatum HS on Lynbya have been largely qualitative. Anecdotal evidence from Florida residents suggests that there is a connection between the loss of M. spicatum and the appearance of L. wollei in spring-fed Kings Bay,

FL (Evans et al 2007). In the current study, M. spicatum leachates caused a statistically significant inhibition of Lyngbya at multiple doses.

Ulothrix sp. was unaffected in all treatments except for the highest dose of M. spicatum. This submerged macrophyte produces multiple algistatic metabolites (Nakai et al 2001; Nam et al 2008) including the polyphenol tellimagrandin II (Gross et al 1996), which is known to affect PS II in eukaryotes.

T. latifolia: Our results showed a significant decrease in the growth rates of all five cyanobacterial genera at various doses by green shoots (Fig. 15), brown shoots (Fig.

17), or root treatments (Fig.22). Ulothrix growth rates were not affected by any Typha treatment.

Temporal variations of phosphorus and carbon content have been documented between the shoots and roots of Typha spp. throughout the growing season (Asaeda et al

2008). For example, the phosphorus content of Typha spp. shoots has been observed to peak after approximately 50 days of growth (Weng et al 2006); thus, some temporal variability in Typha nutrient addition and HS content is expected. Given that TN loading

was similar in all brown shoot treatment doses, changes in TN were not explained solely by plant additions.

Green Shoots: Microcystis, Lyngbya, and Arthrospira growth rates were all significantly inhibited by 2.5 g/L of Typha green shoots. Findings of Microcystis inhibition are consistent with laboratory analyses of leaf application or whole plant extracts from Typha orientalis, where M. aeruginosa growth rates decreased by >75%

(after 19 days) and 50%, respectively (Chen et al 2012; Li and Hu 2005).

Brown Shoots: Microcystis and Lyngbya growth rates were inhibited by all Typha brown shoot treatments over three independent experiments. Some variability in the inhibition of both Oscillatoria and Arthrospira was evident between temporally separated experiments, where treatments in experiment 1 were more effective at equal or smaller doses than in experiments 3 and 5. While results in experiments 3 and 5 are consistent with a long-term mesocosm assay showing no effect of T. latifolia on O.angustissima

(Verb et al 2001), Typha brown shoots significantly inhibited Oscillatoria and

Arthrospira in experiment 1 (2.5 g/L). Shoots for experiment 1 were harvested in early

July, while brown shoots for experiments 3 and 5 were collected in early August and early September, respectively. The additional 1-2 months of decomposition time allotted to shoots in experiments 3 and 5 during the peak of summer would have reduced the relative HS content of these treatments, particularly as non-living plants lose their inhibitory effectiveness in natural settings after about 3 months. Thus, it may be assumed

that results found in experiment 1 are more representative of inherent inhibitory content of Typha shoots, and doses below 2.5 g/L may show effective cyanobacterial inhibition in future studies. Another possible explanation for inhibitory variance amongst temporally separated experiments includes the HS mediated inactivation of alkaline phosphatase, which would limit access to phosphorus more severely in experiment 1, where SRP averaged 0.79 µg/L, compared to experiment 5, where SRP averaged 3.3-3.6 µg/L. T. latifolia brown shoot treatment introduced little to no phosphorus, most likely because phosphorus had already been translocated to the plants roots, leached back into the sediment or adjoining plants, or consumed by bacteria and fungi.

Typha Roots: Root treatments significantly inhibited Microcystis in experiments

1 and 3, is consistent with the significant inhibition of M. aeruginosa in laboratory applications of T. angustifolia root extracts at a dose of 1 kg/L (Nakai et al 2010). Our root treatments were effective at 1/100 the dose, which may indicate a consistency with the trend that lab assays underestimate the toxicity of treatments in natural settings

(Barrett et al 1999; Martin and Ridge 1999). However, we harvested T. latifolia for our experiment from the shores of Viking Lake; relatively low lake nutrient levels may have increased the exudates present in these roots. Concentrations of exudates released from

Typha sp. have been shown to be higher when grown in low nutrient water for the first 28 days (Wu et al 2012).

When exposed to T. latifolia roots, both Oscillatoria and Arthrospira growth rates significantly decreased in experiment 1 (16.7 g/L), but were unaffected and significantly

increased (respectively) in experiment 3 (10 g/L). This may indicate a temporal variation in humic substances, the effects of alkaline phosphatase inactivation as described above, or a hormesis effect (a common toxicological dose-response trend where low dose provide stimulatory, positive responses, and larger doses elicit negative or inhibitory responses).

Low data resolution in root treatments may have been caused by microscopic root components that often littered the field of view during microscopic analysis. In addition, proportions of root stocks to rhizoids applied in treatment replicates were not considered in treatment preparation. Thus, variance amongst replicates in root treatments may be due to differences in these proportions.

General phytoplankton trends

Microcystis growth rates decreased significantly in all experimental treatments yet had the highest growth rates in controls. Microcystis may be particularly sensitive to HS due to its higher growth rates. High growth rates indicate higher metabolic demands and electron transport rates. The primary source for electrons is satisfied by PSII in autotrophic organisms, and HS can directly affect the efficiency by which electrons are transported though the photosynthetic electron transport chain. This can lead to uncontrolled electron flow to oxygen and production of ROS (Steinberg et al 2008).

Oscillatoria was generally unaffected by HS; this may be explained by several dynamics, including lower metabolic rates, HS defense mechanisms, and underestimation of toxicity due to low data resolution of controls. Oscillatoria population control groups exhibited stagnant or negative growth rates in Myriophyllum and all Typha experiments (exp. 1, 3, 4, and 5), indicating slower cyanobacterial metabolic rates and thus potentially less opportunity for electron transport disruption. However, significant inhibition of Oscillatoria did not occur during positive control growth rates of barley applications (exp. 2); rather, it was significantly inhibited by Typha root and brown shoot treatments (exp. 1) despite negative control growth rates. Thus, Oscillatoria’s limited susceptibility to HS does not seem to be explained by lower metabolic rates.

Additionally and as previously mentioned, Oscillatoria filaments were commonly entangled in particulate debris in control samples, and control populations may have been underestimated.

An alternative explanation of Oscillatoria’s resistance to HS may be explained by the presence of a HS defense mechanism. Marine cyanobacteria have been shown to exhibit both laccase and polyphenol oxidase activity (Afreen and Fatma 2013), and a homologous gene is present in the freshwater cyanobacterium Microcystis aeruginosa

SPC777. These two enzymes degrade lignins and polyphenols (components of HS), and high activities in the Oscillatoria species from the field site could potentially explain its relative resistance, particularly given Oscillatoria’s evolution in a niche of sedimentary substrates that are generally higher in HS (Whitton et al 2012). The work to date on

laccases and polyphenol oxidases has been limited to laboratory studies. Future studies on natural cyanobacterial assemblages could potentially explain variability in HS sensitivity.

The hepatotoxin microcystin is known to be produced by both Microcystis and

Oscillatoria, and the presence of microcystin has been confirmed in the lake system used in this study (Sarnelle and Wandell 2008). While this study may show the significant inhibition of toxin producing cyanobacteria by plant humics, effects of HS on cyanobacterial toxins themselves have received little attention.

Management Implications

Given the results of the current study, successful application of plant HS in the mitigation of cyanoHABs should include the following considerations: (1) application of

HS in environmental conditions essential for chemical transformations, (2) the use of macrophytes that limit nutrient loading, (3) limiting the removal of aquatic plants (or compensating for necessary plant removal with other native species) and (4) the application of macrophytes in ecological systems so as to provide sustained HS release, e.g. through the valuation and restoration of wetlands and riparian plants as well as additional active management strategies described below.

(1) As described above, exposure to light at wavelengths between 300-500 nm and adequate dissolved oxygen are essential for the production of inhibitory levels of

ROS. Meeting these conditions allows the potential for any plant production – regardless of whether they contain directly allelopathic chemicals – of compounds that inhibit cyanoHABs.

(2) As nutrient abatement is a primary management strategy in the amelioration of cyanoHABs, it is essential to limit allochthonous phosphorus sources for long term management, particularly as HS – if not continually replenished – has been shown to lose its effectiveness after several months (Iredale et al 2012). As mentioned previously, total phosphorus additions varied substantially across effective treatments (Figs. 11-15).

Barley application has been successful in mitigating cyanoHABs, but it generally introduces unwanted phosphorus to aquatic systems, requires repeated applications, and is often difficult to commercially acquire. While commercially cultivated barley tends to have higher phosphorus contents than local plants, nutrient input varies with the type and source (including cultivation methods and fertilization rates) of plant material, as well as temporal nutrient translocation. Despite the use of unfertilized barley straw, Typha and

Myriophyllum treatments consistently demonstrated the most efficient ratio of cyanobacterial inhibition to P loading. Thus, barley straw should not be considered a first choice amongst treatment options. The growth of native riparian plants will not only supply inhibitory HS, but will also absorb substantial quantities of nutrients from the aquatic system. Typha spp. nutrient absorption dynamics are described by multiple

studies (e.g. Curia et al 2011; Debing et al 2009; Gottschall et al 2007). Local and native plants are preferential choices in the prevention and mitigation of cyanoHABs.

(3) Limiting the removal of living or newly decomposing plant material from riparian areas will aid in the prevention and mitigation of cyanoHABs. Myriophyllum spicatum is a ubiquitous submerged macrophyte that is often an aggressive nuisance species, inspiring much attention by lake managers as to removal strategies (Newman

2004; Delbart et al 2013). However, removal of M. spicatum may also remove emergent properties that inhibit cyanoHABs, thus trading one nuisance for another more toxic nuisance. If M. spicatum removal is deemed necessary, immediate compensations in the form of alternative plants should be applied. While all plants have the potential to produce inhibitory compounds such as ROS, care and consideration should be given to plant selection. For example, while T. latifolia effectively inhibits cyanoHABs (and is native to northern Michigan), some Great Lakes marshes have become overrun with aggressive Typha x glauca species. These non-native species have been shown to reduce native macrophyte species density through the accumulation of its brown shoot litter

(Vaccaro et al 2009).

(4a) Wetland valuation: Increases of HS in some aquatic ecosystems may not only successfully inhibit cyanoHABs, but they may also provide positive impacts to the larger ecological system. Increases in HS may lead to shallower thermoclines, providing less area for productivity (Beisner et al 2003) and a higher susceptibility to epilimnion mixing. Shallower thermoclines may help reduce advantages otherwise conferred to

38 Microcystis and filamentous cyanobacteria by more stable stratification (Moreno-Ostos et al 2008; Posch et al 2012) as anthropogenic climate warming continues into the future.

The addition of aquatic macrophyte HS has been shown to increase the redox potential of some aquatic systems potentially increasing the rate of as demonstrated by a 218 mV increase in the presence of Typha domingensis (Aldridge and Ganf 2003), which may encourage phosphorus precipitation (Aldridge and Ganf 2003).

A hormesis effect (where low doses of a substance are stimulatory and high doses are inhibitive) is observed with HS. High concentrations of HS can cause oxidative stress in a number of freshwater organisms (Timofeyev et al 2007), moderate exposure to HS stress is positively correlated with immune enhancing responses and longevity in aquatic organisms (Steinberg et al 2008; Minois 2000).

Fringing wetlands are currently valued for their role in removal of nutrients, as a sediment trap, a sink for pollutants, and potentially pathogenic bacteria (e.g. coliform bacteria and Salmonella) from incoming waters. Ecosystem services are acknowledged for some trophic levels, such as habitat for aquatic life and support for plant richness and food webs (Sheldon et al 2005; Hudon et al 2012). Current wetland valuation paradigms have yet to acknowledge that shifts in phytoplankton dominance to cyanobacteria (along with potential production of associated toxins) can affect all trophic levels of an aquatic ecosystem; and thus, the inhibition of cyanoHABs is a valuable ecosystem service.

(4b) Active management strategies: Communities and lake managers should encourage the proliferation of riparian plants and wetland areas in the interest of water clarity and cyanoHAB prevention. Highly developed lake perimeters should buffer land-water interfaces from lawns and constructed surfaces with native plants not only for nutrient abatement, but also for the ‘steeping’ of plant HS into the aquatic system. The initiation of lake management programs to encourage, guide, and assist transitions from lawns (or other substrates) to riparian plant corridors may be particularly successful in light of cyanoHABs prevention and mitigation.

Direct, active application of plants may be a preferred or necessary action in highly urban areas or reservoirs that lack adequate substrates for plant growth. While barley straw bale application may be effective in these cases, it introduces additional phosphorus into the aquatic system, requires repeated application, and may be subject to availability. Alternatively, native and local plant HS supplementation can be viably applied in the form of floating plant islands (Hwang and LePage 2011; Nakai et al 2008,

2010; Hubbard et al 2004; Dunqui et al 2012). Floating islands are constructed from lightweight, permeable materials that support the growth of various native plants; these islands have great potential to mediate the steeping and transformation of plant HS necessary for cyanoHABs prevention and mitigation.

Conclusion

In addition to nutrient abatement, wetlands and lake-perimeter macrophytes (and even nuisance submerged plants) may have an additional value in providing HS to aquatic systems, which has demonstrated potential to reduce the extent and negative effects of cyanoHABs and provide additional benefits. The research presented here suggests that protection and/or restoration of fringing wetlands may be a viable and efficient method for cyanoHAB mitigation.

Figure 1: H. vulgare (barley straw) treatment effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira. Horizontal bars indicate statistically similar treatments.

Figure 2: Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) 2 of Microcystis (R = 0.6899; y = -0.0959x + 0.2512; p = 0.000819).

Figure 3: Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira (R² = 0.545; y = -0.0551x + 0.0889; p = 4.22e-10).

Figure 4: Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Lyngbya (R² = 0.858; y = -0.0463x - 0.0106; p = 1.52e-5).

Figure 5: Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Oscillatoria (R² = 0.694; y = -0.0208x + 0.0581; p = 0.000770).

Figure 6: Linear correlation of H. vulgare treatment doses (g/L) and growth rates (µ/d) of Arthrospira (R² = 0.799; y = -0.0567x + 0.0648; p = 3.07e-6).

Figure 7: H. vulgare (barley straw) treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen.

Figure 8: Myriophyllum spicatum treatment effects on the growth rates of Microcystis, Anabaena, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira. Horizontal bars indicate statistically similar treatments.

Figure 9: Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Microcystis (R² = 0.367; y = -0.1028x + 0.1493; p = 0.0369).

Figure 10: Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Ulothrix (R² = 0.523; y = -0.0219x + 0.1814; p = 0.00789).

Figure 11: Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Lyngbya (R² = 0.568; y = -0.027x + 0.0542; p = 0.00467).

Figure 12: Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Arthrospira (R² = 0.491; y = -0.0391x + 0.1378; p = 0.0111).

Figure 13: Linear correlation of M. spicatum treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira (R2 = 0.154; y = -0.0371x + 0.0664; p = 0.00436).

Figure 14: M. spicatum treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen.

Figure 15: Typha latifolia (cattail) green shoot treatments effect on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira. Horizontal bars indicate statistically similar treatments.

Figure 16: T. latifolia green shoot treatment impacts on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen.

Figure 17: Typha latifolia (cattail) brown shoot treatment effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira. Horizontal bars indicate statistically similar treatments.

Figure 18: Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Microcystis (R2 = 0.663; y = -0.161x + 0.2559; p = 0.00127).

Figure 19: Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Lyngbya (R² = 0.586; y = -0.0175x - 0.0133; p = 0.00370).

Figure 20: Linear correlation of T. latifolia brown shoot treatment doses (g/L) and growth rates (µ/d) of Microcystis, Lyngbya, Oscillatoria, and Arthrospira (R2 = 0.162; y = -0.0484x + 0.0833; p = 0.00459).

Figure 21: T. latifolia brown shoot treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen.

Figure 22: Typha latifolia (cattail) root treatments effects on the growth rates of Microcystis, Ulothrix, Lyngbya, Oscillatoria, and Arthrospira. Horizontal bars indicate statistically similar treatments.

Figure 23: T. latifolia root treatments on (A) total phosphorus, (B) soluble reactive phosphorus, and (C) total nitrogen.

Figure 24: Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Microcystis. Treatment doses are as follows: (A) Typha brown shoots, 1 g/L (B) M.spicatum, 12 g/L WW (C) Typha root, 10 g/L WW (D) M. spicatum, 18 g/L (E) M. spicatum, 24 g/L (F) Typha green shoot, 2.5 g/L (G) H. vulgare, 7 g/L. Numbers above bars indicate average percent decrease of cyanobacterial colonial volume.

Figure 25: Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Lyngbya. Treatment doses are as follows: (A) Typha brown shoots, 3 g/L (B) M. spicatum, 12 g/L WW (C) M. spicatum, 24 g/L WW (D) H. vulgare, 7 g/L. Numbers above bars indicate average percent decrease of cyanobacterial biovolume.

Figure 26: Increases in total phosphorus (µg/L) from controls in treatments inhibiting Oscillatoria. Treatment doses are as follows: (A) Typha brown shoot, 2.5 g/L (B) Typha root, 16.7 g/L WW (C) H. vulgare, 5 g/L (D) H. vulgare, 7 g/L. Numbers above bars indicate average percent decrease of cyanobacterial biovolume. Asterisks indicate statistically significant treatments.

Figure 27: Increases in total phosphorus (µg/L) from controls in treatments significantly inhibiting Arthrospira. Treatment doses are as follows: (A) Typha brown shoots, 2.5 g/L (B) H. vulgare, 3 g/L (C) M.spicatum, 24 g/L WW (D) Typha green shoots, 2.5 g/L (E) H. vulgare, 5 g/L (F) H. vulgare, 7 g/L. Numbers above bars indicate average percent decrease of cyanobacterial biovolume.

63 Table 1: Dates, temperature, pH, and dose levels for all treatments conducted over five mesocosm batches. All dosages are g/L dry weight (DW) measurements unless indicated otherwise. * M. spicatum WW to DW equivalents: ~1.3, 2, and 2.7 g/L DW.

64 Table 2: Tukey’s HSD p-values of experimental treatments. Values in italics were obtained from Holm-Bonferroni sequentially adjusted, Welch's t-tests. Significant values (p<0.05) are in bold. ‘NP’: not present in experiments.

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