Switching Harmful Algal Blooms to Submerged Macrophytes in

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Switching Harmful Algal Blooms to Submerged Macrophytes in Shallow Waters using Geo-Engineering Methods: Evidence from a 15N tracing study Honggang Zhang, Yuanyuan Shang, Tao Lyu, Jun Chen, and Gang Pan Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b04153 • Publication Date (Web): 12 Sep 2018 Downloaded from http://pubs.acs.org on September 13, 2018

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Environmental Science & Technology

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Switching Harmful Algal Blooms to Submerged Macrophytes in Shallow Waters

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using Geo-Engineering Methods: Evidence from a 15N tracing study

3

Honggang Zhang†, Yuanyuan Shang†, Tao lyu‡,§, Jun Chen†, Gang Pan*, †,‡,§

4

†

5

Beijing 100085, China

6

‡

7

Brackenhurst Campus, NG25 0QF, UK

8

§

9

and Environmental Sciences, Nottingham Trent University, Brackenhurst Campus,

Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences,

School of Animal, Rural and Environmental Sciences, Nottingham Trent University,

Centre of Integrated Water-Energy-Food Studies (iWEF), School of Animal, Rural

10

NG25 0QF, UK

11

* Corresponding author: [email protected] (GP)

1

ACS Paragon Plus Environment


12

ABSTRACT: Switching the dominance from algae to macrophytes is crucial for lake

13

management to human-induced eutrophication. Nutrients from algal sources can be

14

utilized in the process of transition from algal blooms to macrophytes, thereby

15

mitigating eutrophication. However, this process rarely occurs in algal bloom

16

dominated waters. Here, we examined the hypothesis that the transition of algal

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blooms to macrophytes and the transfer of nutrients from algae at different

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temperatures (8°C and 25°C) can be facilitated by using geo-engineering method. The

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results showed that the combination of flocculation and capping treatment could not

20

only remove Microcystis aeruginosa blooms from eutrophic waters but also facilitate

21

algal decomposition and incorporation into submerged macrophyte (Potamogeton

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crispus) biomass. The flocculation-capping treatment could trigger algal cell lysis. As

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compared with the control groups, the photosynthesis and respiration rate of algae

24

were inhibited and chlorophyll-a (Chl-a) concentrations were significantly reduced in

25

the flocculation-capping treatment groups. The

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and 34.8% of algae-derived nitrogen could be assimilated by Potamogeton crispus at

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8°C and 25°C, respectively. The study demonstrated that flocculation-capping method

28

can facilitate the switchover from algae- to macrophyte-dominated state, which is

29

crucial for restoring the aquatic ecosystem.

15

N tracing study revealed that 3.3%

2


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TOC

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INTRODUCTION

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Harmful algal blooms (HABs) in natural waters pose serious threats to the aquatic

34

ecosystem, environment, and public health throughout the world.1, 2 The formation of

35

algal blooms restricts light penetration into the deeper water layers, which could

36

suppress the growth of submerged macrophytes owing to the decreased

37

photosynthetic rates.2,

38

submerged macrophytes, which play a crucial role in sustaining the clear state of lakes.

39

4, 5

40

through a natural process, owing to slow and uncontrolled algal bloom die-off.

41

Moreover, the nutrients in the algae-dominated waters is always priority used for

42

algae growth rather than that of submerged macrophytes.8 In addition, the algal

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blooms induced limit light and low level of oxygen at bottom layers of water, which

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could heavily inhibit macrophyte seed germination and growth.2 Thus, removing algal

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blooms and recovering the clarity of water effectively are important for submerged

3

Restoration of clear water can trigger the growth of

However, it is difficult to achieve in water bodies with established algal blooms

3


6, 7


46

macrophytes growth and aquatic ecosystem restoration.

47

Over the past few decades, many efforts have been made to remove the algal

48

blooms, including reducing nutrient concentrations from water bodies.9 In-lake

49

geo-engineering methods have preferably tackled both eutrophication and HABs by

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adding solid-phase phosphorous (P) sorbents,

51

substances,12 and algaecides13 into water. However, the side-effects caused by the use

52

of non-biodegradable metal salts or other chemical substances have become

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increasingly concerned.11, 14 Moreover, it has been widely recognized that reducing

54

the nutrient concentrations even lower than the level when degradation of the

55

vegetation occurred is often insufficient for restoring the vegetated clear state.15, 16

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Some studies eliminated the HABs out of water column through flocculation and

57

sedimentation by the modified clay/soil.17-19 Considering that a substantial proportion

58

of nutrients in water is stored in algal cells during algal blooms,8 the modified

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clay/soil methods can speed up the algal blooms, together with nutrients inside the

60

cells settling onto the sediments in an environmentally-friendly way with lesser side

61

effects.19,

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temperature and survive on the lake bottom in a certain period, which may seed algal

63

blooms in the following years.21 Moreover, the released nutrients from the decayed

64

algal biomass could fuel the growth of algae and sustain the eutrophic status of

65

lakes.22

66

20

10

Al- or Fe-salts,11 chemical

However, many settled algal cells may tolerate the low light at low

If the algal biomass-derived nutrients could be assimilated by submerged 4


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macrophytes, it is possible to facilitate the ecosystem restoration by transferring

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excess algae-sourced nutrients into the food web .23-25 However, the nutrients released

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from the decayed HABs are always priority favored by algae rather submerged

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macrophytes during the next growing season,

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switching from the dominance of algae to that of macrophytes. Capping with natural

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soils after settling HABs has been suggested to prevent algal floc/sediment

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resuspension and reduce nutrient release into the water column.26 In addition to

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enhancing the transparence by flocculation, capping with soil or clay could improve

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sediment anoxia, 27 which makes it possible to construct suitable habitats for restoring

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submerged macrophytes.28, 29 Additionally, once the algal flocs were capped by the

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natural soil, the algal biomass should be buried and decomposed under the capping

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layer, and the nutrients released from decayed algal biomass could be retained to the

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sediments. These excess nutrients from algae have high potential for use in submerged

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macrophytes growth. Thus, the reconstruction of submerged vegetation would be

81

facilitated by reestablishing habitats with suitable light and oxygen level, and

82

available nutrients.28, 29 However, to the best of our knowledge, the effects of such

83

geo-engineering methods on the nutrient transformation process and mechanisms

84

from established algal blooms to submerged macrophytes remain largely unexplored.

8

which aggravates the difficulty of

85

In this study, algal biomass vitality and nitrogen assimilation experiments were

86

conducted to explore the process and mechanisms of switching from HABs (M.

87

aeruginosa) to the dominance of submerged macrophytes (Potamogeton crispus) in 5



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water-sediment columns. The HABs were treated by using a combination of modified

89

soil flocculation and capping with natural soils under different temperatures of 8°C,

90

25°C, and 35°C. The chlorophyll-a concentration, morphology, and photosynthesis

91

and respiration rates of the algal cells were investigated in the control, flocculation

92

treatment, and flocculation-capping treatment groups. The

93

conducted to explore the process and efficiency of Microcystis-derived nitrogen

94

uptake by the submerged macrophytes at two temperatures, i.e. 8°C and 25°C. Based

95

on these results, this study aims to examine the synergetic effects of the flocculation

96

and capping treatment on switching HABs into submerged macrophytes and

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demonstrate that the switch from HABs to submerged macrophytes could be

98

facilitated by using geo-engineering technology.

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MATERIALS AND METHODS

15

N tracing study was

100

Algae, soils, and flocculants. Microcystis aeruginosa is a well-known freshwater

101

bloom-forming cyanobacteria. The M. aeruginosa strain (FACHB-905) was obtained

102

from the Institute of Hydrobiology, Chinese Academy of Sciences, and cultured in

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autoclaved BG11 medium with 98% 15N as Na15NO3 (Sigma-Aldrich)in the laboratory.

104

All the algal batch cultures used in this study were maintained at 25 ± 1°C under cool

105

white fluorescent light of 2000–3000 lx on a 12 h light/12 h darkness regime in an

106

illuminated incubator (LRH-250-G, Guangdong Medical Apparatus Co. Ltd., China).

107

The BG11 medium with 98% 15N as Na15NO3 (Sigma-Aldrich) was supplemented in

6


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algal batch cultures 3 days before the assimilation experiment to compensate the

109

medium loss.

110

Soil was collected from the bank of Lake Taihu (China), washed with deionized

111

water, and dried for 10 h at 90°C. The soils used for flocculation and capping were

112

ground and sieved through 180 meshes (380 µm),

113

respectively. Chitosan (C56H103N9O39, Qingdao Haisheng Bioengineering Co. Ltd.,

114

China) was dissolved by adding 100 mg of chitosan into 100 mL of 0.5% HAc (1 g/L)

115

and stirring until all the chitosan had dissolved. To modify the soil, 100 mL soil

116

suspension (100 g/L) was added to 300 mL chitosan solution (1 g/L). The mixture was

117

freshly prepared for each experiment. All the containers and materials were sterilized

118

before use.

119

Algal biomass vitality experiment. Algal cultures in the mid- to late-exponential

120

growth phase were used. The experiment was conducted for 60 days in 27 plexiglass

121

cylinders with an inner diameter of 8.4 cm and height of 50 cm (Figure S1a).

122

One-liter bloom water with Chl-a concentration of 5670 µg/L (7.3–7.7×107 cells/mL)

123

was filled into the columns. The incubation temperatures of 8°C, 25°C, and 35°C were

124

selected to simulate the real temperature in Lake Taihu (China) at spring, early

125

summer, and midsummer, respectively. Lake Taihu has serious HABs annually, where

126

the algae start to grow fast in spring and the bloom happens in summer.

127

Eighteen columns were selected randomly and treated by modified soils for

7



128

flocculation. The modified soil suspension was added to the bloom water and stirred

129

by using a glass rod. The final concentrations of the modified soils in each column

130

consisted of 3 mg/L chitosan and 100 mg/L soil. The flocculated columns were kept

131

standing for 3 h to allow the sedimentation of the algal flocs. Subsequently, nine

132

columns from the flocculated columns were covered with a 1 cm-thick layer of

133

natural soil. The nine columns were treated only with flocculation were labeled “F-no

134

capping” and the nine columns treated by flocculation-capping treatment were labeled

135

“F-capping”. The remaining nine columns without any treatment were set as the

136

“control”.

137

All the columns were encircled with a cloth about 15 cm from the bottom to

138

ensure darkness. Thereafter, the nine columns from each treatment group (control,

139

F-no capping, and F-capping) were separated equally to three parts (three columns for

140

each) and incubated under 8°C, 25°C, and 35°C with fluorescent light (2000–3000 lx,

141

12 h light/12 h darkness) (Figure S1 a). In order to allow the algae vitality in different

142

treatments at the same level and make the algal cultures adapt to the new temperature,

143

all the columns were stabilized under corresponding temperature condition for 10 h

144

before sampling.

145

The vitality experiment lasted for 60 days and Chl-a concentrations were

146

measured at day 0, 15, 30, 45, and 60. The samples for day 0 were taken from 10 cm

147

below the surface water of the control group and filtered with a 0.45 µm membrane.

148

The samples from the F-no capping and F-capping groups were the deposited algal 8


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flocs at the bottom of the columns. For Chl-a concentration analysis of all the samples

150

were extracted by acetone (90%) for 24 h at 4°C and measured with a

151

spectrophotometer.16 The same samples from day 0 and 60 were used to analyze the

152

morphology, photosynthesis, and respiration. The samples were centrifuged at 6000

153

rpm for 3 min and pre-fixed with 2.5% glutaraldehyde for 4 h, and washed with

154

phosphate buffer solution. Thereafter, the samples were post-fixed with 1% osmium

155

tetroxide for 2 h, and again washed with phosphate buffer solution. The washed

156

samples were dehydrated twice through a series of 30%, 50%, 70%, 85%, 95%, and

157

100% ethanol solutions and dried with a vacuum drier. Completely dry samples were

158

then mounted on a copper stub, coated with gold, and examined with a scanning

159

electron microscope (SEM, S-3000N, Hitachi, Japan). For photosynthesis and

160

respiration analysis, the samples were added to the micro-breathing bottle (4 mL), and

161

cultured under the same conditions as those for algal batch cultures. After transferring

162

the sampling bottles into the incubator, photosynthetic and respiratory rates were

163

measured with a micro-respiration system (MRS, Unisense, Danmark). The O2

164

concentration was measured continuously for 60 s every 2 min in each sample by an

165

O2 microsensor within a whole culture cycle (i.e., 10-h light/10-h darkness regimen).

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Nitrogen assimilation experiment. After incubation with algae, the

167

aeruginosa cells were collected by a 30 µm mesh and rinsed at least ten times with

168

deionized water to remove the unassimilated 15N-NO3. The resulting δ15N value of the

169

labeled M. aeruginosa was 1072 ± 13‰ (n = 2), and a certain dosage of algae were 9


15

N-labeled M.


170

added into filtered lake water (30 µm), which was used to form bloom water (7.3–

171

7.7×107 cells/mL). In total, 40 columns (diameter 8.4 cm and height 50 cm) were

172

filled with 10 cm of sediment and 1.6 L of bloom water and stabilized for 3 days

173

before the experiment. The sediment was collected from Lake Taihu, China. A 15-cm

174

above the bottom of the column was encircled with a cloth to avoid the effects of

175

ambient light on the sediment. All the columns were treated by modified soil for

176

flocculation and then capped with a 1 cm thick layer of nature soil. Twenty columns

177

were planted with Potamogeton crispus seedlings after the capping treatment and

178

named as the “vegetated” group. The other 20 columns remained unvegetated and

179

were named as the “unvegetated” group. Thereafter, 10 columns each from the

180

vegetated and unvegetated groups were cultured in the illuminated incubator at 8°C.

181

The remaining 10 columns from each group were incubated at 25°C (Figure S1b).

182

Both incubation conditions were set under fluorescent light (2000–3000 lx, 12 h

183

light/12 h darkness). Temperatures of 8°C and 25°C were selected to simulate the

184

germination and rapid growth stage of P. crispus in Lake Taihu (China) at spring and

185

early summer. Plant (seedling) and sediment samples (the top 5 cm) were collected

186

right after the flocculation-capping treatment (day 0) and day 10, 17, 27, and 45.

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During each sampling event, two random columns (treated as duplicates) were visited,

188

and the entire plant biomass was harvested from them.

189

The sediment and plants were separately homogenized, dried, and analyzed for

190

stable nitrogen isotope ratio (15N/14N) using a Delta Plus Advantage mass 10


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15

191

spectrometer (Finnigan MAT) connected to a Flash EA1112 elemental analyzer.

192

abundance was expressed using the conventional delta notation against the

193

atmospheric nitrogen standard:

194

δ 15N (‰) = ( 15 N

195

Moreover, to compare the labeling 15N accumulation in the sediments and plants

196

from different treatment groups, the excess 15N should be calculated for the absolute

197

amount of the incorporated labeling

198

concentration of

199

equations: 30

15

14

Nsample

15

N

14

N

N s tan dard − 1) × 1000

15

N. The data are presented as excess

N in the dry sample and calculated according to the following

µmol of N in sample × 200

201

at%15 N sample-at%15 N control

100 gram of dry sample

Excess 15 N(µmol / g ) =

at%15 N sample=

(1)

100 × Rair × (

(2)

δ 15 N sample

1+Rair+Rair×

+1 ) 1000 δ 15 N sample

(3)

1000

202

The analytical error between repeated measurements was typically within

203

±0.1‰. where at%15Ncontrol represents the value on day 0, and δ15N is expressed as an

204

excess value relative to the atmospheric nitrogen ratio, Rair = 0.0036765.

205

Statistical analysis. Origin 8.0 (OriginLab, Northampton, MA, USA) and SPSS 16.0

206

(IBM Corporation, Armonk, NY, USA) were used for plotting and data analysis,

207

respectively. Significance levels for all the comparisons were set at P < 0.05. In the

208

algal biomass vitality experiment, a two-way ANOVA with post-hoc Duncan’s 11



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multiple range test was used to compare the Chl-a concentrations, photosynthesis and

210

respiration rates separately among the different treatment groups at different

211

temperatures. In the test, the groups (control, flocculation, and flocculation + capping)

212

and temperatures (8°C, 25°C, and 35°C) were the two independent factors, and Chl-a

213

concentration, photosynthesis, and respiration rates were the dependent factors in each

214

analysis. The Chl-a concentration in the same treatment group at each temperature

215

was tested by a one-way ANOVA with post-hoc Turkey’s test. In the nitrogen

216

assimilation experiment, the difference between the ability of

217

by P. crispus was analyzed by a two-way ANOVA with post-hoc Duncan’s multiple

218

range test. The treatments (vegetated and unvegetated) and temperatures (8°C and

219

25°C) were the independent factors, and

220

factor. Moreover, linear correlation analysis was conducted to test the relationship

221

between 15N assimilation rate and P. crispus biomass through the experiment.

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RESULTS

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Algal biomass vitality. The initial Chl-a concentration in all the columns was 5670

224

µg/L. In the control groups, Chl-a concentrations were significantly higher than the

225

initial values and reached 7397 and 6731 µg/L at 8°C and 25°C, respectively, on day

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60 (Figure 1). Chl-a concentrations at 35°C improved significantly to 9224 µg/L at

227

day 15 and decreased gradually to approximately 2243 µg/L at day 60. In both F-no

228

capping and F-capping groups, the concentrations of Chl-a showed continuous

229

declines with sampling time under all the incubation temperatures. A significant

15

15

N assimilation rates

N assimilation rate was the dependent

12


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difference was observed for the corresponding samples between F-no capping and

231

F-capping groups at day 60 (P < 0.05) (Table S1). At 8°C, 25°C, and 35°C, Chl-a

232

concentrations decreased to around 3444, 2277, and 18 µg/L, respectively, in the

233

F-capping group, and the values were 4203, 2574, and 500 µg/L, respectively, in F-no

234

capping group. Moreover, higher temperature accelerated the decrease in Chl-a

235

concentrations in both the F-no capping and F-capping groups (P < 0.05) (Figure 1).

236

The interaction of temperature and treatment did not show significant effects on the

237

Chl-a concentrations changes (P > 0.05) (Table S1). 8ºC

Chl-a (µ µ g/L)

10000

25ºC

35ºC

8ºC

10000

8000

6000

6000

6000

4000

4000

4000

0

0 10

20

30

40

Time (d)

50

60

35ºC

F-capping

F-no capping

0

25ºC

2000

2000

0

8ºC

10000

8000

Control

239

35ºC

8000

2000

238

25ºC

0

10

20

30

40

Time (d)

50

60

0

10

20

30

40

Time (d)

50

60

Figure 1. Concentrations of Chl-a in different treatment groups along the experiment.

240

The algal cells collected on day 0 showed intact morphology with no obvious

241

differences among the control, F-no capping, and F-capping groups (Figure S2). At

242

the end of the experiment, most algal cells collected from the three systems generally

243

showed intact morphology at 8°C (Figure 2). However, the algal cells collected from

244

the F-capping group incubated at 25°C were obviously deformed and lysed as

245

compared with those from the control group. Moreover, more lysed cells were

246

observed in the F-capping group incubated at 35°C than in the control and F-no 13



247

capping groups (Figure 2).

c

248 249

Figure 2. SEM images of algal cells in different treatment systems incubated for 60 days at 8oC,

250

25oC and 35oC.

251

At the beginning of the experiment, the M. aeruginosa cells collected from all

252

the three groups could sustain normal photosynthesis and respiration rate, which

253

showed that the oxygen produced in the light stage was sufficient to maintain the

254

respiration of the algae in the dark phase (Figure S3). The algal cells collected from

255

both control and F-no capping groups sustained photosynthesis at 8°C and 25°C after

256

60 days, as reflected by the positive oxygen change rates in the light stage (Figure 3 a,

257

b, c, and d). However, the photosynthesis efficiency was eight times lower in the F-no

258

capping group at 25°C than that in the control group. Although the cells collected 14


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from the F-capping group could sustain photosynthesis in the light phase at 8°C, the

260

efficiency was much lower than those in the control. It should be noted that the O2

261

change rates for the cells in F-capping group incubated at 25°C showed negative

262

values even in the light incubation phase (Figure 3h), indicating that the death of algal

263

biomass had occurred. Death and decay of algal cells were found in all the three

264

systems after 60 d of incubation at 35°C, which was reflected by negative O2 change

265

rates (Figure 3 c, f, and i). However, flocculation-capping treatment accelerated algal

266

cell death and decay, which could be reflected by the significantly higher

267

consumption rates of O2 from the F-capping group than from those either of the F-no

268

capping or control (P < 0.05). It should be noted that the interaction between

269

temperature and treatment significantly influenced the photosynthesis and respiration

270

rates (P < 0.05) (Table S1). O2 Change Rates (mg/L/h) 18

32

a

12

Dark

Light

O2 Change Rates (mg/L/h)

8

12

16

d

Dark

Light

0 -6 0

32

12

16

6

8

0

18

4

8

12

16

20

Dark

Light

6

Dark

Light

8

12

16

18

4

8

12

16

Dark

Light

16 6

0

8 0 4

8

12

Time (h)

16

20

0 -6 0

8

12

16

20

32

f

24

Dark

Light

16 8

0 -6 20 0 32

h

24 12

6

0 4

8 0

0 -6 20 0 32

g

12

16 8

16 6

8 0 4

18

Dark

24 12

16 6

0

24

Light

0 -6 0 32

e

24 12

-6 0

271

20

32

c

18

0 4

8

12

16

i

Dark

4

8

12

16

20

32

24 12

Light

O2 Concentration (mg/L)

4

18

24 12

8 0

-6 0

-6 0

32

b

16 6

0

18

O2 Concentration (mg/L)

24 12

6

18

18

24

Dark

Light 16 6

16

8 0

8

20

0 -6 0

Time (h) 15


0 4

8

12

Time (h)

16

20


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Figure 3. Photosynthesis and respiration of M. aeruginosa cells in different groups after

273

incubation for 60 days (a: 8oC-control, b: 25oC-control, c: 35oC-control; d: 8oC-F-no capping,

274

e: 25oC-F-no capping, f: 35oC-F-no capping; g: 8oC-F-capping, h: 25oC-F-capping,

275

i:35oC-F-capping).

276

Nitrogen assimilation. After the flocculation-capping treatment, the δ15N enrichment

277

in the sediment samples from all the columns were 18.47‰. The values were

278

continuously decreased along the experiment and reached approximately 14.9, 12.9,

279

8.1, and 7.3‰ in unvegetated-8°C, vegetated-8°C, unvegetated-25°C, and

280

vegetated-25°C groups, respectively (Figure 4). Generally, the vegetated groups

281

showed a significantly lower δ15N in the sediments than that in the unvegetated

282

groups after day 10 under both the incubation temperatures. Moreover, both the

283

vegetated and unvegetated groups showed a significantly lower δ15N at 25°C than that

284

at 8°C from day 10 to 45 (P < 0.05)(Table S1). unvegetated-8℃ unvegetated-25℃

20 18

vegetated-8℃ vegetated-25℃

14 12

δ

15 N (‰)

16

10 8 6 0

285 286

10

17

27

45

Time(d) Figure 4. The δ15N in sediment collected from different columns during the 45-d experiment. 16


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287

A total of 1.38 µmol15N/g labeled algae was filled in each column before 15

288

flocculation. The mass balance calculation of excess

289

conducted at the end of experiment (Figure 5). The excess 15N (0.48 µmol 15N/g) in P.

290

crispus collected at 25°C was 10 times greater than that at 8°C (0.045 µmol15N/g).

291

Approximately 20% of the excess

292

8°C. However, approximately 55–75% of excess 15N was unaccounted in the columns

293

incubated at 25°C.

15

N in each system was

N was unaccounted in the system incubated at

Unaccounted

P. crispus

Sediment

Excess labeled material (%)

100 80 60 40 20 0

294 295

d ed d ed getat -vegetate nvegetat -vegetate e v n u u ℃ ℃ 5 8 ℃ 2 8℃ 25 Figure 5. Mass balance of 15N in the different treatments after 60 days of incubation.

296

In addition, the increase rate in δ15N in P. crispus at 25°C was five times higher

297

than that at 8°C during the experiment (Figure 6). The increase in 15N enrichment was

298

significantly correlated with the increase in P. crispus biomass (r = 0.915, P < 0.05),

299

where the biomass of P. crispus grown at 25°C was double of that grown at 8°C.

17


δ15N(‰) in 8℃

δ15N(‰) in 25℃

biomass in 8℃

biomass in 25℃ 1.50

40

1.25

30

1.00

15

δ N (‰ )

50

Page 18 of 32

0.75

20

Biomass (g)


0.50

10

0.25 0

10

17

27

45

Time (d)

300 301

Figure 6. The δ15N in P. crispus and the plant dry biomass during the experiment.

302

DISCUSSION

303

HABs sedimentation using modified clay/soils. The removal and management of

304

the growth of blooms, especially cyanobacterial blooms, is an important step in the

305

recovery of eutrophic lakes before the re-emergence of macrophytes. In this study, the

306

modified soil was selected for accelerating the sedimentation of algal blooms. The

307

soil particles provided the algal biomass with sufficient ballast to counteract the

308

buoyancy of the M. aeruginosa cells in the water columns. The algal flocs settled to

309

the bottom of the columns, whereas M. aeruginosa was mainly suspended in the

310

control water columns (Figure 1). Although the Chl-a concentrations in the F-no

311

capping and F-capping treatment groups showed similar declining trends at each

312

temperature after the application of the modified soils, the Chl-a concentration in each

313

sampling point from the F-no capping group was slightly higher than those from the

18


Page 19 of 32


314

F-capping group. It was attributed to the fact that more M. aeruginosa cells survived

315

in the F-no capping columns than in the F-capping columns. The surviving algal

316

biomass may return to the water columns, especially in shallow waters, where wind

317

and wave-induced turbulence occurred.26

318

The modified soil flocculation and capping with natural soils caused little

319

damage to the M. aeruginosa cells, as reflected by the intact cell morphology and

320

normal photosynthesis and respiration at day 0 (Figure S3). This contributed to the

321

visually observation that no homogeneously green or yellow color appeared around

322

the flocs, which suggested cell lysis in this type of laboratory experiments.

323

However, other analyses, such as dissolved Chl-a, toxins, and nucleic acids should be

324

conducted to prove the visual observation in the further studies. The observed intact

325

algal cells after the flocculation-capping treatment may be important for preventing

326

the intracellular cyanotoxins or excess nutrients abruptly released to the environment

327

in practice. 18, 32 However, the chitosan, which was used to modify natural soils in this

328

study, may possess antimicrobial activities against some bacteria,32,

329

cyanobacteria species.31,34 The dose of flocculants should be considered seriously in

330

practice. It was reported that a higher dose of chitosan (e.g., >8 mg/L) could lead to

331

cell lysis of M. aeruginosa. 32 The lower dose (3 mg/L) of chitosan in the present

332

experiment is similar to those reported by Miranda et al. (2017), who found no

333

detrimental

334

chitosancombining natural soils could lower the toxic risk on the aquatic organisms

effects

on

Microcystis.35 Moreover,

19


in

our

33

10, 31

including

previous

study,


335

exerted by chitosan alone further.36

336

Vitality changes in settled M. aeruginosa. Capping with soils can keep the settled M.

337

aeruginosa cells in darkness, which is a key factor affecting the photosynthetic rates.3

338

In the present study, the photosynthesis and respiration rate of M. aeruginosa cells

339

were severely hindered after the flocculation and capping treatment, which may

340

trigger the decomposition of algal cells. The photosynthesis and respiration effects of

341

M. aeruginosa cells could be inhibited in F-no capping group as reflected by the

342

significantly lower change rate of O2 respiration than those in control. However, a

343

significant lower rate of photosynthesis and respiration rate was observed in the

344

F-capping groups, which indicated an even higher photo-inhibition effect through the

345

F-capping treatment (Figure 3 g–i). The results mentioned above confirmed the

346

hypothesis that flocculation-capping treatment can accelerate the algal bloom die-off.

347

It should be noted that the interference of other bacteria, such as heterotrophic

348

bacteria, on the algal cell respiration should be investigated in further study.

349

The interaction between the treatment and temperature could significantly affect

350

the photosynthesis and respiration rates of the deposited M. aeruginosa cells in our

351

experiment (P < 0.05) (Table S1). Temperature is a crucial factor for the vital

352

activities of cyanobacteria in natural waters. In the present study, the three

353

temperatures (8°C, 25°C, and 35°C) were established to simulate the real temperature

354

at spring, early summer, and midsummer in Lake Taihu, China. The dominant

355

cyanobacteria fast growth in spring and the blooms occur annually during the summer 20


Page 20 of 32

Page 21 of 32


356

in Lake Taihu.37 The present results showed that the deposited algal biomass tends to

357

be tolerant to low light at lower temperatures, as reflected by the normal morphology,

358

photosynthesis, and respiration in the control group (Figure 2 and 3). Similarly, Ma et

359

al (2016) found that most cyanobacteria sank to the sediment layer and remained

360

dormant as viable inoculants (akinetes) below 12.5°C.38 These deposited algal cells

361

can return to the water column as a potential source of bloom formation when higher

362

temperatures occurred.38 The higher temperature stimulated the growth of M.

363

aeruginosa cells, as reflected by the faster and higher increasing rates of Chl-a in

364

controls at 35°C than at 25°C before 15 days (Figure 3). The consumption of O2 in

365

F-capping systems also increased as the temperature increased, and the O2 respiration

366

became negative at 25°C, especially at 35°C, after 60 days of incubation (Figure 3),

367

indicating that higher temperatures accelerate the respiration rate of algal blooms

368

buried under the capping layer.

369

Assimilation of nitrogen from algae by submerged vegetation. In lakes, nutrient

370

cycling occurs in the sediment, with algal sedimentation showing a strong effect on

371

biogeochemical processes. The decomposition of algal blooms can release nutrients

372

directly, which could lead to changes in nutrient composition cycling in sediment and

373

water.39-41 In this study, we found that nitrogen was released into the sediment from

374

the settled M. aeruginosa, and then absorbed by P. crispus (Figure 4, 5). The

375

unaccounted

376

perturbation and mineralization.42 The organic nitrogen, including

15

N through the

15

N balance calculation may be due to benthic

21


15

N, could be


Page 22 of 32

377

degraded to inorganic fractionation with net loss through nitrification and

378

denitrification reactions into gaseous phases.43 Another fraction of the unaccounted

379

15

380

Moreover, higher temperatures could trigger greater Microcystis-derived nitrogen

381

release from sediments, which is consistent with the findings of other studies

382

suggesting that the nutrient cycling rates increased with the addition of settled algal

383

blooms and elevation of temperatures.39

384

N may because of the nutrients released into the overlying waters (Figure 5).

The uptake of nutrients by macrophytes plays a vital role in the mitigation of

385

internal nutrient loads in vegetated sediment of lakes.5 In this study, excess

386

detected in the P. crispus biomass, which suggests that the flocculation-capping

387

treatment could transfer nutrients from HABs into submerged macrophyte growth.

388

Thus, it is possible to reduce excess N from algae released into water columns

389

(Figures 4–6). This is the accepted method of restoring a healthier ecological system

390

dominated by submerged vegetation in shallow waters in previous studies.28, 44 The

391

rapid uptake of δ15N at both 8°C and 25°C mainly occurred within the first 10 days in

392

this study (Figure 4), which is consistent with the findings of the rapid uptake of

393

labeled ammonium and nitrate by common reeds.45 Additionally, the assimilation of

394

nitrogen by submerged vegetation can occur directly by the uptake of nitrogen from

395

water columns.46 This may attribute to the partial decrease in the unaccounted labeled

396

N content in the vegetated groups (Figure 5). Higher temperature could facilitate the

397

assimilation of Microcystis-derived nitrogen into P. crispus (Figure 6). It may be 22


15

N was

Page 23 of 32


398

partially because of the decomposition of deposited algal biomass and organic

399

nitrogen mineralization process in the sediment, which could be facilitated at higher

400

temperatures to produce more nutrient source for P. crispus. In addition, most aquatic

401

plants grow from the spring to midsummer in temperate lakes, which is consistent

402

with our result that the growing rate of P. crispus was twice as high at 25°C than at

403

8°C (Figure 6). The growth rate significantly affects the incorporation of δ15N in P.

404

crispus, as reflected by the five-fold higher δ15N‰ found at 25°C than at 8°C. Further

405

studies should focus on the mineralization rate of deposited algal blooms associated

406

with P. crispus growth after the proposed treatment.

407

Implications for lake restoration. Generally, the change from the dominance of

408

algae to that of macrophyte in lakes subjected to human-induced eutrophication can

409

be difficult to achieve under natural conditions due to persistent excessive growth of

410

algal biomass. Restoration of such lakes from an established algal bloom to a desired

411

state dominated by submerged macrophytes requires significant intervention, even

412

after reducing external nutrient inputs. Therefore, many in-lake geo-engineering

413

methods have been widely used as environmentally-friendly, efficient, and

414

economical methods of accelerating the removal of algal blooms from water.19, 20, 28

415

The improvement in transparency and dissolved oxygen concentrations in bottom

416

water resulted from the application of modified clay/soil technology,

417

facilitate (e.g., establishing a certain period for plant germination and growth)

418

reconstruction of submerged macrophytes. Flocculation-capping methods, as shown 23


19, 28

which can


419

in this study, can not only eliminate the algal biomass, but also facilitate their

420

degradation. Then, the nutrients released from decayed algal biomass can be utilized

421

for the growth of submerged vegetation. Higher temperatures accelerated both the

422

decomposition and incorporation of algae into plant biomass, implying that

423

application of such geo-engineering method combined with seeding macrophytes

424

during the period of algal blooms can facilitate such transformation due to the overlap

425

of growing seasons between algae and submerged vegetation, especially in temperate

426

lakes. However, a pilot field experiment is necessary to test the potential effects of

427

such in-lake geo-engineering methods for both controlling algal blooms and

428

facilitating the transition from the state of algal dominance to macrophyte dominance

429

state in lakes.

430

ASSOCIATED CONTENT

431

Supporting Information

432

Figures showing SEM images of algal cells at the beginning of the experiments (0

433

day), Photosynthesis and respiration of M. aeruginosa cells in different systems: a:

434

25oC-control-0d, b: 25 oC-F-no capping-0d, c:25oC-F-capping-0d. Table showing

435

results of the analysis of variance (ANOVA) on the effects of the Temperature and

436

Treatment, and their interactions on Chl-a, photosynthesis and respiration rate, δ15N in

437

sediment, and δ15N in P. crispus.

438

AUTHOR INFORMATION

439

Corresponding Author

440

*Corresponding author: Tel.: +86 10 62849686; Fax: +86 10 62849686; E-mail 24


Page 24 of 32

Page 25 of 32


441

address: [email protected]

442

Notes

443

The authors declare no competing financial interest.

444

ACKNOWLEDGEMENTS

445

The research was supported by the National Natural Science Foundation of China

446

(41877473,41401551);

447

(XDA09030203); the National Key Research and Development Program of China

448

(2017YFA0207204); and Beijing Natural Science Foundation (8162040).

449

AUTHOR CONTRIBUTIONS

450

G.P. designed the research; H.Z., Y.S and J.C, performed research; H.Z. analyzed the

451

data; H.Z. wrote the paper, G.P. and T.L. contributed significant revision and language

452

improvement.

the

Strategic

Priority

Research

453

25


Program

of

CAS


454

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