critical review of paper for wastewater class project

CIVL 7230 PROJECT

Critical Review of “Enhanced salinities, as a proxy of seawater desalination

discharges, impact coastal microbial communities of the Eastern

Mediterranean Sea” Article Authors: Natalia Belkin, Eyal Rahav, Hila Elifantz, Nurit Kress, and Ilana Berman-Frank

Review by: Brock Horsley

11/27/2015

Belkin, N., Rahav, E., Elifantz, H., Kress, N. and Berman-Frank, I. (2015), Enhanced salinities, as a proxy of seawater desalination discharges, impact coastal microbial communities of the eastern Mediterranean Sea. Environmental Microbiology, 17: 4105–4120. doi: 10.1111/1462-2920.12979

1

Objectives

Desalination plants discharge effluent brine into surface waters often at high

concentrations of salinity relative to ambient levels. While the environmental impact of

desalination effluent has previously been discussed, there are few laboratory studies on this

topic. Salinity has been considered the most important driver of the global distribution of

bacteria and other microbial life. The objective of the study was to perform a laboratory study to

examine the impact of increased salinity on planktonic microbial life in the coastal Eastern

Mediterranean Sea (EMS) of Israel.

Methods

Two experiments were performed: one from April 23rd

to May 5th

and another from July

7th

to July 18th

of 2013. The first one will be referred to as mixed-spring and the second as

summer-stratified. Time periods were selected to reflect different conditions in surface water

bodies. It was expected that initial conditions at each time period would influence microbial

environments. Both experiments contained three types of samples: a control seawater sample, a

5% added salinity sample, and a 15% added salinity sample. Three copies of each sample were

made for later statistical analysis. Each sample was a bag of seawater roughly 1 m3 in size, all in

a 16 m3 pool of circulated seawater. To account for the effects of isolation on microbial life in

each sample, the 5% and 15% added salinity samples were compared to the control samples. The

5% and 15% experiments were chosen to simulate conditions that were found at a nearby

desalination discharge site. The seawater was pumped from a site 300 m off of the coast and at a

depth of 2 m. To prevent evaporation, dilution, or contamination, the samples were covered with

polyethylene tops that still allowed gas exchange with the atmosphere. The tops also allowed for

50% surface light penetration.

2

Samples were taken from each bag two hours after brine was added. This time was

allotted so that each sample’s properties could be fully characterized. Each sample was

approximately 5 to 10 L, collected gravitationally for 11 to 12 days. Samples taken each day

were measured for salinity, temperature, chlorophyll (Chl a), bacterial productivity (BP), and

primary productivity (PP). DNA samples were also taken determine bacterial and eukaryotic

compositions. Samples were frozen until time to analysis to prevent decomposition. Quality

assurance was performed by national laboratories in the US, Canada, and Japan.

Results & Conclusions

Autotrophs experienced a decrease in photochemical efficiency, and heterotrophs saw

enhanced bacterial activity. Salt stress can inactivate the photosynthetic reaction centers and

inhibit protein synthesis. The data collected suggested that the immediate effect of salt addition

was salt stress on autotrophs and suppression of photosynthesis. For the 15% increased

experiment, PP decreased significantly in spring and summer tests. For mixed-spring, PP went

from 3.4 to 1.7 ug C/L/h, and stratified-summer went from 2.9 to 0.9 ug C/L/h. The decrease was

attributed to the immediate death of salt-sensitive phytoplankton. In heterotrophs, BP increased

by 2.5 times in mixed-spring and by 1.5 times in stratified-summer. The rapid BP increase was

attributed to the need of bacteria to self-regulate in response to the influx of salt. This includes

osmotic stress on cells, internal pH, and energetic potential of membranes. As for community

composition, increased salinity did not have a significant effect in mixed-spring when compared

to control. However, salinity was the dominant driver of community composition in stratified-

summer. For the control in stratified-summer, bacterial communities decreased in diversity by

51%, while eukaryotic communities increased in diversity by 252%. In the 15% addition

experiment for stratified-summer, bacterial diversity decreased by 87% and eukaryotic diversity

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decreased by 70%. In the 15% addition, cyanobacterial OTUs increased by 140%, while they

decreased by 90% in control. However, proteobacterial OTUs decreased by 30% in the 15%

addition, while they increased by 45% in control.

Although cyanobacterial OTUs increased in the 15% addition, Prochlorococcus marinus

(a major cyanobacterial order) disappeared after the first day of treatment. Studies have shown

that P. marinus contributes 9 to 18% of global marine carbon fixation. Among heterotrophs,

Pelagibacter declined significantly after six days in the 15% addition. Pelagibacter accounts for

roughly 30% of the bacteria in global oceans. It has been theorized that Pelagibacter consumes

osmolytes produced by P. marinus. Therefore, planktonic food webs near effluent sites of

desalination plants may change dramatically in the stratified-summer season of the EMS.

Turning to other microbial life, the diatom Gyrodinium was relatively low in abundance in

control during the stratified-summer experiment. In the 15% addition experiment, Gyrodinium

population was up 12% compared to other eukaryotic OTUs recorded. Some species of

Gyrodinium are known to form red tides and other harmful blooms. Thus, their increased

appearance in highly saline environments is of concern in desalination plant discharge sites.

The results of the experiments suggest that microbial communities have higher functional

plasticity in response to salinity in the mixed-spring, while they can be controlled by salinity

changes in the stratified-summer. Since biodiversity can help maintain the stability of an

environment to changes in the ecosystem, maintaining abundance of adaptive species can help

stabilize an environment that experiences change salinity. Salinity conditions can be a driving

factor in community composition of planktonic communities. An increase in salinity may reduce

species abundance, which would make to environment select for organisms that thrive in high-

salinity environments. Seasonal changes in community composition can buffer against these

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types of environmental stress. Ultimately, decrease in biodiversity that is caused by an increase

in salinity may destabilize the aquatic food web. The study recommended that other discharges

from desalination plants, such as coagulants and anti-scalants should be examined in future

studies.

Analysis

For strengths, the study had accurate modeling of the real-world scenario. The

concentrations tested were based on conditions at actual effluent sites. Seawater used was from

the EMS itself. The microbial community composition was seasonally consistent in spring and

summer. Continuous mixing was done to emulate sea mixing. The reporting had professional

statistical analysis and quality assurance. All tested means employed analyses of variance

(ANOVA). UNIFRAC Monte Carlo tests were used for comparing biological communities. The

study also took consideration for observational errors. All parameters were compared to controls.

Quality assurance was performed by national laboratories in the US, Canada, & Japan. They

assessed methodology for each parameter that had standardized testing. The study had good

explanations of tangible concerns, and provided direct links to the effects of increasing salinity.

Connections between change in microbial environment and known effects were made. The two

main connections were causation of red tides and changes in the food web. The study managed

to linked increases and decreases in microbial taxa to change in salinity. Species with elastic

response to change in salt will thrive, while some essential species in the local food web are

inelastic and become eradicated.

For shortcomings, the study presents other concerns about desalination plants with

limited discussion. It brings up how other substances in discharges from desalination plants may

be impact microbial communities. It seems to mention this as a major conclusion from the paper.

5

While it is a topic related to desalination plants, it is unrelated to the effects of changing salinity

in water. This concern should have presented as a side-thought, rather than among conclusions

from the research. Some of the laboratory methods did not have enough explanation for those

unfamiliar with the research. To those unaccustomed to lab techniques, some detailed were

understated or unclear. For instance, it was unclear how covers on experiment samples both

prevent contamination of atmospheric inputs but maintain gas exchange with the atmosphere.

Other details seemed unexplored, such as the effects of freezing the samples might have on

anticipated results.

Turning to limitations of the research, as desalination processes are a growing practice,

research into effects of discharge are not abundant. Many studies discuss implications of

increased salinity, but few actual lab studies exist. There are also few field measurements to

determine the impact of effluent discharge. It is hard to determine long-term conditions of

seawater when field data is limited. Since the study is contained to a 16 m3 zone, the effects of a

larger boundary zone are also unknown. It would be of interest to see if the same effects on a

microbial community would occur if there was no clearly defined boundary, as in a real seawater

scenario.

One of the unique findings of the paper involves how the increase in salinity led to

immediate decreased photochemical potential. A decrease in Chl a followed, reflecting death of

salinity-sensitive phytoplankton. It was previously known that salinity elevation leads to reduced

Chl a in some algae and cyanobacteria. However, there were no previous records found of rapid

changes within 2 hours of salt addition.

Critical Review of “Enhanced salinities, as a proxy of seawater desalination discharges, impact

coastal microbial communities of the Eastern Mediterranean Sea”

Review by: Brock Horsley

Objectives

Objectives

• Perform lab study on impact of increased salinity on microbial community in Easter Mediterranean Sea (EMS)

– As a result of the discharge of high salinity from desalination plants

– Impact of desal plants has been discussed, few lab studies

Methods

Methods

• Two Experiments Performed

– APR 23 – MAY 5: Mixed-Spring

– JUL 7 – JUL 18: Stratified-Summer

– Initial conditions influence environments

Methods

• Each Experiment:

– 3 control samples

– 3 samples with 15% additional salinity

– 3 samples with 5% additional salinity

– Chosen based on conditions at effluent sites found in other studies

Methods

– Approx. 1m3 in size for each sample type

– 16 m3 circulating pool

– Pumped from a site 300 m off coast, depth 2 m

– Polyethylene covers on each experiment

• Prevent contamination but allow atmospheric exchange

• Allowed 50% surface light penetration

Methods

– Samples taken from each bag 2 hr after start

– Taken in 5 to 10 L sizes, collected gravitationally

– Collected for next 11 to 12 days

– Parameters measured every day: • Salinity

• Temperature

• Cholorophyll (Chl a)

• Bacterial productivity (BP)

• Primary Productivity (PP)

• DNA for determination of taxa


• Autotrophs saw decrease in photochemical efficiency

• Heterotrophs saw enhanced bacterial activity

– Expected, as salt stress can inhibit photosynthetic reaction centers


• 15% addition experiment:

– PP decreased significantly in spring and summer

– 3.5 to 1.7 ug/L in summer

– 2.9 to 0.9 ug/L in spring

– Attributed to death of salt-sensitive phytoplankton


• 15% addition experiment (for heterotrophs):

– BP increased 2.5x compared to control in spring

– BP increased 1.5x compared to control in summer

– Attributed to need of bacteria to self-regulate due to influx of salt

• Osmotic stress on cells, internal pH, energy potential of membranes


• Community Composition:

– Increased salinity had no significant effect in spring compared to control

– Salinity was dominant driver in summer

• Summer Control: – 51% decrease bacterial diversity

– 252% increase eukaryotic diversity

• Summer 15% Addition: – 87% decrease bacterial diversity

– 70% decrease eukaryotic diversity



– Summer Experiment:

• Cyanobacterial OTUs: – Control: decreased 90%

– 15% addition: increased 140%

• Proteobacterial OTUs: – Control: increased 45%

– 15% addition: decreased 30%



– Summer Experiment:

• Although cyanobacterial OTUs increased by 140% in the 15% addition, Procholorococcus marinus disappeared after first day of treatment

• P. marinus is a major cyanobacterial order

• Contributes 9 to 18% of global marine carbon fixation


• Community Composition: – Summer Experiment:

• Pelagibacter, a heterotroph, declined 6 days later in the 15% addition experiment

• Pelagibacter account for roughly 30% of bacteria in global oceans

• Theorized that Pelagibacter consumes osmolytes produced by P. marinus

• Conclusion: planktonic food webs near effluent sites of desalination plants may shift profoundly in the stratified-summer season of the EMS


• Community Composition: – Summer Experiment:

• Gyrodinium: – Control: relatively low (exact

number unreported)

– 15% addition: Gyrodinium population up 12% compared to other eukaryotic OTUs

– Some species of Gyrodinium known to form red tides

– Increased appearance of Gyrodinium in high salinity environments is of concern at desalination discharge sites


• Final Conclusions:

– Microbial communities have higher functional plasticity to increased salinity in spring than in summer

– Maintaining an abundance of adaptive species can help stabilize an environment that experiences changes in salinity


• Final Conclusions:

– Decrease in biodiversity caused by increase in salinity may destabilize aquatic food web

– Recommendation for future study of other discharges from desalination plants (coagulants, anti-scalants) on microbial communities

Analysis

Strengths

Accurate modeling of real-world

scenario • Rather than random salt concentrations,

based on conditions at effluent sites

• Pulled actual seawater from EMS for experiment

• Microbial community composition seasonally consistent

• Continuous mixing to emulate sea mixing

Professional Statistical Analysis and

Quality Assurance • Analyses of variance (ANOVA)

– Provides a statistical test for multiple means to determine statistical significance

– To determine if a p-value is above significance level

– Probability of obtaining extreme results

– UniFrac Monte Carlo tests for comparing biological communities

Professional Statistical Analysis and Quality Assurance

• Consideration for observational errors

• All parameters tested compared to controls

• Quality assurance performed by national laboratories in US, Canada, & Japan

– Assessed methodology for each parameter determination

Good Explanations of Tangible

Concerns and Link to Study • Connections between change in microbial

environment and known effects – Red tides

– Change in food web

• Linked increases/decreases in microbial taxa to change in salinity – Species with elastic response to change in salt will

thrive

– Some essential species in food web are inelastic / eradicated

Weaknesses

Study Presents other Concerns with

Limited Discussion • Brings up how other substances in discharges

from desal plants may be of concern

• Seems to mention is as a major conclusion from the paper

• It is a topic related to desal plants, but unrelated to effects of changing salinity in water

• Presentation of this concern should have been presented as a side-thought, not among general conclusions

Some Laboratory Methods Under-

Explained • To those unaccustomed to lab techniques,

some detailed were understated

– How do covers on experiment samples both prevent contamination of atmospheric inputs but maintain gas exchange with the atmosphere?

– What effects did freezing the samples have on anticipated sample results?

Limitations

Lack of Comparative Laboratory Studies & Field Measurements

• As desalination processes are a growing practice, research into effects of discharge are not abundant

• Many studies discuss implications of increased salinity, but few actual lab studies

• Few field measurements of impact of effluent discharge

Lack of Comparative Laboratory Studies & Field Measurements

• Hard to determine what to expect and long-term conditions of seawater when field data is limited

• Since study is contained to a 16 m3 zone, effects of a larger boundary zone is unknown

– Would we see the same effects on a microbial community that is not allowed to migrate from a more open boundary?

Unique Findings

Increase in salinity led to immediate decreased photochemical potential

– Decrease in Chl a followed, reflecting death of salinity-sensitive phytoplankton

– Previously known that salinity elevation leads to reduced Chl a in some algae and cyanobacteria

– No previous records found of rapid changes within 2 hours of salt addition

• For 15% addition:

• PP from 3.4 to 1.7 ug/L of Chl a in mixed-spring

• PP from 2.9 to 0.9 ug/L of Chl a in stratified-summer

Knowledge Gaps

Lack of Labs Studies & Field Data

• Increase in BP from salt addition not know to be, but hypothesized to be, from elevated organic consumption bacteria need to adjust to osmotic stress on cells

– Based on findings of a paper from (del Giogrio and Bouiver, 2002)

– Outside of first few hours, salinity elevations did not significantly alter BP production

Effects of losing Pelgibacter and P. marinus are speculative / leading

– Noted that they are responsible for 9 to 18% of marine carbon fixation

– Does not note if other microbes that adjust to highly saline environments that would out-compete these two would be able to replace them

Not clarified if influx of Gyrodinium is

really an issue – Argues that some species of Gyrodinium are

known to form red tides

– Not specified which Gyrodinium species are found in EMS

– Unclear if the ones in this experiment would actually contribute to red tide formation

Questions?

Article: Belkin, N., Rahav, E., Elifantz, H., Kress, N. and Berman-Frank, I. (2015), Enhanced salinities, as a proxy of seawater desalination discharges, impact coastal microbial communities of the eastern Mediterranean Sea. Environmental Microbiology, 17: 4105–4120. doi: 10.1111/1462-2920.12979 Backdrops: Image 1: http://teachmiddleeast.lib.uchicago.edu/foundations/geography/images/geography-05.jpg Image 2: http://m.doosan.com/common/img/intro/status1.jpg Image 3: http://tinyurl.com/ngjn2e4

http://teachmiddleeast.lib.uchicago.edu/foundations/geography/images/geography-05.jpg




http://m.doosan.com/common/img/intro/status1.jpg

http://m.doosan.com/common/img/intro/status1.jpg

http://tinyurl.com/ngjn2e4

http://tinyurl.com/ngjn2e4

Enhanced salinities, as a proxy of seawaterdesalination discharges, impact coastal microbialcommunities of the eastern Mediterranean Sea

Natalia Belkin,1 Eyal Rahav,2 Hila Elifantz,1

Nurit Kress2 and Ilana Berman-Frank1*1Mina and Everard Goodman Faculty of Life Sciences,Bar-Ilan University, Ramat Gan 52900, Israel.2Israel Oceanographic and Limnological Research,National Institute of Oceanography, Haifa 31080, Israel.

Summary

Seawater desalination plants increase local coastalsalinities by discharging concentrated brine back tothe sea with ∼50% higher than ambient salinities. Theimpacts of high salinities on microbial coastal popu-lations of the eastern Mediterranean Sea (EMS) wereexamined in two mesocosm experiments; first, duringthe mixed-spring and second, during the stratified-summer periods with average salinity of ∼39. Ambientsalinities were increased by 5% and 15%. Highersalinity (15%) mesocosms induced rapid (within 2 h)declines in both primary productivity (PP) and algalbiomass parallel to an increase in bacterial produc-tivity. Subsequently, for the duration of the experi-ments (11–12 days), both Chlorophyll a and PP ratesincreased (2 to 5 and 1.5 to 2.5–fold, respectively)relative to unamended controls. The initial assem-blages of the ambient microbial populations andintensity of salinity enrichments influenced the com-munity responses. During the mixed-spring experi-ment, the composition of prokaryotic and eukaryoticpopulations shifted only slightly, suggesting highfunctional plasticity of the initial populations.While during the stratified-summer experiment, highsalinity changed the composition and reduced thebiodiversity of the microbial communities. In an ultra-oligotrophic environment such as the EMS, salinityinduced declines in microbial diversity may provide atipping point destabilizing the local aquatic food web.

Introduction

Large-scale seawater desalination is an effective solu-tion to the freshwater shortage of many countries aroundthe world. Desalination in the Mediterranean Sea com-prises 17% of the world’s total desalination and is one ofthree semi-enclosed basins with intensive desalinationactivity that are anticipated to raise ambient salinities ofthe coastal habitats (Bashitialshaaer et al., 2011). InIsrael alone, seawater desalination by reverse osmosis(SWRO) currently provides ∼450 million m3 (Mm3) y−1

of fresh water along the easternmost Mediterraneancoastline. By 2025, water production supplied by five toseven large-scale coastal plants is forecast to reach750 Mm3 y−1 constituting ∼30% of Israel’s freshwatersupply or c. 80% of the domestic and industrial needs(Dreizin et al., 2008).

The main by-product of all desalination processes is thelarge quantity of concentrated brine that is dischargedback to the marine coastal environment (Ahmad andBaddour, 2014). With typical water recovery rates of40–50% in the desalination process, SWRO plants dis-charge brine to the sea with nearly twice the salt concen-tration of the ambient seawater. This discharge generallyincludes other chemicals used in the process (e.g. coagu-lants, anti-foulants and anti-scalants) (NRC, 2008; UNEP,2008; Spiritos and Lipchin, 2013). Along coastlines, waterloss due to its utilization by desalination plants, combinedwith brine discharge back to the coastal environment,increases the ambient salinity around the outfall areas(Lattemann and Höpner, 2008). The temporal and spatialdispersion pattern of the discharged brine differs amongsites and seasons due to the discharge technology of theplant, changes in local currents and annual physical–chemical characteristics of the water column (Dawoudand Al Mulla, 2012).

The long-term environmental and ecological impacts ofdesalination plants on the marine ecosystem have beenpoorly documented (Roberts et al., 2010). Yet, theelevated salinity at discharge sites, frequently combinedwith the chemicals applied during the desalinationprocess, may impact marine life and water quality aschanges in the physical and/or chemical environment areoften followed by shifts in the composition and production

Received 14 January, 2015; accepted 2 July, 2015. *For correspond-ence. E-mail [email protected]; Tel. (+972)-3-5318214;Fax (+972)-4-6914842.

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Environmental Microbiology (2015) 17(10), 4105–4120 doi:10.1111/1462-2920.12979

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd

mailto:[email protected]

of biological communities. Sensitive coastal environmentsmay especially be prone to alterations in salinity gradi-ents, as observed for the low-salt tolerance Mediterra-nean seagrass Posidonia oceanica and Cymodoceanodosa (Sanchez-Lizaso et al., 2008; Garrote-Morenoet al., 2014). Alterations in biodiversity and succession ofdifferent species may also induce harmful cyanobacterialor algal blooms further affecting coastal water quality(Zingone and Enevoldsen, 2000).

Despite myriad studies discussing the potential foradverse environmental impacts of desalination plant efflu-ents, laboratory studies or field measurements assessingthese impacts are scarce (Roberts et al., 2010; Elimelechand Phillip, 2011; Liu et al., 2013). Moreover, scant infor-mation has been published showing the impacts of desali-nation discharge on the autotrophic and heterotrophiccoastal communities (Drami et al., 2011; van der Merweet al., 2014). Yet, salinity is considered the most importantdriver of global distribution patterns of bacteria as well asother microorganisms (Lozupone and Knight, 2007) andcan regulate functional performance, growth ratesand shifts in bacterial community composition (Bouvierand del Giorgio, 2002; Langenheder et al., 2003). Here,we examined the impacts of increased salinity on thestructure and function of natural assemblages of plank-tonic microbial populations from the coastal Eastern Medi-terranean seawater (EMS) in 1 m3 mesocosms during two11–12 day experiments. To simulate the natural salinityincreases near desalination plant outfalls along the IsraeliMediterranean coastline (Roberts et al., 2010; Dramiet al., 2011; Kress et al., 2011), we enhanced salinities inour experimental mesocosms by 5% and 15% aboveambient salinity (∼41 and 45 respectively). To account forseasonal differences that influence the typical assem-blages of the ambient microbial communities, we imple-

mented two identical experiments: one in early springwhen the water column is fully mixed and one in summerwhen the water column is thermally stratified (termedhereafter: mixed-spring experiment and stratified-summerexperiment respectively).

Results and discussion

Initial state and rapid physiological responses of themicrobial community to salinity

Following the salt additions, the water properties (tem-perature and nutrients) retained the typical seasonalvalues of the EMS surface waters (Table 1). Due to thesalt additions, inorganic nutrient concentrations wereslightly elevated but not significantly different (P > 0.05) inthe treated mesocosms relative to the control mesocosmsat the beginning of the experiment (T0) (elevations up to0.05 μmol L−1 PO4, 0.5 μmol L−1 NO2 + NO3, 0.59 μmol L−1

Si(OH)4) (Table 1).Seasonality imprinted the initial nutrient concentrations

and microbial assemblages of autotrophs andheterotrophs, and these differences affected the subse-quent microbial community responses. At T0, both bacte-rial and eukaryotic representatives of the mixed-springexperiment were more diverse than planktonic communi-ties of the stratified-summer experiment (average effec-tive number of bacterial and eukaryotic species 1715versus 1137; and 553 versus 137 – during mixed-springand stratified-summer experiments respectively). Insummer, the bacterial community was a subset of thespring community (Fig. S1) with a total overlap of 40.2 %between bacterial operational taxonomic unit (OTU) fromthe two seasons. A greater seasonal distinction was foundbetween the diatom and dinoflagellate populations com-prising each experiment with only 13.4% and 16.2%

Table 1. The initial physical, chemical and biological properties as measured on the first day of each of the two mesocosm experiments(mixed-spring and stratified-summer experiments), 2 h after brine additions.

Measured parameter

Mixed-spring experiment Stratified-summer experiment

Ambient 5% 15% Ambient 5% 15%

Salinity 38.8 ± 0.0 40.5 ± 0.1 40.5 ± 0.2 39.3 ± 0.2 41.3 ± 0.4 45.5 ± 0.5Temperature (°C) 22.7 ± 0.0 22.7 ± 0.0 22.7 ± 0.0 29.9 ± 0.2 29.9 ± 0.1 30.0 ± 0.2PO4 (μmol L−1) 0.13 ± 0.01 0.12 ± 0.02 0.18 ± 0.04 0.02 ± 0.00 0.02 ± 0.00 0.05 ± 0.01NO2 + NO3 (μmol L−1) 0.39 ± 0.11 0.55 ± 0.21 0.50 ± 0.06 BDL 0.21 ± 0.12 0.60 ± 0.21Si(OH)4 (μmol L−1) 1.57 ± 0.11 1.69 ± 0.11 1.79 ± 0.09 1.78 ± 0.08 1.86 ± 0.01 2.37 ± 0.04Diversity of bacterial species 1762 ± 513 1656 ± 212 1729 ± 179 939 ± 322 1173 ± 212 1299 ± 122Diversity of eukaryotic species 528 ± 138 480 ± 11 651 ± 31 173 ± 74 108 ± 57 131 ± 19Chlorophyll a (μg L−1) 0.24 ± 0.02 0.12 ± 0.03 0.17 ± 0.01 0.24 ± 0.02 0.25 ± 0.02 0.17 ± 0.01Primary productivity (μg C L−1 h−1) 3.36 ± 0.42 3.89 ± 0.58 1.77 ± 0.24 2.86 ± 0.39 2.74 ± 0.60 0.86 ± 0.15PSII quantum yield (Fv/Fm) 0.12 ± 0.01 0.13 ± 0.02 0.07 ± 0.03 0.13 ± 0.00 0.11 ± 0.01 0.06 ± 0.02Bacterial productivity (μg C L−1 h−1) 0.89 ± 0.61 2.52 ± 0.21 2.26 ± 0.30 2.34 ± 0.37 2.91 ± 0.69 3.26 ± 0.80Bacterial abundance (105 cells ml−1) 6.3 ± 0.7 6.3 ± 0.3 5.7 ± 0.3 4.2 ± 1.1 4.3 ± 1.0 4.3 ± 0.5

All parameters are averages ± SD of three mesocosms per treatment.BDL, below detection limit; Diversity, effective numbers of species that were calculated from Shannon Entropy Index.

4106 N. Belkin et al.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 4105–4120

respective identity in the composition of OTUs betweenthe mixed-spring community and stratified-summer com-munity, with lower read numbers in the stratified-summerexperiment (Fig. S2).

The seasonal difference we detected in the OTU abun-dance and composition of in-situ phytoplankton commu-nities is consistent with published Mediterranean Seadata demonstrating an increase in species diversity withthe annual enrichment of nutrients from winter mixing(reviewed in Siokou-Frangou et al., 2010), which pro-duces, along the Israeli coastline, a community comprisedof diatoms, dinoflagellates (micro-phytoplankton) andnano-phytoplankton (3–20 μm) (Azov, 1986). In thestratified-summer experiment, the phytoplankton diversitywas lower (Table 1), reflecting the typical summer plank-tonic assemblages of the coastal Levantine waters domi-nated by the small (< 3 μm) cyanobacteria and pico-eukaryotic algae (Azov, 1986; Kimor et al., 1987; Herutet al., 2014). Although diversity indices were higher for themixed-spring community than the stratified-summer com-munity (Table 1), the initial algal biomass (Chl a) andprimary productivity (PP) were similar in our experimentalcontrols (ambient mesocosms) for both experiments(0.24 μg Chl a L−1 and 3.15 μg C L−1 h−1, respectively,and Table 1) and may actually reflect the coastal origins ofthe waters that are more uniformly mixed, receive terres-trial and anthropogenic nutrient inputs and depend to alesser extent than offshore populations on up-welled deepnutrients.

In the EMS, heterotrophic bacteria contribute signifi-cantly to the food web structure and may compete suc-cessfully with phytoplankton when nutrients are limited(Thingstad et al. 2005). Moreover, during thermal summerstratification, a heterotrophic microbial food web sup-ported by intensive recycling of organic carbon and nutri-ents dominates the Levantine oligotrophic waters (Tanakaet al., 2007; Ignatiades et al., 2009; reviewed byPulido-Villena et al., 2012). This was reflected in theheterotrophic bacterial productivity (BP) that wasinitially significantly higher in the stratified-summer experi-ment than BP measured in the mixed-spring experiment(2.3 μg C L−1 h−1 vs. 0.9 μg C L−1 h−1 respectively;P < 0.005 – Table 1).

Regardless of the seasonal difference in the initialdiversity and composition, the salinity increases causedan immediate functional response (within 2 h of saltaddition) followed by subsequent changes in both thecomposition and function of the microbial communitiesthroughout the duration of the experiments. Moreover,similar rapid physiological responses were detected inboth sets of experiments. Two hours after the brine addi-tion, flow cytometry and molecular analyses revealed thatthe microbial community structure remained unchanged(Table 1, Table S1). Yet, rapid physiological responses

were recorded that were not a result of dilution. Theseincluded suppression of photosynthesis and decreasedphotochemical efficiency in the autotrophs and enhancedheterotrophic bacterial activity.

Salt stress can inhibit various aspects of the photosyn-thetic process (Sudhir and Murthy, 2004). Changes inphotosynthesis were reflected in the photosystem II(PSII) photochemical efficiency (Fv/Fm) that decreasedsignificantly (P < 0.005) within 2 h in the 15% treatmentsamples (Table 1). Salt stress can inactivate both photo-synthetic reaction centres – PSI and PSII, and inhibit thede novo synthesis of proteins, specifically the synthesisof the D1 protein in PSII (Allakhverdiev et al., 2002). Saltstress can also enhance the oxygenase activity ofribulose 1,5-bisphosphate carboxylase/oxygenase whilecurtailing its carboxylase activity and reducing carbonfixation (Sivakumar et al., 2000). In cyanobacteria,acclimation processes to salt stress can extend from12 to 24 h, during which the cells activate osmolyte syn-thesis (Hagemann, 2011). Our data suggest that theaddition of brine induced an immediate (within 2 h) saltstress on autotrophic microorganisms and suppressedphotosynthesis.

Moreover, the photosynthetic pigment and indirectproxy of algal-biomass (Chl a) decreased within the first2 h (0.24 versus 0.17 μg Chl a L−1, in controls and in the15% treatments, respectively, both in mixed-spring andstratified-summer experiments) (Table 1). Changes in PPparalleled those of Chl a. In the 15% mesocosms, PPdecreased significantly (P < 0.05) after 2 h from 3.4 to1.7 μg C L−1 h−1 in the mixed-spring experiment and from2.9 to 0.9 μg C L−1 h−1 in the stratified-summer experiment(Table 1). The decrease in Chl a probably reflects theimmediate death of salinity-sensitive phytoplanktonspecies (Brand, 1984) following the salt addition. Moreo-ver, salinity elevation reduces cellular Chl a content insome algae and cyanobacteria (McLachlan, 1961) similarto salt-susceptible plants. Yet, we could not find previouspublished records of such rapid changes as we recordedhere.

Higher salinity also induced an immediate responsefrom the heterotrophic populations. Bacterial productionrates increased in the 15% treatments relative to controls(by 2.5 and 1.5-fold in mixed-spring and stratified-summerexperiments respectively) (Table 1). Our results showthat under the experimental conditions, the abundanceof mixed-spring and stratified-summer coastal Mediterra-nean bacterial communities was maintained at theelevated salinities and was in the range of previous reportsin the EMS (Mapelli et al., 2013), while BP increased in thehigher salinity treatments (Table 1 – BAand BP ), indicatinghigher cell specific bacterial activity. The rapid increase inBP may be related to the elevated organic carbon con-sumption required by bacteria to maintain the changing

High salinity impacts coastal microbial populations 4107


osmotic stress of cells and to regulate the internal pH aswell as changing the energetic potential of the cells’ mem-branes (del Giorgio and Bouvier, 2002).

Physiological changes throughout the experiments

Salinity increases produced similar overall responses forboth experiments regardless of the season or initial micro-bial inocula (Chl a; PP and ultra-phytoplankton biomasscomposition) (Fig. 1). This similarity overshadowed somemicrobial responses (e.g. BP) that were seasonallydependent. During the course of each experiment, thesalinity remained unchanged from the initial values, whiletemperature changed (up to 1.34°C) due to daily weatherfluctuations. At the end of the experiments (Tend), due tobiological utilization, inorganic nutrient concentrations(PO4 and NO2 + NO3) were lower than at T0 and similar inall mesocosms (Table S2). Si(OH)4 concentration was sig-nificantly lower than T0 only in the 15% treatments indi-cating changes in functional compositions (discussedbelow).

In both experiments, the elevations in salinity resultedin higher algal biomass (derived from Chl a concentration)

that was indicative of the adaptive capacity of theautotrophic community. The 5% treatment caused a fastand significant increase in the algal biomass after 2 days(Fig. 1A) in both experiments, while autotrophs exposedto 15% salinity lagged for several days before a significantelevation in Chl a concentrations was recorded by day 5(P < 0.001 and P < 0.005 for 5% and 15% respectively;Fig. 1A). Subsequent increases in chlorophyll concentra-tions to values ∼5-fold higher than control mesocosmswere measured at Tend (Fig. 1A).

The increase in autotrophic biomass was especiallypronounced in the ultra-phytoplankton which are impor-tant primary producers in the Levantine basin (Azov,1986; Siokou-Frangou et al., 2010) with small (< 5 μm,pico-phytoplankton) cyanobacteria and larger (>5 μm,nano-phytoplankton) eukaryotic algae dominant alongthe Israeli coast (Herut et al., 2012). By Tend, ultra-phytoplankton biomass increased significantly in the 15%treatments (P < 0.01; Fig. 1B), contributed mostly bylarger (>5 μm) eukaryotic algal cell numbers (Table S1). Inthe 15% higher salinity mesocosms these eukaryotic phy-toplankton were probably diatoms as the stimulation ofalgal biomass was accompanied by silica consumption.

Fig. 1. Temporal changes in: A. Average Chl a concentrations normalized to controls; B. Carbon content of the ultra-phytoplankton communityat the outset and the end of the experiments (*denotes values are significantly different from other treatments P < 0.01); C. average primaryproductivity (PP) normalized to controls. All averages include six biological replicates of mixed-spring and stratified-summer periodexperiments combined ± SE; D. Average bacterial productivity (BP) normalized to controls, averages include three biological replicates ofmixed-spring and stratified-summer period experiments separately ± SE.



At Tend silica concentrations (in the 15% treatmentmesocosms) were 50–90% lower than T0 values and were0.94 μmol L−1 and 0.11 μmol L−1 for the mixed-spring andstratified-summer experiments respectively (Table S2).The differential response in silica uptake is consistent withthe ability of diatoms to survive and thrive at the highersalinity treatments (Kirst, 1989; Schapira et al., 2010) andwith the compositional changes of eukaryotes derivedfrom molecular analysis further described.

Concurrent with biomass changes, PP rates were 1.5-fold higher in the 5% mesocosms throughout most of theincubation period compared with T0 and parallel controlmesocosms (Fig. 1C). The most apparent and significantchange in PP rates was observed in the 15% mesocosms.In these mesocosms, PP rates increased by 1.5 to fivefoldcompared with control mesocosms 5 days after the T0

and were two to threefold higher than controls for thenext 7 days (P < 0.0005; Fig. 1C). In the first 5 days, PPwas significantly lower (by 0.7–0.8) than the controls(P < 0.05), suggesting that the autotrophic communityfunction was initially inhibited by the salinity elevation(Fig. 1C), similar to rates measured 2 h after experimentalinduction (Table 1). These changes were likely caused byosmotic stress that triggered species-specific metabolicchanges followed by structural changes of the communityduring acclimation (Sudhir and Murthy, 2004; Kaartokallioet al., 2005).

The observed lag time of 5 days prior to the enhancedphotosynthetic activity in the 15% treatments (Fig. 1C)may have also been due to the higher energetic demandof the phytoplanktonic community under elevated saltconditions as reported from brackish water communities(Pilkaityte et al., 2004). However, despite the enhancedPP (after 5 days) in the stratified-summer experiment,the PSII photochemical quantum yields (Fv/Fm) of theautotrophic community were significantly lower in the15% treatments than in the control mesocosms(0.09 ± 0.03 versus 0.21 ± 0.07 respectively; P < 0.05).PSII photochemical efficiency decreases when phyto-plankton grow under stress, e.g. nutrient limitation orhigh light (Kolber et al., 1988), indicating that salinity-induced physiological stress caused the decline in Fv/Fm

in the 15% treatments. Similarly, exposure of the greenalga Chlorococcum cells to N-deficiency combined withelevated salinities caused an inhibition of cell divisionand a strong depression of photosynthetic activity(Masojidek et al., 2000).

Excluding the rapid increases in BP measured duringthe first hours of the experiments (discussed above),salinity elevations did not significantly alter bacterialheterotrophic production in most treatments. Seasonalityimpacted the bacterial production potential as significantelevation in bacterial production was measured only forthe mixed-spring experiment exposed to 15% treatments

from day 4 to Tend (P < 0.0005; Fig. 1D). No increase wasmeasured in BP for the stratified-summer experiment. Thecontrasting response in the metabolic signature of bacte-ria between seasons may have occurred due to the dif-fering composition of the initial bacterial communities(Fig. 2) and their functional plasticity.

Temporal changes in community composition

Compositional shifts within the different communitiesderived from ribosomal ribonucleic acid (rRNA) geneswere examined using principal coordinate analysis(PCoA). The distances (calculated from beta diversity)between the beginning and the end (T0 and Tend) of theexperiment for each treatment indicate the extent ofthe shift undergone by the communities (Table 2 andFig. 2). Throughout the mixed-spring experiment, thecomposition of bacterial and eukaryotic groups wassimilar for all treatments and for the control mesocosms(Fig. 2A and B). These results indicate that salinityincreases barely affected the composition of these com-munities and changes were primarily dictated by thetime of sampling.

In contrast, during the stratified-summer experiment,salinity was a dominant driver impacting community com-position as seen by the large change both in bacteria andphototrophic eukaryote composition from the 15%mesocosms (largest distances that significantly differedfrom the controls; P < 0.005). After 6 days (Tm), both bac-terial and eukaryotic communities in the 15% mesocosmsdeviated from the control and 5% mesocosm (Fig. 2C andD). Further analyses demonstrated shifts in both thedirection and magnitude of bacterial and eukaryotic alphadiversity. This significantly changed from T0 to Tend

(Table 2). Thus, over the course of the experiment,diversity of the ambient (controls) bacterial communitydecreased by 51% while that of eukaryotes increased by252% (Table 2). Concurrently, in the high-salinity (15%)mesocosms, both bacterial and eukaryotic diversitydeclined by 87% and 70% respectively (Table 2).

The three major phyla comprising 96–98% of all bacte-rial OTUs from all stratified-summer mesocosms werethe Proteobacteria, Cyanobacteria and Bacteroidetes,which are consistent with earlier reports from the EMS(Feingersch et al., 2010). Significant shifts appearedin the specific composition of proteobacteria andcyanobacteria appeared during the experiment (12 days).High salinity (15% addition) caused a 140% increase inthe relative abundance of cyanobacterial OTUs, whilecyanobacterial OTUs declined by 90% in the controlmesocosms (Table 2). In contrast, the relative abundanceof proteobacterial OTUs increased by 45% in the controlsand were negatively impacted by high salinity (15% addi-tions caused a 30% decline).



The major cyanobacterial order in all mesocosms wasthe Synechococcales (in particular Prochlorococcusmarinus), Oscillatoriophycideae and Nostocales. In the15% treatment, P. marinus disappeared almost com-pletely after 1 day, while the abundance of filamentouscyanobacterial OTUs, specifically Oscillatoriophycideaeand Nostocales, known to adapt to high-salinity conditions(Hagemann, 2011; Jeffries et al., 2012), increased com-pared with the controls (Fig. 3A, Fig. 4).

Among the heterotrophic bacterial groups, thealphaproteobacterium (Pelagibacteraceae) Pelagibacterand the gammaproteobacteria Altermondales declinedsignificantly after 6 days in the high-salinity (15%)mesocosms of the stratified-summer experiment, (Fig. 3B,Fig. 4). These were replaced by Rhodobacterales andspecifically Roseibacterium elongatum and Paracoccusmarinus of the Alphaproteobacteria (Fig. 3B), which areaerobic, chemoheterotrophic, bacteriochlorophyll-containing bacteria (Suzuki et al., 2006) and aerobicbacteria producing the carotenoid adonixanthindiglucoside (Khan et al., 2008). The disappearance of bothPelagibacter (Pelagibacteraceae) and P. marinus from the

Fig. 2. Principal coordinate analysis based on beta diversity of species composition derived from analyses of bacterial 16S rRNA andeukaryotic 18S rRNA genes in mixed-spring [A (16S) and B (18S)] and stratified-summer experiments [C (16S) and D (18S)]; Thecommunities exposed to 15% salinity elevation from the final day of the experiment of each experiment are marked by ellipsoids. T0: 2 h fromthe beginning, T1: 1 day from the beginning, Tm: middle of the experiment, Tend: final day of the experiment.

Table 2. Temporal changes in microbial communities during thestratified-summer experiment.

% Relative change Tend from T0

Salinity addition above control

Treatment Control 5% 15%

16S True diversity −51% 82% −87%Distance (median) 0.32 0.42* 0.59*Cyanobacteria OTUs −90% −45% 140%Proteobacteria OTUs 45% 13% −30%

18S True diversity 252% 156% −70%Distance (median) 0.33 0.39 0.60*Diatom algae OTUs −90% −86% 15%Dinoflagellates OTUs 300% 400% 150%

Relative changes are recorded from the start of the experiment (T0) tothe end (Tend = 11 days) and presented as % change from T0 oftrue diversity for bacteria (16S) and eukaryotic (18S) species; calcu-lated distances between communities are based on beta diversity;and the relative changes (% change Tend from T0) in the relativeabundance of OTUs of the major representative taxa: cyanobacteria,proteobacteria, diatoms and dinoflagellates.*Significant community shift, relative to changes that occurred incontrols (Bonferonni corrected unifrac Monte Carlo significance testP < 0.005).



15% treatments is notable as Pelagibacter comprises∼30% of the bacteria in the global oceans and dominatesthe heterotrophic bacterial community composition (Morriset al., 2002) including the EMS (Feingersch et al., 2010).The cyanobacteria Prochlorococcus contribute globallyfrom 9% to 18% of net primary production, marine carbonfixation and oxygen evolution (Partensky et al., 1999;DuRand et al., 2001; Flombaum et al., 2013). Adaptationsto high-salt environments have been documented in theProchlorococcus genome, potentially encoding forosmolyte production (Scanlan et al., 2009; Klahn et al.,2010). The Pelagibacter genome encodes for transportersof several osmolytes (Giovannoni et al., 2005) along withthe ability to metabolize these compounds (Sun et al.,2011). Pelagibacter may actually consume osmolytes pro-duced by Prochlorococcus and other phytoplankton as asource of energy and nutrients (Thompson et al., 2013).Thus, their sensitivity to high-salinity environments, suchas we observed here after 6 days (Tm), is intriguing andsuggests a profound shift in the planktonic food webs thatmay occur near marine outfalls of desalination plants

during the stratified season of the eastern Mediterranean(May–Dec).

During the stratified-summer experiment, dominantautotrophic eukaryotic groups (diatoms and dinoflagel-lates) changed concurrently with the altered bacterialcommunities (Table 2). The initial populations of theexperiment were consistent with the coastal waters thatare relatively rich in the number of diatom species(Gomez, 2003; Ignatiades et al., 2009; Herut et al., 2012).In the 15% elevated salinity mesocosms, diatom OTUsrelative abundance increased by 15%, parallel to adecrease of 86% to 90% in respective OTUs in the 5%and control mesocosms. Here, the chain-forming diatomLeptocylindrus spp. (50% to 60% of all diatom OTUs)dominated the OTUs throughout the experimental period(Fig. 3C) in contrast with the control mesocosms wheretheir relative OTU abundance declined from ∼50% at T0 to0.2% at Tend (Fig. 5). Leptocylindrus spp. is the predomi-nant diatom in some waters along the eastern andwestern coastlines of the Mediterranean (Ignatiadeset al., 2009; Aktan, 2011; Herut et al., 2011; 2014). At the

Fig. 3. Temporal dynamics in the abundanceof major microbial lineages as obtained by therelative OTU abundance ± SD (of total A, B:bacterial, C, D: eukaryotic OTUs) throughoutthe 11 days of the stratified-summerexperiment for the 15% enhanced-salinitymesocosms. A: cyanobacteria; B:proteobacteria; C: diatoms and D:dinoflagellates.



Fig. 4. Temporal changes in the composition of major bacterial lineages (Cyanobacteria and Proteobacteria) during the stratified-summerexperiment determined by the relative OTU abundance from 16S rRNA analyses at T0 and Tend – after 11 days for the control and 15%enhanced-salinity mesocosms.

Fig. 5. Temporal changes in compositions of major eukaryotic lineages (diatoms and dinoflagellates) during the stratified-summer experimentas determined by the relative OTU abundance from 18S rRNA gene analyses at T0 and Tend – after 11 days for the control and 15%enhanced-salinity mesocosms.



15% salt mesocosms, Leptocylindrus was replaced byincreasing abundance of Minutocellus polymorphus (< 5%at T0 to > 30% of eukaryotic OTUs after 8 days). Thispennate diatom is typical of enclosed and semi-enclosedbasins or estuarine waters, which may at times be heavilyeutrophied, and was previously found to bloom in a Medi-terranean lagoon (Sarno et al., 1993).

Dinoflagellates comprised the second most abundantgroup of eukaryotic phytoplankton in the summer experi-ment with relative OTU abundances increasing by 300–400% in the controls and 5% treatment communities andby 150% in the 15% mesocosms from T0 to Tend (Table 2).The two main genera identified in the 15% mesocosmswere Gymnodinium spp. (1% OTUs at T0) and Gyrodiniumspp. (1% OTUs at T0) (Fig. 3D, Fig. 5). Gyrodinium OTUsincreased to 12% of all eukaryotic OTUs by the end of theexperiment (Fig. 3D, Fig. 5) and dominated the dinoflagel-late OTUs, while in control and 5% mesocosms theirrelative abundance remained low (Fig. 5). Both thesegenera are generally associated with warm and stratifiedwaters (Estrada, 1991), defined as truly phagotrophic, andmay constitute a main part of the microzooplankton (Sherrand Sherr, 2007). Several Gymnodinium and Gyrodiniumspecies form red tides and harmful blooms and areeurohaline (Zingone and Enevoldsen, 2000; Nagasoeet al., 2006). Thus, their resiliency or increases underhigh-salinity conditions should be of special concern whenmonitoring outfall areas.

Our results present two types of responses of the micro-bial community throughout the experiments. The microbialcommunity present during the mixed-spring experimentwent through minimal compositional changes yetincreased its metabolic activity in response to salinityelevations. This suggests a high functional plasticity of aninitially resistant community, while the structural shiftduring the stratified-summer experiment indicates a func-tionally redundant microbial community that was controlledby the salinity changes (Allison and Martiny, 2008). Biodi-versity acts as a buffer against environmental fluctuationsand maintains the stability of ecosystem processes(Tilman, 1999; Loreau et al., 2001). These principlesappear valid also for microbial systems (Bell et al., 2005;Saikaly and Oerther, 2011). Higher diversity increases thechance that some species will be resistant to changes,allows species to compensate for one another and facili-tates processes such as recruitment, thus enhancingrecovery over longer timescales. Maintaining the abun-dance of species with an adaptive capacity, i.e. a combi-nation of phenotypic plasticity, physiological responses,distributional shifts and rapid evolution of traits bettersuited to new conditions (Berga et al., 2012; Bernhardt andLeslie, 2013) can stabilize community function (functionalredundancy) in a varying environment such as that withaltered salinity (McNaughton, 1977; Hooper et al., 2005).

In contrast, a negative correlation between salinity anddiversity may diminish the community resilience to addi-tional disturbances such as increased temperaturesor extreme weather events associated with climatechange (Solan et al., 2004). Thus, decreases in bacterialand phytoplanktonic diversities that can occur at theoutfall locations of desalination plants, and were alsoexemplified in the high-salinity treatments of our stratified-summer experiment (Figs 4 and 5), may reduce the com-munities’ ability to overcome additional stressors.

Conclusions

Here, we demonstrated that salinity conditions similar tothose produced along the EMS coastline by desalinationbrine disposal could be a driving factor shaping thecomposition and function of the microbial planktonic com-munities. We provide evidence for rapid physiologicalresponses (timescale of hours) that may occur even whenthe residence time of plankton at the discharge site isrelatively short. Moreover, while brine discharges mayfluctuate temporally and spatially, their continual and long-term (chronic) input to the coastal areas probably main-tains the phytoplankton and bacterial communities under acontinuous state of salinity stress especially during periodsof low turbulence. Higher chronic salinity may reduce thenumber of species and thus diversity, which would pre-dominately select for high-salinity resilient organisms.Adaptation to salinity fluctuations is crucial for planktonicorganisms and acts as a decisive factor regulating func-tional properties of communities inhabiting a constantlyfluctuating system as found in coastal environments(Brand, 1984). Thus, seasonal changes in composition andflexibility in metabolic responses, as we measured, can‘buffer’ against sudden environmental stress and increaseacclimation over longer periods of time.

Salinity-induced declines in biodiversity of primary andbacterial producers in an ultra-oligotrophic environmentsuch as the Levantine basin may produce a tipping pointand destabilize the local aquatic food web includinggrazers that may be directly impacted by higher salinities(Hart et al., 1998). Furthermore, desalination outflowsalso frequently discharge brine containing coagulants andanti-scalants (i.e. iron salts and polyphosphonates,respectively) that could further modify the responses ofplanktonic communities to the altered salinity gradientsand should also be examined.

Experimental procedures

Mesocosm set-up

Acid washed polyethylene bags (nine bags, each ∼1 m3) sup-ported by cylindrical plastic frames were deployed within acontinuously circulating seawater 16 m3 pool to maintain



ambient temperature and illumination. Surface coastal sea-water was pumped to these mesocosm bags from TelShikmona (Haifa, Israel) at 2 m depth, approximately 300 mfrom the coastline (32°49′34N, 34°57′20E). Bags were filledwith water using plastic tubing by alternating the filling amongthe bags every 2 min to achieve a homogenous distribution.The entire filling procedure lasted for ∼3 h. Polyethylenecovers prevented evaporation or dilution as well as externalcontamination (atmospheric inputs, etc.), yet maintained gasexchange with the atmosphere and allowed for ∼50% ofsurface light to penetrate the water. The mesocosms werelocated at the National Institute of Oceanography of the IsraelOceanographic and Limnological Research IOLR in Haifa,Israel.

Experimental design and sampling

Coastal mixed-spring shallow waters are expected to beaffected by the seasonal mixing processes of the openwaters, reflected in mixed-spring and stratified-summerperiods, when oligotrophic and ultra-oligotrophic conditionsprevail as well as different microbial communities (Kromet al., 1991). These conditions influence the initial inoculaof the ambient microbial planktonic communities. Twomesocosm experiments were carried out: (i) 23 April–5 May2013 and (ii) 7–18 July 2013. These experiments were char-acterized as mixed-spring and stratified-summer experimentrespectively. Both experiments consisted of a control(ambient water – no salinity amendment) and two salinitytreatments (5% and 15% above ambient salinity), simulatingrelative salinity elevations found at nearby desalinationoutfall sites (Roberts et al., 2010; Kress et al., 2011), each intriplicate mesocosms for statistical analyses. The 5% and15% above ambient salinity (39.05 ± 0.35) represented anaverage salinity of 40.90 and 45.25, respectively, in themesocosms. The salinity elevations were achieved by addi-tion of artificial brine (80) prepared by dilution of seawatersalts (NeoMarine, Brightwell Aquatics) in ambient seawater.The salinity was measured with a Yellow Spring InstrumentsYSI 6000 probe (using the Practical Salinity Scale). Toprevent experimental bias due to dilution of the microbialpopulations in the seawater, we prepared the salt additionswith the same seawater used in the mesocosms so that alltreatments and controls contained the same initial microbialassemblage.

The nine mesocosms were filled and sampled 2 h after thebrine addition to fully characterize the initial properties of thewater and the community composition and function (time 0 –T0). At each sampling, the polyethylene covers were openedand 5–10 L water was collected gravitationally from eachmesocosm using acid washed Masterflex Tygon tubing intoacid washed 5 L plastic containers. Subsequent samplingstook place every 1–2 days at 08:00 am for a period of 11–12days for salinity, temperature, chlorophyll (Chl a), bacterialand primary productivity. Inorganic nutrient concentrations,bacterial and ultra-phytoplankton abundance were measuredat the beginning and end of the mesocosm experiments.Deoxyribonucleic acid (DNA) samples for bacterial andeukaryotic composition were sampled on four occasionsthroughout the experiment: days 0, 1, 6 or 7 (middle) and 11or 12 (end), referred as: T0, T1, Tm and Tend respectively. To

minimize biases due to sampling frequency causing a reduc-tion of water levels, the water level within the mesocosmswas retained above 90% of the total volume throughout thewhole experiment (i.e. less than 100 L were taken out of1000 L). To detect the community responses caused onlyby the treatments, we compared all our results to theunamended controls (incubated under the same conditions),reducing the changes that occurred due to the enclosure ofthe community or natural succession (Calvo-Díaz et al.,2011).

Laboratory analyses

Seawater for inorganic nutrient analyses (ortho-phosphate,nitrate + nitrite, nitrite and silicic acid) was sampled into acid-washed scintillation vials. To prevent microbial decompositionof organic matter, the samples were immediately frozen andkept frozen until the day of analysis when they were thawed.Nutrient concentrations were determined using a segmentedflow, Seal Analytical AA-3 System following the methodsdescribed in Kress and Herut (2001). Quality assurance ofthe methods was confirmed by the results of intercomparisonexercises (National Oceanic and Atmospheric Administration(NOAA), USA and the National Research Council ofCanada (NRC), Japan, Quasimeme). The precision of thenitrate + nitrite and nitrite, orthophosphate and silicic acidmeasurements were 0.02, 0.003 and 0.06 μM, respectively,while the limits of detection were 0.08 μM, 0.008 μM and0.03 μM respectively.

Chl a concentrations were determined by the non-acidification method (Welschmeyer, 1994). Mesocosmsamples (250 ml) were vacuum filtered through GF/F 25 mmfilters (Whatman) with a nominal pore size of 0.7 μm. Thepigments were extracted from the filters in 5 ml of 90%acetone, at 4°C, for 24 h in the dark. Chlorophyll a concen-tration was determined using a Luminescence Spectrometer(Trilogy Laboratory Fluorometer, CA) at 436 nm excitationfilter, 680 nm emission filter.

Photosynthetic carbon fixation rates were measured bya modified 14C incorporation method (Steemann-Nielsen,1952). For each mesocosm, we filled quadruplicatepolycarbonate bottles (50 ml; Nalgene) with water duringmorning (∼09:00), inoculated with 5 μCi of NaH14CO3 tracer(Amersham, CFA3), and incubated for 4 h under ambientirradiance and temperature (20.5–23.5°C and 28.5–30°C forspring and summer experiments respectively). After incuba-tion, particulate matter was collected on GF/F filter. The totalradioactivity in this fraction was determined by liquid scintil-lation (Packard Tri carb 2100 TR liquid scintillation analyser)and converted to rates of PP as described by Lagaria andcolleagues (2011) taking into account dark incorporation andzero time controls.

PSII photochemical quantum yields (Fv/Fm) were deter-mined using a Fluorescence Induction and RelaxationSystem (FIRe – Satlantic, Halifax, Canada) to analyse thephotosynthetic characteristics of the autotrophic microorgan-isms (Kolber et al., 1998). After 15 min of dark acclimation,the samples were analysed with a FIRe system set to deliversaturation flash sequences of 100 ms−1 with 1 ms−1 intervalsbetween flashes with the maximum gain (2400) utilized for allsamples. Fluorescence parameters measured were as



follows: Fluorescence F0 intrinsic fluorescence [arbitrary units(a.u.)], the maximum fluorescence (Fm), when all PSII reac-tion centres are photochemically reduced. Based on theseparameters variable fluorescence [Fv = Fm – F0 (a.u.)] wasdetermined. Fv/Fm was calculated after blanks (0.2 um filteredseawater) were subtracted from the initial, dark, adaptedsamples (Cullen and Davis, 2003).

Bacterial production rates were measured using the 3Hleucine incorporation technique (Kirchman et al., 1985).Briefly, 1.7 ml triplicate samples and a control were incubatedwith a mixture of L-[4,5-3H] leucine (Perkin Elmer, specificactivity 160 Ci mmol−1) at a final concentration of 100 nmolLeucine L−1. Samples were incubated in the dark at roomtemperature (∼25 °C), fixed and treated following the micro-centrifugation protocol (Smith and Azam, 1992). Potentialbacterial production rates were calculated using a conversionfactor of 1.5 kg C mol–1 with an isotope dilution factor of 2.0(Simon and Azam, 1989).

Ultra-phytoplankton and bacterial abundances were enu-merated by flow cytometry. Samples of 1.8 ml were immedi-ately fixed after sampling with 5 μl of 50% glutaraldehyde(Sigma G-7651), incubated at room temperature for 10 min,subsequently frozen in liquid nitrogen and kept at −80°C untilanalysis. Prior to the analysis, fixed samples were fastthawed at 37°C. Analysis was performed using a flowcytometer – FACScan Attune, fitted with argon lasers (405and 488 nm). Beads of 1 μm diameter (Polysciences) servedas standards (Marie et al., 2005; Stambler, 2006). The taxo-nomic discrimination was based on cell side scatter – a proxyof cell volume; forward scatter – a proxy of cell size; andorange and red fluorescence of phycoerythrin and Chl a(filters: 574/26 band pass and 640 long pass respectively).Heterotrophic bacteria were stained (300 μl of the initialsample) with SYTO 9 Green Fluorescent Nucleic Acid Stain(Marie et al., 1997) and enumerated by discrimination basedon green fluorescence (530/30 band pass filter) and sidescatter.

Pico/nano phytoplankton carbon biomass was calculatedfrom cell counts assuming 175 fg C cell−1 for Synechococcuscells, 53 fg C cell−1 for Prochlorochococcus cells and2100 fg C cell−1 for nanoeukaryotes (Campbell, 2001).

Statistical analyses

Prior to the statistical comparison, data were binned into twogroups: T0–T5 and T6–Tend, to reduce the effects of minorobservation errors. To assess treatment-dependent signifi-cant changes for each physiological parameter one-wayanalyses of variance (ANOVAs) with permutations were per-formed for each group using the R software package(lmPerm, ver. 2.15). The permutation test allowed applyingANOVA with no assumptions about the data set normality orhomogeneity of variance and corrected for the multiple com-parisons. For comparison of each parameter between differ-ent treatments, statistical analysis was done by post-hocTukey honestly significant difference test after ANOVA test.

DNA isolation and high-throughputphylogenetic analyses

To analyse the microbial community diversity by 16S and 18SrRNA genes, water samples (2 L) were filtered on 47 mm

0.2 μm pore size Supor filters (Pall Gelman, Ann Arbor, MI),frozen in liquid nitrogen and kept at −80°C until DNA extrac-tion. Community genomic nucleic acids were isolated fromthe filters, and media using a phenol–chloroform extractionmethod modified according to Massana and colleagues(1997) and Brinkhoff and Muyzer (1997). For initial amplifica-tion, the broadly conserved bacterial primers 27F and 1100Rwere used to amplify the 16S rRNA gene region (Lane, 1991;Dowd et al., 2008), and eukaryotic primers 360F and 1492Rfor 18S rRNA gene region (Edgcomb et al., 2011). ThermoScientific Phusion high-fidelity DNA polymerase was used toamplify these segments. A secondary polymerase chain reac-tion (PCR) was performed for next-generation sequencing(Ion Torrent Life Technologies, USA) using specially designedfusion primers with different tag sequences as: LinkerA-Tags-27F and LinkerB-338R for 16S (Hamady et al., 2008);LinkerA-Tags-528F and LinkerB-706R for 18S rRNA gene(Cheung et al., 2010). Polymerase chain reactions were per-formed under the following conditions: 95°C for 3 min fol-lowed by 25 cycles and 20 cycles (first and secondary PCR,respectively) of 95°C for 30 s; 60°C for 30 s and 72°C for1 min and a final elongation step at 72°C for 5 min. Aftersecondary PCR, all amplicon products were purified usingAgencourt Ampure magnetic purification beads (AgencourtBioscience Corporation, MA, USA) to exclude primer dimers.Products of DNA for the 16S and 18S fractions weresequenced to get a representative view of the bacterial andeukaryotic community composition.

Sequence analyses. Sequences were processed and ana-lysed using ‘quantitative insights into microbial ecology’ (QIIME)an open-source software pipeline (Caporaso et al., 2010).Sequences were removed if they were < 200 or > 400 bp, hada quality score of < 25, contained ambiguous characters or anuncorrectable bar code or did not contain the primersequence. Remaining sequences were assigned to samplesby examining the bar codes. Mean number of sequences persample passing quality filters were 2823 and 5919 withaverage sequence length 265 bp and 255 bp for bacteria andeukaryotes respectively. Similar sequences were clusteredinto OTUs using UCLUST (Edgar, 2010) with a minimum cov-erage of 99% and a minimum identity of 97%. A representativesequence was chosen from each OTU then was aligned usingPYNAST (Caporaso et al., 2010), the Greengenes (DeSantiset al., 2006) and Silva databases (bacteria and eukaryota,respectively) with a minimum identity of 80%. Chimera check-ing was applied to remove chimera sequences, followed byLane mask to screen out hypervariable regions after align-ment. Taxonomy was assigned using the Ribosomal DatabaseProject (RDP) [Michigan State University] classifier with aminimum support threshold of 80% (Wang et al., 2007) and theRDP taxonomic nomenclature.

Quantifying and comparing diversity. Several metrics wereapplied to test the communities’ compositional shifts in themesocosms. All parameters tested were compared with thecontrol communities. To evaluate the diversity within commu-nities (Alpha diversity), we employed rarefaction plots andbranch length-based phylogenetic diversity measurements(Faith, 1992). Effective number of species (referred to asdiversity) were calculated from Shannon index of entropy (H)



according to Jost (2007) using the conversion equation:exp(H), to examine and compare the diversity of each com-munity from different time points during the incubation period(Moreno and Rodríguez, 2011). To determine the amount ofdiversity shared between two communities (pairwise sampledissimilarity – beta diversity), we employed the weightedUNIFRAC metric (Lozupone and Knight, 2005; Lozupone andKnight, 2007; Lozupone et al., 2007; Lozupone et al., 2011).To control for sampling effort in beta diversity measurements,rarefaction and jackknifing analysis were applied (Lozuponeet al., 2006). We performed significance tests and PCoAusing UNIFRAC (Lozupone and Knight, 2005). To furtherunderstand the significant shifts, we calculated relativechanges in diversity of each treatment normalized to theinitial diversities. In addition, we calculated the extent of thesecompositional shifts by testing the distances measured bybeta diversity between the beginning and the end of theincubation for each treatment and tested which of the shiftswas significantly different from others (when P < 0.05) usingthe UNIFRAC Monte Carlo significance test. Finally, we exam-ined the OTUs of major representative taxa that changedtheir abundance significantly (when P < 0.05) throughout theincubation using Bonferroni corrected ANOVA and calculatedthese relative changes.

Acknowledgements

We acknowledge the Israel Water Authority grant number4500445459 for partial funding to Ilana Berman-Frank (IBF).We thank Dan Miller for technical help and samplingsduring the experiments. This research is part of the PhDrequirements for Natalia Belkin from Bar Ilan University(BIU). Natalia Belkin (NB) was supported by a President’sFellowship from BIU and The National Fellowship GraduateProgram for Marine Conservation in the Mediterranean.

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Supporting information

Additional Supporting Information may be found in the onlineversion of this article at the publisher’s web-site:

Fig. S1. Weighted Venn diagram based on 16S rRNA genesequences comparing the initial bacterial composition (asdetermined by OTUs) in control samples from T0 in mixed-spring (red) experiment to samples from stratified-summer(blue) experiment. The overlap of weighted Venn circlesreflects the OTU reads originating from the same lineage.Circle sizes are proportional to the relative amount of eachsample’s OTUs in the data set. The accompanying tabledetails the percent of OTUs contributed by each season’sdata set, out of all compared OTUs per lineage (100%) andthe overlap of OTUs shared between the two seasons, aswas calculated for the comparative weighted Venn diagram.The diagram was created using CoVennTree methoddescribed by Lott et al. (2015).Fig. S2. Weighted Venn diagram based on 18S rRNAgene sequences comparing the initial eukaryotic phyto-plankton composition (as determined by OTUs) in controlsamples from T0 in mixed-spring (red) to samples fromstratified-summer (blue) experiments. The overlap ofweighted Venn circles reflects the OTU reads originatingfrom the same lineage. Circle sizes are proportional to therelative amount of each sample’s OTUs in the data set. Theaccompanying table details the percent of OTUs contributedby each season’s data set, out of all compared OTUs perlineage (100%) and the overlap of OTUs shared betweenthe two seasons, as was calculated for comparativeweighted Venn diagram. The diagram was created usingCoVennTree method described by Lott et al. (2015).



Table S1. Changes in the abundance of ultra-phytoplankton(Synechococcus, Prochlorochococcus and eukaryotic algae)as obtained by flow cytometry. Detailed data are average cellcounts ± SD from triplicate mesocosms of each treatment atthe beginning (T0) and the end (after 11 and 12 days = Tend)of the mixed-spring and stratified-summer experiments.Table S2. Temporal changes in the average inorganicnutrient concentrations ± SD measured from triplicate

mesocosms of each treatment at the beginning (T0) and theend (after 11 and 12 days = Tend) of the mixed-spring andstratified-summer experiments.Supporting Information Reference. Lott, S.C., Vob, B.,Hess, W.R., and Steglich, C. (2015) CoVennTree: a newmethod for the comparative analysis of large datasets. FrontGenet 6: 1–8.



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