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Review

Evidence-Based Framework to Manage Cyanobacteria and Cyanotoxins in Water and Sludge from Drinking Water Treatment Plants

by
Farhad Jalili
1,
Saber Moradinejad
1,*,
Arash Zamyadi
2,
Sarah Dorner
1,
Sébastien Sauvé
3 and
Michèle Prévost
1
1
Department of Civil, Mineral and Mining Engineering, Polytechnique Montréal, Montréal, QC H3C 3A7, Canada
2
Faculty of Engineering and Information Technology, University of Melbourne, Melbourne, VIC 3010, Australia
3
Department of Chemistry, University of Montréal, Montréal, QC H3C 3J7, Canada
*
Author to whom correspondence should be addressed.
Toxins 2022, 14(6), 410; https://doi.org/10.3390/toxins14060410
Submission received: 9 May 2022 / Revised: 12 June 2022 / Accepted: 13 June 2022 / Published: 15 June 2022
(This article belongs to the Special Issue Management of Cyanobacteria and Cyanotoxins in Waters)
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Abstract

:
Freshwater bodies and, consequently, drinking water treatment plants (DWTPs) sources are increasingly facing toxic cyanobacterial blooms. Even though conventional treatment processes including coagulation, flocculation, sedimentation, and filtration can control cyanobacteria and cell-bound cyanotoxins, these processes may encounter challenges such as inefficient removal of dissolved metabolites and cyanobacterial cell breakthrough. Furthermore, conventional treatment processes may lead to the accumulation of cyanobacteria cells and cyanotoxins in sludge. Pre-oxidation can enhance coagulation efficiency as it provides the first barrier against cyanobacteria and cyanotoxins and it decreases cell accumulation in DWTP sludge. This critical review aims to: (i) evaluate the state of the science of cyanobacteria and cyanotoxin management throughout DWTPs, as well as their associated sludge, and (ii) develop a decision framework to manage cyanobacteria and cyanotoxins in DWTPs and sludge. The review identified that lab-cultured-based pre-oxidation studies may not represent the real bloom pre-oxidation efficacy. Moreover, the application of a common exposure unit CT (residual concentration × contact time) provides a proper understanding of cyanobacteria pre-oxidation efficiency. Recently, reported challenges on cyanobacterial survival and growth in sludge alongside the cell lysis and cyanotoxin release raised health and technical concerns with regards to sludge storage and sludge supernatant recycling to the head of DWTPs. According to the review, oxidation has not been identified as a feasible option to handle cyanobacterial-laden sludge due to low cell and cyanotoxin removal efficacy. Based on the reviewed literature, a decision framework is proposed to manage cyanobacteria and cyanotoxins and their associated sludge in DWTPs.
Keywords:
cyanobacteria; cyanotoxins; pre-oxidation; sludge; accumulation; management; water treatment plant
Key Contribution: Cyanobacteria and cyanotoxins management in DWTPs is a triangular activity that includes monitoring, treatment, and sludge handling and interrelation.

1. Introduction

A cyanobacterial bloom occurrence may result in metabolite (cyanotoxins and taste and odor agents) production and release, which is considered a widespread problem in drinking water resources around the world [1,2,3,4,5,6,7,8,9,10].
Conventional treatment processes, including coagulation, flocculation, sedimentation, and filtration, are widely applied to remove cyanobacterial cells and cell-bound cyanotoxins [11,12,13,14]. However, conventional treatment processes may not be able to remove dissolved metabolites (e.g., cyanotoxins) efficiently [12,15,16,17,18,19,20,21,22]. Moreover, toxic cyanobacterial breakthrough has been reported in the effluent of conventional treatment processes and even after post-oxidation [18]. Therefore, additional treatment such as oxidation or powdered (PAC)/granular activated carbon (GAC) may be required to control dissolved metabolites [23,24,25,26].
Pre-oxidation enhances cyanobacteria cell removal during the coagulation/sedimentation process [27,28,29,30,31,32,33,34] and may decrease the cell breakthrough potential from the downflow processes. However, it is reported that pre-oxidation may cause cyanobacteria cell damage (decrease in cell viability) and cell-bound cyanotoxin release [35,36,37]. The level of cell lysis/damage and cyanotoxin degradation/release following pre-oxidation depends on the oxidation exposure (CT as residual concentration x contact time), and it is the driver to find the best pre-oxidation practice against cyanobacteria and cyanotoxins [38,39].
Furthermore, conventional treatment processes cause cell accumulation in drinking water treatment plants’ (DWTPs) sludge, even in DWTPs with low cyanobacterial cell numbers in the intake water [13,18,40,41,42,43,44]. Several studies have demonstrated that cyanobacterial cells could survive in the stored sludge and release cyanotoxins for up to 12 days [13,40,45,46,47,48,49,50,51,52]. Recent studies revealed a new challenge on the probability of extended survival time and even cyanobacterial growth during sludge storage [53,54,55]. Thus, recycling the supernatant of stored cyanobacteria-laden sludge to the head of the DWTPs can increase health-related concerns [55,56]. Such challenges highlight the importance of the treatment and management of cyanobacteria-laden sludge [57,58,59].
The objectives of this study are to: (1) critically review shreds of evidence of the pre-oxidation impact on the cultured-based and natural bloom studies, (2) perform a critical review of the fate of cyanobacteria and cyanotoxins in conventional treatment plants’ sludge and during sludge storage, and (3) develop an operational decision framework to determine the best practice to minimize risks associated with cyanobacteria and cyanotoxin presence in DWTPs.
This critical review provides insight into the fate of cyanobacteria and their associated metabolites throughout DWTPs and their sludge; furthermore, a practical decision framework to mitigate health and operational risks is developed.

2. Impact of Conventional Treatment on Cyanobacteria and Cyanotoxin Accumulation in Sludge

Different studies have reported that conventional treatment processes can remove 62–99% of the cyanobacterial cells in DWTPs [13,18,40,41,42,56]. It has been demonstrated that potential toxic cyanobacterial cells such as Microcystis, Dolichospermum, and Aphanocapsa can be removed using conventional processes [12,56,60,61].
The long-term monitoring of a high-risk DWTP (Lake Champlain—Quebec) during cyanobacterial bloom seasons from 2008 to 2011 showed an extreme accumulation of cyanobacteria cells (up to 107 cells/mL) and cyanotoxins (up to 60 μg/L microcystin-LR (MC-LR)) in the sludge of the clarifier [12,18]. Monitoring of the same DWTP in 2017 showed that cyanobacteria cell accumulation in the sludge holding tank was up to 31-fold higher than taxonomic cell counts in the intake water [56]. An investigation of four DWTPs in the Great Lakes (Ontario) with low cyanobacterial cell influx (<1000 cells/mL) revealed that cyanobacterial cells and cyanotoxins accumulated in the sludge by up to 100 and 12 times higher than the raw water, respectively [44]. Zamyadi, et al. [57] reported a 406% and 2600% cell count increase in the thickened and centrifuged sludge, respectively, in a DWTP equipped with dissolved air flotation (DAF). A similar high accumulation was also reported for the backwash of the direct filtration process [45].
Pre-oxidation may decrease the risk of cyanobacterial accumulation in the clarifier and sludge [43]. Two DWTPs (the same source for intake water with low cyanobacteria cells; maximum < 500 cells/mL) with chemically enhanced conventional treatment processes were studied, but only one of the DWTPs was equipped by pre-ozonation [43]. Pre-ozonation (initial concentration: 0.3–0.8 mg/L, contact time: 6.3 min) decreased cell accumulation in the surface of the clarifier and filters by up to 1450 times as compared to the DWTP without pre-ozonation. Accordingly, an up to 7 times lower cell accumulation was observed in the sludge of the DWTP with pre-ozonation (Figure 1).
In many DWTPs, the supernatant of the stored sludge is recycled to the head of the plant as the spent filter backwash water [45,57,62]. A full-scale study on a low-risk DWTP (3400 cells/mL at the intake water) documented that cyanobacterial cell counts in intake water increased by up to 43% after recycling the supernatant. Surprisingly, 80% of the transferred cells from the supernatant water were viable [60]. A recent laboratory investigation on intake water that contained 1 × 106 cells/mL of cultured M. aeruginosa reported that although conventional treatment maintained the treated effluent parameters at below WHO and USEPA guidelines, recycling of the sludge supernatant resulted in an additional increase in cells and cyanotoxins levels in the influent by up to 7 × 104 cells/mL and 0.26 µg/L MC-LR, respectively [63].

3. Pre-Oxidation Impact on Cyanobacteria Cells, Viability, and Cyanotoxins

3.1. Impact of Pre-Oxidation on Cyanobacteria Cell Counts

Cyanobacterial entry into DWTPs can be dampened by using pre-oxidation. Pre-oxidation may cause cell lysis, damage, and cyanotoxin release and degradation. Several studies were conducted to evaluate the pre-oxidation impact on cyanobacteria (e.g., cell viability and lysis) and cyanotoxins (release and degradation). A recent study tried to map the treatment barriers against cyanobacteria cells and cyanotoxins in drinking water facilities [10]. The results showed that the efficiency of the multi-barrier approach depends on the species present, metabolite concentration, and pre-oxidation dose [10]. Table 1, Table 2, Table 3 and Table 4 summarize the literature on the impact of pre-oxidation on cyanobacteria (cultured-based and natural blooms) for four common oxidants (chlorine, ozone, potassium permanganate, and hydrogen peroxide). Table 1, Table 2, Table 3 and Table 4 show that CT (residual concentration × contact time) is a main driver of cyanobacterial cell lysis, damage, cyanotoxin release, and degradation.
Although some studies have reported a reduction of more than 90% in taxonomic cell counts following pre-oxidation in the lab-cultured cells, Zamyadi, et al. [35] reported a cell reduction of 70% at high chlorine exposure (CT 296 mg min/L). Fan, et al. [37] showed a limited impact of chlorine exposure (CT 104 mg min/L) on the taxonomic cell counts of Microcystis aeruginosa (logarithmic phase). These observations might be related to the cyanobacteria stage of life and agglomeration. Furthermore, comparing the taxonomic cell count percentage in cultured-based and natural bloom studies demonstrates the lower impact of pre-oxidation during natural blooms. Figure 2 exhibits a lower impact of pre-ozonation (2 mg/L) on cell number reduction in a natural bloom in comparison with lab-cultured cyanobacteria.
Table 1, Table 2, Table 3 and Table 4 and Figure 2 show that pre-oxidation, even at high CTs, may not be able to cause complete cell lysis. Consequently, it is important to clarify how far the pre-oxidation can cause viability loss and cyanotoxin release.
Table
Table 1. Summary of the literature on the impact of pre-chlorination on cyanobacteria (cultured and natural blooms). HV: high viability, LV: low viability, DV: development stage, MA: maintenance stage.
Table 1. Summary of the literature on the impact of pre-chlorination on cyanobacteria (cultured and natural blooms). HV: high viability, LV: low viability, DV: development stage, MA: maintenance stage.
Dominant Cyanobacteria
(Cell Density)		Lab/FieldCl2 Dose (mg/L)Contact Time (min)CT (mg min/L)Cell Count Reduction %Cell Viability %ToxinsReferenceComment
Microcystis
(2 × 106 cells/mL)		Lab1–2-min. 15
max. 90		-min. 83
max. 18.4		99% degradation[66]Saline solution; exact dose and contact time were not provided; no residual; CT evaluation weak; no cell-bound
D. circinalis
(46,000 cells/mL)		Lab2
3		0–60min. 1.8
max. 50		-min. 15%
0 for CT 5.8		>100% release (CT 5.8)
>90 degradation (CT 50)		[67]River water; using fluorescein diacetate (FDA) for viability
Microcystis
(6 × 104 cells/mL
(2.5 × 105 cells/mL)
(5 × 105 cells/mL)		Lab2
4.5
10		0–60min. 3
max. 296.1		max. 76%->100% release (CT 5)
>90 degradation (CT 35)		[35]River water, ultrapure water; no viability was reported
Microcystis
(7 × 105 cells/mL)		Lab3, 4, 51, 2, 5, 10, 20, 30, 60min. 2.8
max. 104		Limited impact<5% (CT 4)25% degradation (CT 2.8)
Complete degradation (CT 104)		[37,68]Ultrapure water
Microcystis
(2 × 106 cells/mL)		Lab0.5
0.7
1.5		5, 11, 50, 60, 120min. 2.5
max 180		-<5% (CT 180)10% degradation
40% increase in released		[69]Lake water; no CT reported
Microcystis
(106 cells/mL)		Lab0.2, 0.4, 0.8Range 0–480min. 12
max. 396		-18% (at CT 12)
0.1% (at CT 396)		-[70]Lake water; no CT reported; no cell count; no toxin
Microcystis
(106 cells/mL)		Lab1, 2, 4, 81, 2, 4, 8, 16, 32, 60HV
min. 0.98
max. 361
LV
min. 0.98
max 200		-HV
95–0% (CT > 15)
LV
44–0% (CT > 15)		HV
CT↑—degradation↑
Complete (CT 108)
CT↑—degradation↑
> 50% release CT > 7
> 62% degradation at highest CT		[71]Ultrapure water; two viability range
Microcystis
(1 × 106 cells/mL)
(2 × 106 cells/mL)		Lab1, 2, 4, 81, 2, 4, 8, 16, 32, 60DV
min. 3.8
max 356
MA
min 3.7
max 293		>95% reduction (CT > 13.3)
>95% reduction (CT > 11.9)		No cell viability after oxidationSame as cell death[72]Ultrapure water; two stage of life
Microcystis-Colony
(105 cells/mL)		Lab0.3, 0.5, 1, 2Range 0–20 minmin. 0.97
max. 52		-Depends on colony size
(0–95%)		Release and degradation Colony-size-dependent[73]Lake water; different colony size; no cell count
Natural bloomFieldCl2/DOC: 0.05–3.60–20 minmin. 0.15
max 6.8		>80% increase (CT 6.8)88% reduction Complete release CT:4 (Cl2/DOC: 0.3)[74]No CT provided; CT estimated; Chl-a measured as cell damage surrogate
Natural bloom
US: (3 × 106 cells/mL) - Planktothrix agardhii
Canada: (3 × 105 cells/mL) -
D. spiroides		FieldCl2/DOC:
0.05, 025, 0.15, 0.1, 1		0–20 minUS
min 0.13
max 15
CA
min 0.3 max 21		-Complete degradationComplete degradation
CT 11 (US), CT 7.5 (CA)		[75]No cell viability; no cell count; Chl-a measured as cell damage surrogate
Natural bloom
(3.3 × 105 cells/mL)
D. spiroides
(5.4 × 104 cells/mL)
M. aeruginosa		Field0.2, 0.60–120 minmin 0.15
max 3.84		min. CT 5% decrease
max. CT 34% decrease		min CT: 82%
max CT:55%		CT 3.84: 23% decrease[76]Soft chlorination (low dose)
Natural bloomField2, 50–60 minmin 1.14
max 14.8		min. < 5% reduction
max. > 50% reduction		-2 mg, CT 10, >200% release
5 mg, CT 20, >200% release		[77]No cell viability
Table
Table 2. Summary of the literature on the impact of pre-ozonation on the cyanobacteria (cultured and natural bloom).
Table 2. Summary of the literature on the impact of pre-ozonation on the cyanobacteria (cultured and natural bloom).
Dominant Cyanobacteria
(Cell Density)		Lab/FieldO3 Dose (mg/L)Contact Time (min)CT (mg min/L)Cell Count Reduction %Cell Viability %
(for CT)		ToxinsReferenceComment
Microcystis
(2 × 106 cells/mL)		Lab1
2		-min. 12
max 16		-CT > 54, complete lossCT = 12 complete
degradation		[66]Saline solution; exact dose and contact time were not provided; no residual; CT evaluation weak
Microcystis
(7 × 105 cells/mL)		Lab2, 4, 65min. < 0.22
max. 2.29		-Min CT: 50%
Max CT: 8.5%		>100% release (high CT)
50% degradation		[37,68]Ultrapure water
Microcystis
D. flos-Aquae
(2.5 × 104 cells/mL
(1.5 × 105 cells/mL)		Lab0.5, 2, 40.5–10min. < 0.2
max. 22		32% for 2 mg/L
41% for 4 mg/L		Complete loss,
CT < 0.2		-[64]Ultrapure; no flow cytometry
Microcystis
(2 × 105 cells/mL)
Oscillatoria
(2800 cells/mL)
Lyngbya sp.
(1600 cells/mL)		Lab0.63–524 hmin. 0.5
max 17		100% reduction (CT 0.5)Complete loss,
(CT > 2)		-[78]River water; Chl-a measured as cell damage surrogate; no toxin measurement
Microcystis, Dolichospermum
(4 × 105 cells/mL)		Lab0.5, 1, 25, 10max. 2.5>95% reductionComplete loss-[79]Natural water; no toxin measurement
Microcystis, Dolichospermum
(1.2 × 105–2 × 106 cells/mL)		Field2, 3, 4, 50–10min. 1.4
max 16.8		75% reduction (CT 16.8)CT 3.2: 45%
CT 16.8: 15%		CT < 2, more than 100% release[65]Natural bloom
Natural bloom
US (3 × 106 cells/mL)—Planktothrix agardhii
CA (3 × 105 cells/mL)—
D. spiroides		FieldO3/DOC:
0.05—0.75		0–20US-min. 1.5
max. 3
CA-min 0.2
max. 4.1		-->80% degradation
CT 4.1(CA)		[75]No cell viability; no cell count; Chl-a measured as cell damage surrogate
Natural bloom
(3.3 × 105 cells/mL)
D. spiroides
(5.4 × 104 cells/mL)
M. aeruginosa		Field0.1, 0.30–10max: 0.86max CT 14% decreasemax CT: 79%14% degradation
No release		[76]Soft ozonation (low dose)
Table
Table 3. Summary of the literature on the impact of potassium permanganate on cyanobacteria (cultured and natural bloom).
Table 3. Summary of the literature on the impact of potassium permanganate on cyanobacteria (cultured and natural bloom).
Dominant Cyanobacteria (Cell Density)Lab/FieldKMnO4 Dose (mg/L)Contact Time (h)CT (mg min/L)Cell Count Reduction %Cell Viability %
(for CT)		ToxinsReferenceComment
Microcystis
(2 × 106 cells/mL)		Lab1–2-min. 15
max. 600		-min. CT: 60%,
CT > 60: complete loss		CT: 30 Complete dissolved degradation[66]Saline solution; exact dose and contact time were not provided; no residual; CT evaluation weak
Microcystis
(7 × 105 cells/mL)		Lab1, 5, 100.25–7min. 28.7
max. 2642		14% cell number reduction (CT max)CT 2600: complete lossRelease at CT > 70
Complete degradation CT 2600		[37,68]Ultrapure water
Microcystis, Dolichospermum
(4 × 105 cells/mL)		Lab2, 520max. 45610% reduction at highest CTCT 456: 18% viability-[80]Natural water; no toxin measurement
Microcystis
Bloom from Lake Erie		Lab
Field		0.5–81–5min. 120
max. 1920		-Cell, CT 1920: 2%
Bloom, CT 1920: 40%		-[81]No cell count and toxin; no CT; CT with lower doses was unable to decrease viability
Table
Table 4. Summary of the literature on the impact of hydrogen peroxide on cyanobacteria (cultured and natural bloom). h: hour, d: day.
Table 4. Summary of the literature on the impact of hydrogen peroxide on cyanobacteria (cultured and natural bloom). h: hour, d: day.
Dominant
Cyanobacteria (Cell Density)		Lab/FieldH2O2 Dose (mg/L)Contact TimeCT (mg h/L)Cell Count Reduction %Cell Viability %
(for CT)		ToxinsReferenceComment
Microcystis
(3.7 × 106 cells/mL)		Lab3.4, 174 h, 2 d, 4 dmin. 13.6
max. 1632		min. CT: 8% reduction
max. CT: 89% reduction		K+ release
min. CT: 81%
max. CT: 5%		CT > 816
26% MC release		[82]K release as a surrogate for cell damage; no CT provided
Microcystis
(7 × 105 cells/mL)		Lab10.2, 51, 1020.1 d–7 hmin. 189.3
max. 17,678		Limited changemin. CT: 86%
CT 4770: 7%		No release,
CT 364: >95% degradation		[37,68]Ultrapure water
Pseudanabaena
(107 cells/mL)		Lab3, 5, 10, 202 h, 4 h, 8 h, 2 d, 4 dmin. 6
max. 960		min. CT: No change
max. CT: >90% reduction		CT 120: 2%-[83]Reservoir water; no toxins
Microcystis
(6 × 106 cells/mL)		Lab1–150.1 d–7 dmin. 2.4
max. 2520		CT 1680: 95% reductionmax. CT
3% viability		CT > 1512,
82% degradation		[84]Culture; no CT provided
Microcystis, Dolichospermum
(4 × 105 cells/mL)		Lab5, 106 hmin. 13.9
max. 96.1		<5% reductionmin. CT: 39%
max. CT: 30%		-[79]Natural water
Natural bloom:
(3.3 × 105 Cells/mL)
D. spiroides:
(5.43 × 104 cells/mL)
M. aeruginosa		Field106 h–1 dmin. 47
max. 140.7		max. CT 52% reductionmin. CT: 60%
max. CT: 40%		No release
max.; 15% MC degradation		[76]-

3.2. Chlorination

Figure 3 is a reconstructed graph from the cell viability results following pre-chlorination based on the oxidant exposure (CT). Parameters such as background water quality (e.g., pH and dissolved organic carbon (DOC)) which have an impact on the oxidant demand are included in the CT concept. Therefore, a comparable level of damage should be found by comparing cell viability results using oxidant exposure (CT) for lab-cultured and natural blooms. Figure 3 demonstrates that at the same level of chlorine exposure, the natural bloom is more resistant to pre-chlorination as compared to lab-cultured cells. In other words, lab-cultured studies are not representative of natural bloom pre-chlorination. Fan, et al. [73] reported that the level of cell damage and toxin release depends on the colony size. Figure 3b shows the cultured-based studies fitted with the Chick–Watson equation. Although the results from different unicellular studies are aligned with each other, the colonial Microcystis chlorination shows a different cell damage rate. This could be related to the agglomeration of cyanobacteria cells and increasing the mucilage sheath in colonial cyanobacteria [73]. Despite using the CT calculation to compare the results, Figure 3c demonstrates different cell damage rates for each study of real bloom after chlorination, and the same level of chlorine exposure may not result in the same level of cell damage. Figure 3c shows that cyanobacterial bloom oxidation could be site- and bloom-specific, depending on the agglomeration, cyanobacteria (bloom) stage of life, and metabolic functions. Higher cell damage following pre-oxidation (especially with higher CTs) can lead to higher cyanotoxin release, which cannot be removed during conventional treatment. Soft chlorination showed cell damage by up to 45% and total microcystin (MC) degraded by up to 23%, while no cyanotoxin release was observed [76]. In addition, soft chlorination may cause lower disinfection by-products as a lower oxidant concentration is used in this approach.

3.3. Ozonation

Figure 4 shows the impact of pre-ozonation on cyanobacteria cell damage for cultured-based and natural bloom studies. Figure 4a demonstrates lower cyanobacteria cell damage for a specific ozone exposure for natural blooms as compared to the lab-cultured cyanobacteria. The model fit results (Figure 4b) show a higher cell damage rate for the lab-cultured cyanobacteria in comparison to the natural bloom. As per soft oxidation, soft pre-ozonation was reported to cause up to 21% of cell damage and 14% of MC degradation, while no MC release was observed simultaneously [76]. Such an observation implies the effectiveness of soft pre-ozonation to damage the cells without cyanotoxin release.

3.4. Potassium Permengeanate

Figure 5 demonstrates that the viability loss of the lab-cultured studies harvested in the logarithmic phase is lower than those that harvested in the stationary phase. This observation implies the impact of the cyanobacteria stage of life on pre-oxidation efficiency. A comparison of the cell viability results of the lab-cultured with natural bloom studies following potassium permanganate pre-oxidation confirms the higher resistance of real cyanobacterial bloom cells (Figure 5). In addition, the degradation rate constant of dissolved MCs was higher than that released by MCs for high potassium permanganate doses [36,81].

3.5. Hydrogen Peroxide

Matthijs, et al. [86] reported that a concentration of 2 mg/L H2O2 was able to decrease cyanobacteria (natural bloom) by two logs within 3 days. In addition, cyanobacteria remained at a low abundance level for 7 weeks following H2O2 addition. Figure 6 demonstrates that natural blooms are more resistant to H2O2 than the lab-cultured cyanobacteria, as observed for other oxidants (Figure 2, Figure 3, Figure 4 and Figure 5). Foo, et al. [87] reported that the impact of H2O2 on cyanobacteria is dependent on the residual concentration (C) and contact time (T). In addition, the authors concluded that toxic and non-toxic Microcystis aeruginosa are impacted by H2O2 with the same trend. Zhou, et al. [84] stated that a low dose of H2O2 (<5 mg/L) would have a low and recoverable impact on the lab-cultured Microcystis. On the other hand, the higher the H2O2 dose (>8 mg/L), the higher necrosis, cell death, and consequent cyanotoxin release. A medium dosage of H2O2 with low to medium contact time can activate apoptosis-like programmed cell death (AL-PCD) [84]. The cellular energy required for AL-PCD is provided from the transcriptional, biochemical, and structural changes. Zhou, et al. [84] documented the maximum cell death with low MC production by AL-PCD activation. Zamyadi, et al. [17] studied the impact of H2O2 on blooms and lab-cultured cyanobacteria (Microcystis aeruginosa). The results highlighted a delayed impact of H2O2 on cyanobacteria cells after complete depletion of H2O2 during stagnation (up to one week) [17]. Chl-a and phycocyanin (PC) fluorescence were significantly declined by 93% and 74% in natural bloom and lab-cultured samples, respectively. Additionally, the lab-cultured results revealed delayed MC release during stagnation [17].
Besides the current oxidants, peracetic acid (PAA) has been used in wastewater treatment facilities as a disinfection alternative for chlorine [88]. Almuhtaram and Hofmann [89] studied the impact of PAA and PAA/UV on cyanobacteria and cyanotoxin removal. The results show that 10 mg/L of PAA with 60 min contact time was able to degrade MC-LR by 80% (3.46 M−1 s−1 lower reaction rate as compared to HOCl 1.2 × 102 M−1s−1). In addition, the results elaborated that PAA alone can barely remove cyanobacteria, except at a high dose (10 mg/L) and with lower cyanobacterial cell counts (105 cells/mL).

3.6. Considerations on the Impact of Pre-Oxidation on Downflow Processes

The impact of pre-oxidation on downflow processes should also be considered as it may influence the removal of cyanobacteria by coagulation, flocculation, and sedimentation. Previous studies have been reported that pre-oxidation has a positive impact on enhancing cyanobacterial removal through coagulation/flocculation and sedimentation [27,28,31,32,34,90]. Pre-oxidation can cause morphological deformation [82] and changes in the surface charge of the cells, leading to increased cell removal efficiency during coagulation/flocculation [37].
KMnO4 increases the binding potential to the coagulant by oxidizing organic matter (extracellular and released cell-bound) to lower molecular weight fractions, as well as forming colloids (by MnO2) to be adsorbed to the cyanobacterial cells and forming larger flocs [32,34,81]. Xie, et al. [27] reported that KMnO4 exposure (CT: 10 mg min/L, estimated) could increase cyanobacteria cell removal by 22% during coagulation/flocculation. In addition, pre-ozonation with CT: 4, 10, and 20 mg min/L (estimated) led to an increase in cyanobacteria cell removal during coagulation by 14%, 20%, and 24%, respectively [27]. Cyanobacteria cell removal during coagulation was improved in a full-scale DWTP equipped by pre-ozonation systems (CT: 2.52–3.78 mg min/L (estimated)) [43]. Pre-oxidation may cause metabolite release (organic matter and cell-bound cyanotoxins) following cyanobacterial cell damage. Besides the challenge to remove dissolved cyanotoxins, coagulation efficiency can be compromised by high algal organic matter release following pre-oxidation. Xie, et al. [27] showed that due to pre-ozonation with CT > 4 mg min/L (estimated), cyanobacteria cell viability was completely degraded, and consequently, organic matter concertation increased. Further, Barešová, et al. [91] demonstrated that pre-ozonation (CT < 40 mg min/L (estimated)) could interrupt the coagulation (Al/Fe-based) efficiency of DOC removal (in comparison with higher CTs) due to the degradation of high molecular weight algal organic matter to low molecular weight compounds.
It is noteworthy to recall that H2O2 can have a delayed impact on cyanobacteria and, potentially, cyanotoxin release after complete degradation of the oxidant [17]. This delayed cyanotoxin release should be considered in the downstream processes, as well as in sludge handling.
The oxidant exposure must be adjusted to maximize cell damage and cyanobacteria cell removal (directly or after coagulation) and minimize cyanotoxin release and cell accumulation in the sludge, simultaneously. Figure 7 summarizes the pre-oxidation (soft and normal) advantages/disadvantages of cyanobacteria and cyanotoxins during water treatment. In fact, soft pre-oxidation (low CT of Cl2 and O3) can (1) partially degrade cyanobacteria cells, (2) cause low cyanotoxin release, (3) improve coagulation efficiency to remove cells, and (4) cause low cell accumulation in the downflow processes.

4. Sludge Storage, Oxidation, and Handling

Cyanobacteria and cyanotoxins (cell-bound) accumulate in the sludge of clarifiers throughout the flocculation/coagulation/sedimentation processes. This cyanobacteria-laden sludge remains in the sludge holding tank before disposal. In addition, potential options to treat cyanobacteria-laden sludge need to be considered. Furthermore, safe (healthy, both operationally and environmentally) cyanobacteria-laden sludge handling approaches are required.

4.1. Fate of Cyanobacteria and Cyanotoxins during Sludge Storage

Several studies (Table 5) demonstrated that cyanobacteria cells could stay viable within 2–12 days in the stored sludge. The loss of viability and consequent cyanotoxin release caused an increase in dissolved cyanotoxin concentrations during sludge storage [13,40,45,46,47,48,49,50,51,52]. However, dissolved cyanotoxins in stored sludge can be adsorbed onto the remained PAC injected into the intake water [56], flocs [50] or it can be biodegraded by cyanotoxin degrader species [58,92].
Besides cell survival potential during sludge storage, some studies have hypothesized that cyanobacteria can also grow in stored sludge [53,54,55]. Water Research Foundation (WRF) and Water Research Australia [53] documented that concentrations of DOC, MC-LR, and cylindrospermopsin in stored coagulated sludge contained M. aeruginosa and C. raciborskii exceeded the expected concentrations by 4–10-fold based on the cell quota (if all cell-bound metabolites are released) within 7–16 days, respectively. Dreyfus, et al. [55] studied the fate of stored sludge that contained cultured M. aeruginosa, D. circinale, and C. raciborskii within 18 days. The authors demonstrated that DOC, MC-LR, MC-LA, and CYN concentrations increased by up to 5-, 2.2-, 1.2-, and 2.5-fold during storage, respectively. Another investigation on stored sludge containing cultured M. aeruginosa and D. circinale reported that taxonomic cell counts increased by up to 4.2-fold in sludge stored in a lagoon within 7 days [54]. The authors also reported that the concentrations of cyanobacterial metabolites increased by up to 5 times in the sludge supernatant within 20 days. In the worst case, cyanobacteria could survive by up to 35 days in the stored sludge [54]. Despite the important findings of the previous studies on cell survival and metabolite release during sludge storage, cyanobacterial cell growth during sludge storage is yet to be explored in detail. In these studies, cell and metabolite increase during sludge storage might be due to the cell growth or to either the (1) underestimation of cell quota, (2) increase of metabolite production per cell during storage, or (3) additional cell settling from the supernatant to the sludge during the storage [53,54,55].
Our recent study on the cyanobacteria-laden sludge of a DWTP documented cell depletion, survival, and growth in different sludge samples [58]. Cell growth was observed in four out of eight sludge samples (different sampling dates) stored in the dark for 7–38 days. In the worst-case scenario, taxonomic cell counts increased from 2.7 × 106 to 5.3 × 106 cells/mL within 16 days (96% cell growth). Cell growth was also confirmed by increasing cyanobacterial biomarkers such as the “Pentose phosphate pathway” marker, which is responsible for the heterotrophic growth of cyanobacteria [93].
Table
Table 5. Impact of sludge storage on cyanobacteria and cyanotoxins. STX: saxitoxin, PACl: colyaluminium chloride, CTSAC: Chitosan-aluminum chloride.
Table 5. Impact of sludge storage on cyanobacteria and cyanotoxins. STX: saxitoxin, PACl: colyaluminium chloride, CTSAC: Chitosan-aluminum chloride.
Initial Characteristics of Cyanobacteria/
Coagulation/Sedimentation Process		Initial Condition of Cyanobacteria and Cyanotoxins in the Stored SludgeObservationReference
Cultured M. aeruginosa
(1 × 106 cells/mL)
(Jar test, 70 mg/L alum)		8 × 106 cells/mL,
2500 µg MC-LR/L		Cell survival (2 days); cell lysis and cyanotoxin release (2 days); degradation of dissolved cyanotoxins (8–10 days)[13]
Cultured D. circinale and C. raciborskii (1.0 × 105 cells/mL)
(Jar test, 40 mg/L alum)		Sludge supernatant:
D. circinale:
1300 cells/mL
STX: 0.4 µg/L		Cells remained viable up to 7 days; cell lysis and toxin release within 3 days[45]
Cultured M. aeruginosa
(2 × 106 cells/mL)
(Jar test, 15 mg/L AlCl3) 		18 µg/L dissolved MCsCell lysis and cyanotoxin release after 6 days[40]
Cultured M. aeruginosa
(1 × 106 cells/mL)
(Jar test, 4 mg/L PACl-optimum dose)		20 µg/L dissolved MCsCell lysis and cyanotoxin release within 6–12 days[46]
Microcystisflos aquae
(5.2 × 105 cells/mL)
(Jar test, 100 mg/L alum)		Sludge supernatant:
MC-RR, MC-YR: < 2 µg/L		Cell survival (5 days); cell lysis and cyanotoxin release (5–10 days); degradation of dissolved cyanotoxins (up to 15 days)[62]
Cultured M. aeruginosa
(1 × 106 cells/mL)
(Jar test, 15 mg/L ALCl3, 4 mg/L PACl)		−0.9 bar vacuum pressure for dewatering the sludge
23 µg/L total MCs		Cell lysis and cyanotoxin release within 4–6 days; optimum sludge storage time for AlCl3 and PACl was suggested to be 4 and 2 days, respectively.[47]
Cultured M. aeruginosa
(1 × 106 cells/mL)
(Jar test, 0–70 mg/L FeCl3)		~1 µg/L dissolved MCsCell lysis and cyanotoxin release (2–8 days); degradation of dissolved cyanotoxins (> 10 days)[48]
Myponga reservoir
Cultured M. aeruginosa (2.3 × 105 cells/mL)
Cell-bound MC-LR: 4.7 µg/L
Dissolved MC-LR: 2.0 µg/L
(Jar test-80 mg/L alum)		Sludge supernatant after 1 day storage:
Cells: 4300 cells/mL
Cell-bound MC-LR: 0.5 µg/L
Dissolved MC-LR: 2.5 µg/L		Cell survival (4 days); cell lysis and cyanotoxin release (4–7 days); degradation of dissolved cyanotoxins (> 4 days)[53]
Myponga reservoir
Cultured M. aeruginosa (3.1 × 105 cells/mL)
DOC: 10.1 mg/L
Cell-Bound MC-LR: 5.0 µg/L
Dissolved MC-LR: 2.9 µg/L
(Jar test-80 mg/L alum)		Sludge supernatant after 1 day storage:
DOC: 5.2 mg/L
Cell: 2760 cells/mL
Cell-bound MC-LR: <DL
Dissolved MC-LR: 4.0 µg/L		Cell growth (within 7–16 days) confirmed by DOC and MC-LR cell quota
Myponga reservoir
Cultured C. raciborskii (3.1 × 105 cells/mL)
DOC: 10 mg/L
Cell-bound CYN: 2.5 µg/L
Dissolved CYN: 0.7 µg/L
(Jar test-80 mg/L alum)		Sludge supernatant after 1 day storage:
DOC: 6.0 mg/L
Cell: 7080 cells/mL
Cell-bound CYN: 1.0 µg/L
Dissolved CYN: 0.8 µg/L		Cell growth (within 7–23 days) confirmed by DOC and CYN cell quota
River Murary
Cultured C. raciborskii (3.1 × 105 cells/mL)
DOC: 8.63 mg/L
Cell-bound CYN: 2.7 µg/L
Dissolved CYN: 0.3 µg/L
(Jar test-80 mg/L alum)		Sludge supernatant after 1 day storage:
DOC: 4.9 mg/L
Cell: 4140 cells/mL
Cell-bound CYN: 0.3 µg/L
Dissolved CYN: 0.9 µg/L		Cell growth (within 15–23 days) confirmed by DOC and CYN cell quota
Cultured M. aeruginosa
(1 × 106 cells/mL)
(Jar test- 15 mg/L AlCl3, 50 mg/L FeCl3, 15 mg/L PAFC)		20 µg/L dissolved MCs
1–4.2 mg/L dissolved polysaccharides
4 mg/L chla		Cell lysis and toxin release (2–10 days)[49]
Cultured M. aeruginosa
(2 × 106 cells/mL)
(Jar test, 2.6 mg/L chitosan- 7.5 mg/L AlCl3 (CTSAC)		9 µg/L dissolved MCs
(after coagulation)
18 µg/L dissolved MCs
(without coagulation); the difference is due to adsorption in CTSAC		Toxin release (0–4 days); degradation of dissolved cyanotoxins (6–10 days)[50]
M. aeruginosa, D. circinale, C. raciborskii
(3.0 × 105 cells/mL)
(Jar test, 80 mg/L Alum)		Sludge supernatant after 1 day storage:
DOC: 5.2–6.5 mg/L
Cell: 2162–7080 cells/mL
Cell-bound MC-LR: <0.5 µg/L
Dissolved MC-LR: 2.5–4.0 µg/L
Cell-bound CYN: 1.0 µg/L
Dissolved CYN: 0.8 µg/L		Increased DOC, MC-LR, MC-LA, and CYN to higher the expected values
(hypothesis: increase of the metabolite production, cell growth or both)		[55]
M. aeruginosa and D. circinale
(8.6 × 104 cells/mL)
(Jar test, 80 mg/L Alum)		Non-coagulated sludge: 5.0 × 106 cells/mL
Coagulated sludge: 5.4 × 105 cells/mL		Cell survival (up to 35 days); 4.2× increase in cell counts in the sludge lagoon within 7 days; increased metabolites to higher the expected values (up to 5×); increased cell counts in the sludge
(hypothesis: cell growth, additional settling, or both)		[54]
Cultured M. aeruginosa
× 105 cells/mL)
(Jar test, 15 mg/L AlCl3, 50 mg/L FeCl3, 15 mg/L PAFC)		1 µg/L dissolved MCs
Cell lysis and toxin release (4–6 days); degradation of dissolved cyanotoxins (6–10 days)[51]
Cultured Oscillatoria sp.
(1.0 × 104 cells/mL)
(Jar test, 5 and 10 mg/L PAFC)		1.0 mg/L chla
2.3 µg/L cell-bound protein
8.6–11.4 µg/L dissolved CYN		Increase in chla level after 4 days, suggesting cell growth; loss of cell integrity after 2 days, while cells remained viable up to 8 days; increase in dissolved CYN, showing toxin release within 4 days[94]
Cultured C. raciborskii
(1 × 106 cells/mL at late exponential phase)
(Jar test, 10 mg/L PAFC)		1.1 µg/L dissolved CYN
2 mg/L cell-bound protein		Cell lysis and toxin release after 6 days; degradation of dissolved cyanotoxins after 10 days[52]
n/aSludge of a DWTP containing natural cyanobacterial blooms stored for 7–35 days in the darkness (8 samples).
0.7 × 105–5.6 × 106 cells/mL
25–7130 ng/L cell-bound MCs
38–349 ng/L dissolved MCs		Cell growth in 4/8 samples after 9–35 stagnation days; cell death in the rest 4/8 samples; degradation of dissolved cyanotoxins after 8 days[58]

4.2. Cyanobacteria-Laden Sludge Treatment

A summary of studies on the treatment of cyanobacteria-laden sludge is presented in Table 6. The available data demonstrated that sludge oxidation could not completely remove cyanobacteria cells and metabolites from the sludge [57,59]. Sludge is often stored after oxidation, while its supernatant can be recycled to the head of the DWTP. Thus, the impact of oxidation on sludge storage should be investigated. Recent findings showed no remarkable benefits in sludge oxidation followed by sludge storage as compared to only sludge storage [59]. The maximum additional taxonomic cell count decreased by a combination of oxidation (KMnO4 or H2O2) and storage was 32% as compared to storage only. However, oxidation/storage could cause a remarkable cell growth (by up to 145%) and toxic gene copy numbers of mcyD increase (by up to 13.0×) in some sludge samples [59]. This phenomenon can be attributed to gene expression regulation due to the presence of oxidative stresses [58,59,95,96]. Similarly, sludge oxidation could not completely remove cyanobacteria and cyanotoxins from the supernatant sludge [59]. Finally, the costs and by-product formation during the oxidation of organic-matter-rich sludge should be considered [58].

4.3. Sludge Handling Challenges

In general, the sludge supernatant is recycled to the head of the DWTP or is discharged into the source [13,56,97]. The solid phase either can be transferred to the WWTP or is applied for landfilling [98,99,100]. Less environmentally friendly approaches such as untreated residual discharge into lakes or ponds can be also applied in some circumstances [101]. The re-use of DWTPs’ residuals is growing [99,102,103,104]. However, an investigation demonstrated that the half-life of MC analogs varies from 8 to 18 days in soil [105]. Since there is a risk of soil and groundwater contamination, landfill and field applications of cyanobacteria-laden sludge should be avoided. Overall, cyanobacteria-laden sludge should be treated before disposal either in situ or via sending it to wastewater treatment plants.
Flocs may have a protective role during sludge storage for M. aeruginosa [40,46,47,48,52]. In contrast, Li, et al. [52] documented that polyaluminium ferric chloride (PAFC) can stimulate the lysis of C. raciborskii and CYN release by up to 94% during sludge storage. This may occur in the sludge and lead to cyanotoxin release. However, all studies have been conducted in laboratory conditions and on cultured-based cyanobacteria. In fact, due to complex parameters such as the presence of various cyanobacterial cells in various forms (aggregated, multicellular), ages, and viabilities, the design of such experiments in full scale is complex.
Stresses such as oxidation and storage can shift cyanobacterial communities towards resistant genera (e.g., Microcystis and Aphanocapsa), which can produce MCs [56,58,59]. Thus, the survival probability of MC producer species can increase during sludge oxidation or storage. The fate of cyanotoxins in the sludge is complex due to the simultaneous occurrence of various phenomena such as cell survival, growth, lysis, cell-bound cyanotoxin release, and released cyanotoxin degradation [53,54,55,56,59,106,107]. Based on the increased risk of cell lysis and cyanotoxin release during sludge storage, some studies have suggested that cyanobacteria-laden sludge should be disposed of prior to 4 days to avoid metabolite release [47,51,108]. However, these studies only focused on metabolite release and not on cell survival/growth phenomena. Additionally, the possibility of sludge disposal can be a technical and financial challenge in large DWTPs.

5. Decision Framework to Manage Cyanobacteria and Cyanotoxins in Drinking Water Treatment Plants

5.1. Framework Basis

Since cyanobacterial cells and their associated metabolites, including cyanotoxins, as well as taste and odor agents such as geosmin and 2-Methylisoborneol (MIB), affect water and sludge quality, monitoring should be applied for the evaluation of the water treatment chain and sludge handling.
Microscopy taxonomic cell count techniques have been widely applied to evaluate the water and sludge in previous studies [12,18,43,44,56,77]. Previous cyanobacterial monitoring guidelines were prepared based on taxonomic cell counts and biovolumes [109,110,111]. A recent study suggested 0.3 mm3/L biovolumes as the vigilance level [109]. However, bias related to human error [112], the negative impact of Lugol’s iodine on biovolumes [113], cell underestimation/overestimation due to the presence of aggregated cells [114], and the presence of debris and sediments, especially in the sludge samples [56], may affect the results. More importantly, the significant time required for taxonomic cell counts is a major barrier in using them for a real-time/practical approach.
In situ fluorometry using on-line probes is a compromising technique for measurement of PC based on relative fluorescence units (RFU) in water resources [18,115,116,117,118,119]. However, the correlation between RFU and biovolume is complex and site-specific [119,120,121]. Previous investigations have reported that various RFUs ranging from 0.7 to 1.8 could correlate with a 0.3 mm3/L biovolume in different sources and bloom events [116,119]. Therefore, it is recommended to perform a correlation between on-line probe readings (RFU) and biovolumes for each water resource. It is noteworthy that the limits of detection and quantification of on-line probes should be considered [116,118].
MC concentration has been introduced in several guidelines such as those of the WHO (1.0 μg/L MC-LR) [122,123] and Health Canada (1.5 μg/L) [124]. Geosmin and MIB negatively affect water quality, raise complaints about taste and odor, and decrease the public’s confidence about the treated water safety [125,126,127,128,129]. Thus, they should be monitored throughout the treatment chain and the recycled sludge supernatant. Using enzyme-linked immunoassay (ELISA) tests for cyanotoxins measurement has been accepted for cyanotoxin monitoring [44,130,131,132]. The reported thresholds for geosmin and MIB are 1.3–4.0 ng/L and 6.3–15 ng/L, respectively [133,134,135]. Since the taste and odor agents are not harmful, but increase complaints and concerns about the water quality [126,127,129,134], olfactory detection can be considered for monitoring and detection [128,136].

5.2. Decision Framework

A decision framework to manage cyanobacteria and cyanotoxins in DWTPs is presented in Figure 8. The objective of this framework is to minimize cell breakthrough and accumulation throughout DWTPs and sludge. The three steps, (i) source water risk assessment, (ii) treatment breakthrough assessment and management, and (iii) sludge and supernatant risk assessment and management, should be taken for cyanobacteria and cyanotoxin control in DWTPs and sludge. This framework can help water utilities to understand appropriate approaches/strategies against cyanobacteria and cyanotoxins in DWTPs.
Overall, taxonomic cell counts, MCs, and taste and odor agents should be monitored in the (i) intake water, (ii) treatment chain, and (iii) sludge supernatant. These points are subjected to cyanobacteria and cyanotoxin accumulation, leading to a negative impact on water quality.
The optimization of conventional processes may include coagulant dose adjustment, applying aid-flocculants, and lowering the sedimentation and filtration rate during cell breakthrough [12,137,138,139]. Secondary barriers such as pre-oxidation, PAC injection, and GAC, in case of metabolite breakthrough, should be applied [15,25,39,65,140,141,142,143,144]. The impact of supernatant recycling or discharging on the source/intake water quality should be considered during toxic cyanobacterial blooms. Supernatant treatment may be required in the presence of cyanotoxins or taste and odor agents. MC concentration levels should be monitored in the sludge (solids) in case of landfilling or land application. In the case of elevated concentrations of MCs, sludge treatment is required.

6. Conclusions

○
Using the exposure unit (CT) is recommended for cyanobacteria and cyanotoxins oxidation studies, rather than using dose or contact time individually.
○
Regardless of the oxidant type, lab-cultured studies cannot depict the complete picture of natural cyanobacterial bloom behavior during oxidation and may overestimate the oxidation efficiency. In addition, cyanobacterial bloom oxidation is site- and bloom-specific, which could be related to the level of agglomeration, cyanobacteria (bloom) stage of life, and metabolic functions.
○
Soft pre-chlorination and pre-ozonation can compromise cell viability with no or limited cyanotoxin release. Overall, soft pre-oxidation may cause lower disinfection by-products compared to normal pre-oxidation.
○
The cyanobacteria in stored sludge can not only survive, but also grow and release cyanotoxins, even in the dark. Although dissolved cyanotoxins can be degraded during sludge storage, the potential risk of growth and cyanotoxin release should be considered. In fact, the cell growth/depletion in stored sludge is complex and not easy to predict. Therefore, the worst-case scenario should be considered during sludge handling.
○
Due to the low efficacy of sludge oxidation as compared to only stored sludge, as well as the occurrence of cell growth, and gene expression regulation during oxidation/storage, oxidation cannot be a reliable approach in sludge treatment and management.
○
Management of cyanobacteria and cyanotoxins in sludge should be initiated with the minimization of cyanobacteria and cyanotoxin accumulation throughout DWTPs.
○
To control the negative impacts of cyanobacterial accumulation in DWTPs, recycling sludge supernatant to the head of the DWTPs should be regulated during cyanobacterial seasons. Suitable treatment and disposal approaches should be set into guidance and regulations for sludge-containing cyanotoxins.

Author Contributions

Conceptualization, F.J., S.M., S.D., A.Z., S.S. and M.P.; methodology, F.J., S.M., S.D. and M.P.; validation, F.J., S.M., A.Z., S.D. and M.P.; formal analysis, F.J., S.M. and M.P.; investigation, F.J., S.M., S.D., A.Z. and M.P.; writing—original draft preparation, F.J. and S.M.; writing—review and editing, F.J., S.M., S.D., A.Z., S.S. and M.P.; visualization, F.J. and S.M.; supervision, S.D., S.S. and M.P.; project administration, S.S., S.D. and M.P.; funding acquisition, S.D. and S.S. All authors have read and agreed to the published version of the manuscript.

Funding

The authors sincerely acknowledge the financial supports of Algal Blooms, Treatment, Risk As-sessment, Prediction, and Prevention through Genomics (ATRAPP) projects, Génome Québec, Genome Canada and the NSERC Industrial Chair on Drinking Water at Polytechnique Montréal (grant number ALLRP 560764-20).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Churro, C.; Dias, E.; Valério, E. Risk assessment of cyanobacteria and cyanotoxins, the particularities and challenges of Planktothrix spp. monitoring. In Novel Approaches and Their Applications in Risk Assessment; IntechOpen: London, UK, 2012; pp. 59–85. [Google Scholar]
  2. Kosten, S.; Huszar, V.L.M.; Bécares, E.; Costa, L.S.; Donk, E.; Hansson, L.-A.; Jeppesen, E.; Kruk, C.; Lacerot, G.; Mazzeo, N.; et al. Warmer climates boost cyanobacterial dominance in shallow lakes. Glob. Chang. Biol. 2012, 18, 118–126. [Google Scholar] [CrossRef]
  3. Paerl, H.W.; Paul, V.J. Climate change: Links to global expansion of harmful cyanobacteria. Water Res. 2012, 46, 1349–1363. [Google Scholar] [CrossRef] [PubMed]
  4. Chorus, I.; Fastner, J.; Welker, M. Cyanobacteria and Cyanotoxins in a Changing Environment: Concepts, Controversies, Challenges. Water 2021, 13, 2463. [Google Scholar] [CrossRef]
  5. Mokoena, M.M.; Mukhola, M.S. Current Effects of Cyanobacteria Toxin in Water Sources and Containers in the Hartbeespoort Dam Area, South Africa. Int. J. Environ. Res. Public Health 2019, 16, 4468. [Google Scholar] [CrossRef] [Green Version]
  6. Huisman, J.; Codd, G.A.; Paerl, H.W.; Ibelings, B.W.; Verspagen, J.M.H.; Visser, P.M. Cyanobacterial blooms. Nat. Rev. Microbiol. 2018, 16, 471–483. [Google Scholar] [CrossRef]
  7. O’Keeffe, J. Cyanobacteria and Drinking Water: Occurrence, Risks, Management and Knowledge Gaps for Public Health; National Collaborating Centre for Environmental Health: Vancouver, BC, Canada, 2019. [Google Scholar]
  8. Mishra, S.; Stumpf, R.P.; Schaeffer, B.A.; Werdell, P.J.; Loftin, K.A.; Meredith, A. Measurement of Cyanobacterial Bloom Magnitude using Satellite Remote Sensing. Sci. Rep. 2019, 9, 18310. [Google Scholar] [CrossRef]
  9. Akyol, C.; Ozbayram, E.G.; Accoroni, S.; Radini, S.; Eusebi, A.L.; Gorbi, S.; Vignaroli, C.; Bacchiocchi, S.; Campacci, D.; Gigli, F.; et al. Monitoring of cyanobacterial blooms and assessing polymer-enhanced microfiltration and ultrafiltration for microcystin removal in an Italian drinking water treatment plant. Environ. Pollut. 2021, 286, 117535. [Google Scholar] [CrossRef]
  10. Zamyadi, A.; Glover, C.M.; Yasir, A.; Stuetz, R.; Newcombe, G.; Crosbie, N.D.; Lin, T.-F.; Henderson, R. Toxic cyanobacteria in water supply systems: Data analysis to map global challenges and demonstrate the benefits of multi-barrier treatment approaches. H2Open J. 2021, 4, 47–62. [Google Scholar] [CrossRef]
  11. Westrick, J.A.; Szlag, D.C.; Southwell, B.J.; Sinclair, J. A review of cyanobacteria and cyanotoxins removal/inactivation in drinking water treatment. Anal. Bioanal. Chem. 2010, 397, 1705–1714. [Google Scholar] [CrossRef]
  12. Zamyadi, A.; Dorner, S.; Ellis, D.; Bolduc, A.; Bastien, C.; Prévost, M. Species-dependence of cyanobacteria removal efficiency by different drinking water treatment processes. Water Res. 2013, 47, 2689–2700. [Google Scholar] [CrossRef]
  13. Drikas, M.; Chow, C.W.K.; House, J.; Burch, M.D. Using coagulation, flocculation, and settling to remove toxic cyanobacteria. J. Am. Water Work. Assoc. 2001, 93, 100–111. [Google Scholar] [CrossRef]
  14. Szlag, D.C.; Sinclair, J.L.; Southwell, B.; Westrick, J.A. Cyanobacteria and Cyanotoxins Occurrence and Removal from Five High-Risk Conventional Treatment Drinking Water Plants. Toxins 2015, 7, 2198–2220. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  15. Newcombe, G.; Nicholson, B. Water treatment options for dissolved cyanotoxins. Water Supply Res. Technol.-Aqua 2004, 53, 227–239. [Google Scholar] [CrossRef]
  16. Wert, E.C.; Rosario-Ortiz, F.L. Intracellular organic matter from cyanobacteria as a precursor for carbonaceous and nitrogenous disinfection byproducts. Environ. Sci. Technol. 2013, 47, 6332–6340. [Google Scholar] [CrossRef] [PubMed]
  17. Zamyadi, A.; Greenstein, K.E.; Glover, C.M.; Adams, C.; Rosenfeldt, E.; Wert, E.C. Impact of hydrogen peroxide and copper sulfate on the delayed release of microcystin. Water 2020, 12, 1105. [Google Scholar] [CrossRef]
  18. Zamyadi, A.; MacLeod, S.; Fan, Y.; McQuaid, N.; Dorner, S.; Sauvé, S.; Prévost, M. Toxic cyanobacterial breakthrough and accumulation in a drinking water plant: A monitoring and treatment challenge. Water Res. 2012, 46, 1511–1523. [Google Scholar] [CrossRef] [PubMed]
  19. Henderson, R.K.; Parsons, S.A.; Jefferson, B. The impact of differing cell and algogenic organic matter (AOM) characteristics on the coagulation and flotation of algae. Water Res. 2010, 44, 3617–3624. [Google Scholar] [CrossRef] [Green Version]
  20. Newcombe, G. International Guidance Manual for the Management of Toxic Cyanobacteria; Global Water Research Coalition and Water Quality Research Australia: London, UK, 2009; p. 44. [Google Scholar]
  21. Gonzalez-Torres, A.; Putnam, J.; Jefferson, B.; Stuetz, R.M.; Henderson, R.K. Examination of the physical properties of Microcystis aeruginosa flocs produced on coagulation with metal salts. Water Res. 2014, 60, 197–209. [Google Scholar] [CrossRef]
  22. Pivokonsky, M.; Naceradska, J.; Brabenec, T.; Novotna, K.; Baresova, M.; Janda, V. The impact of interactions between algal organic matter and humic substances on coagulation. Water Res. 2015, 84, 278–285. [Google Scholar] [CrossRef]
  23. Merel, S.; Walker, D.; Chicana, R.; Snyder, S.; Baures, E.; Thomas, O. State of knowledge and concerns on cyanobacterial blooms and cyanotoxins. Environ. Int. 2013, 59, 303–327. [Google Scholar] [CrossRef]
  24. He, X.; Liu, Y.-L.; Conklin, A.; Westrick, J.; Weavers, L.K.; Dionysiou, D.D.; Lenhart, J.J.; Mouser, P.J.; Szlag, D.; Walker, H.W. Toxic cyanobacteria and drinking water: Impacts, detection, and treatment. Harmful Algae 2016, 54, 174–193. [Google Scholar] [CrossRef] [PubMed]
  25. Abbas, T.; Kajjumba, G.W.; Ejjada, M.; Masrura, S.U.; Marti, E.J.; Khan, E.; Jones-Lepp, T.L. Recent Advancements in The Removal of Cyanotoxins from Water Using Conventional and Modified Adsorbents—A Contemporary review. Water 2020, 12, 2756. [Google Scholar] [CrossRef]
  26. Villars, K.; Huang, Y.; Lenhart, J.J. Removal of the Cyanotoxin Microcystin-LR from Drinking Water Using Granular Activated Carbon. Environ. Eng. Sci. 2020, 37, 585–595. [Google Scholar] [CrossRef]
  27. Xie, P.; Ma, J.; Fang, J.; Guan, Y.; Yue, S.; Li, X.; Chen, L. Comparison of permanganate preoxidation and preozonation on algae containing water: Cell integrity, characteristics, and chlorinated disinfection byproduct formation. Environ. Sci. Technol. 2013, 47, 14051–14061. [Google Scholar] [CrossRef]
  28. Lapsongpon, T.; Leungprasert, S.; Yoshimura, C. Pre-chlorination contact time and the removal and control of Microcystis aeroginosa in coagulation. IOP Conf. Ser. Earth Environ. Sci. 2017, 67, 012011. [Google Scholar] [CrossRef]
  29. Petrusevski, B.; van Breeman, A.N.; Alaerts, G.J. Effect of permanganate pre-treatment and coagulation with dual coagulants on algae removal in direct filtration. Water Supply Res. Technol.-Aqua 1996, 45, 316–326. [Google Scholar]
  30. Chu, W.; Yao, D.; Deng, Y.; Sui, M.; Gao, N. Production of trihalomethanes, haloacetaldehydes and haloacetonitriles during chlorination of microcystin-LR and impacts of pre-oxidation on their formation. J. Hazard. Mater. 2017, 327, 153–160. [Google Scholar] [CrossRef]
  31. Lin, J.L.; Hua, L.C.; Hung, S.K.; Huang, C. Algal removal from cyanobacteria-rich waters by preoxidation-assisted coagulation-flotation: Effect of algogenic organic matter release on algal removal and trihalomethane formation. J. Environ. Sci. 2018, 63, 147–155. [Google Scholar] [CrossRef]
  32. Naceradska, J.; Pivokonsky, M.; Pivokonska, L.; Baresova, M.; Henderson, R.K.; Zamyadi, A.; Janda, V. The impact of pre-oxidation with potassium permanganate on cyanobacterial organic matter removal by coagulation. Water Res. 2017, 114, 42–49. [Google Scholar] [CrossRef]
  33. Wang, H.; Masters, S.; Falkinham, J.O., 3rd; Edwards, M.A.; Pruden, A. Distribution system water quality affects responses of opportunistic pathogen gene markers in household water heaters. Environ. Sci. Technol. 2015, 49, 8416–8424. [Google Scholar] [CrossRef]
  34. Xie, P.; Chen, Y.; Ma, J.; Zhang, X.; Zou, J.; Wang, Z. A mini review of preoxidation to improve coagulation. Chemosphere 2016, 155, 550–563. [Google Scholar] [CrossRef] [PubMed]
  35. Zamyadi, A.; Fan, Y.; Daly, R.I.; Prévost, M. Chlorination of Microcystis aeruginosa: Toxin release and oxidation, cellular chlorine demand and disinfection by-products formation. Water Res. 2013, 47, 1080–1090. [Google Scholar] [CrossRef] [PubMed]
  36. Fan, J.; Daly, R.; Hobson, P.; Ho, L.; Brookes, J. Impact of potassium permanganate on cyanobacterial cell integrity and toxin release and degradation. Chemosphere 2013, 92, 529–534. [Google Scholar] [CrossRef] [PubMed]
  37. Fan, J.; Ho, L.; Hobson, P.; Brookes, J. Evaluating the effectiveness of copper sulphate, chlorine, potassium permanganate, hydrogen peroxide and ozone on cyanobacterial cell integrity. Water Res. 2013, 47, 5153–5164. [Google Scholar] [CrossRef] [PubMed]
  38. Moradinejad, S.; Trigui, H.; Maldonado, J.F.G.; Shapiro, B.J.; Terrat, Y.; Sauvé, S.; Fortin, N.; Zamyadi, A.; Dorner, S.; Prévost, M. Metagenomic study to evaluate functional capacity of a cyanobacterial bloom during oxidation. Chem. Eng. J. Adv. 2021, 8, 100151. [Google Scholar] [CrossRef]
  39. Moradinejad, S.; Trigui, H.; Guerra Maldonado, J.F.; Shapiro, J.; Terrat, Y.; Zamyadi, A.; Dorner, S.; Prévost, M. Diversity assessment of toxic cyanobacterial blooms during oxidation. Toxins 2020, 12, 728. [Google Scholar] [CrossRef]
  40. Sun, F.; Pei, H.-Y.; Hu, W.-R.; Ma, C.-X. The lysis of Microcystis aeruginosa in AlCl3 coagulation and sedimentation processes. Chem. Eng. J. 2012, 193–194, 196–202. [Google Scholar] [CrossRef]
  41. Teixeira, M.R.; Rosa, M.J. Comparing dissolved air flotation and conventional sedimentation to remove cyanobacterial cells of Microcystis aeruginosa. Part II. The effect of water background organics. Sep. Purif. Technol. 2007, 53, 126–134. [Google Scholar] [CrossRef]
  42. Chorus, I.; Bartram, J. Chapter 6. Situation assessment, planning and management. In Toxic Cyanobacteria in Water: A Guide to Their Public Health Consequences, Monitoring and Management; WHO: Geneva, Switzerland, 1999; p. 28. [Google Scholar]
  43. Zamyadi, A.; Dorner, S.; Ndong, M.; Ellis, D.; Bolduc, A.; Bastien, C.; Prévost, M. Low-risk cyanobacterial bloom sources: Cell accumulation within full-scale treatment plants. J. Am. Water Work. Assoc. 2013, 102, E651–E663. [Google Scholar] [CrossRef] [Green Version]
  44. Almuhtaram, H.; Cui, Y.; Zamyadi, A.; Hofmann, R. Cyanotoxins and cyanobacteria cell accumulations in drinking water treatment plants with a low risk of bloom formation at the source. Toxins 2018, 10, 430. [Google Scholar] [CrossRef] [Green Version]
  45. Ho, L.; Dreyfus, J.; Boyer, J.e.; Lowe, T.; Bustamante, H.; Duker, P.; Meli, T.; Newcombe, G. Fate of cyanobacteria and their metabolites during water treatment sludge management processes. Sci. Total Environ. 2012, 424, 232–238. [Google Scholar] [CrossRef] [PubMed]
  46. Sun, F.; Pei, H.-Y.; Hu, W.-R.; Li, X.-Q.; Ma, C.-X.; Pei, R.-T. The cell damage of Microcystis aeruginosa in PACl coagulation and floc storage processes. Sep. Purif. Technol. 2013, 115, 123–128. [Google Scholar] [CrossRef]
  47. Sun, F.; Hu, W.; Pei, H.; Li, X.; Xu, X.; Ma, C. Evaluation on the dewatering process of cyanobacteria-containing AlCl3 and PACl drinking water sludge. Sep. Purif. Technol. 2015, 150, 52–62. [Google Scholar] [CrossRef]
  48. Li, X.; Pei, H.; Hu, W.; Meng, P.; Sun, F.; Ma, G.; Xu, X.; Li, Y. The fate of Microcystis aeruginosa cells during the ferric chloride coagulation and flocs storage processes. Environ. Technol. 2015, 36, 920–928. [Google Scholar] [CrossRef] [PubMed]
  49. Xu, H.; Pei, H.; Xiao, H.; Jin, Y.; Li, X.; Hu, W.; Ma, C.; Sun, J.; Li, H. Behaviors of Microcystis aeruginosa cells during floc storage in drinking water treatment process. Sci. Rep. 2016, 6, 34943. [Google Scholar] [CrossRef] [Green Version]
  50. Ma, C.; Pei, H.; Hu, W.; Xu, H.; Jin, Y. The lysis and regrowth of toxic cyanobacteria during storage of achitosan–aluminium chloride composite coagulated sludge: Implications for drinking water sludge treatment. RSC Adv. 2016, 6, 112756–112764. [Google Scholar] [CrossRef]
  51. Xu, H.; Pei, H.; Jin, Y.; Xiao, H.; Ma, C.; Sun, J.; Li, H. Characteristics of water obtained by dewatering cyanobacteria-containing sludge formed during drinking water treatment, including C-, N-disinfection byproduct formation. Water Res. 2017, 111, 382–392. [Google Scholar] [CrossRef]
  52. Li, H.; Pei, H.; Xu, H.; Jin, Y.; Sun, J. Behavior of Cylindrospermopsis raciborskii during coagulation and sludge storage-higher potential risk of toxin release than Microcystis aeruginosa? J. Hazard. Mater. 2018, 347, 307–316. [Google Scholar] [CrossRef]
  53. Water Research Foundation (WRF); Water Research Australia. Optimizing Conventional Treatment for the Removal of Cyanobacteria and Toxins; Water Research Foundation (WRF): Denver, CO, USA, 2015; p. 185. ISBN 978-1-60573-216-9. [Google Scholar]
  54. Pestana, C.J.; Reeve, P.J.; Sawade, E.; Voldoire, C.F.; Newton, K.; Praptiwi, R.; Collingnon, L.; Dreyfus, J.; Hobson, P.; Gaget, V.; et al. Fate of cyanobacteria in drinking water treatment plant lagoon supernatant and sludge. Sci. Total Environ. 2016, 565, 1192–1200. [Google Scholar] [CrossRef]
  55. Dreyfus, J.; Monrolin, Y.; Pestana, C.J.; Reeve, P.J.; Sawade, E.; Newton, K.; Ho, L.; Chow, C.W.K.; Newcombe, G. Identification and assessment of water quality risks associated with sludge supernatant recycling in the presence of cyanobacteria. J. Water Supply Res. Technol.-Aqua 2016, 65, 441–452. [Google Scholar] [CrossRef]
  56. Jalili, F.; Trigui, H.; Guerra Maldonado, J.F.; Dorner, S.; Zamyadi, A.; Shapiro, B.J.; Terrat, Y.; Fortin, N.; Sauvé, S.; Prévost, M. Can cyanobacterial diversity in the source predict the diversity in sludge and the risk of toxin release in a drinking water treatment plant? Toxins 2021, 13, 25. [Google Scholar] [CrossRef] [PubMed]
  57. Zamyadi, A.; Henderson, R.K.; Stuetz, R.; Newcombe, G.; Newtown, K.; Gladman, B. Cyanobacterial management in full-scale water treatment and recycling processes: Reactive dosing following intensive monitoring. Environ. Sci. Water Res. Technol. 2016, 2, 362–375. [Google Scholar] [CrossRef]
  58. Jalili, F. Optimal Treatment Strategies to Prevent and Manage Cyanobacteria and Cyanotoxins in Drinking Water Sludge. Ph.D. Thesis, Polytechnique Montréal, Montréal, QC, Canada, 2022. [Google Scholar]
  59. Jalili, F.; Trigui, H.; Maldonado, J.F.G.; Dorner, S.; Zamyadi, A.; Shapiro, B.J.; Terrat, Y.; Fortin, N.; Sauvé, S.; Prévost, M. Oxidation to Control Cyanobacteria and Cyanotoxins in Drinking Water Treatment Plants: Challenges at the Laboratory and Full-Scale Plants. Water 2022, 14, 537. [Google Scholar] [CrossRef]
  60. Zamyadi, A.; Romanis, C.; Mills, T.; Neilan, B.; Choo, F.; Coral, L.A.; Gale, D.; Newcombe, G.; Crosbie, N.; Stuetz, R.; et al. Diagnosing water treatment critical control points for cyanobacterial removal: Exploring benefits of combined microscopy, next-generation sequencing, and cell integrity methods. Water Res. 2019, 152, 96–105. [Google Scholar] [CrossRef]
  61. Water Research Foundation (WRF); United States Environmental Protection Agency (US EPA); Veolia Water Indianapolis. Strategies for Controlling and Mitigating Algal Growth within Water Treatment Plants; Water Research Foundation (WRF): Denver, CO, USA, 2009; p. 312. [Google Scholar]
  62. Ho, L.; Barbero, A.; Dreyfus, J.; Dixon, D.R.; Qian, F.; Scales, P.J.; Newcombe, G. Behaviour of cyanobacterial bloom material following coagulation and/or sedimentation. J. Water Supply Res. Technol.-Aqua 2013, 62, 350–358. [Google Scholar] [CrossRef]
  63. Pinkanjananavee, K.; Teh, S.J.; Kurobe, T.; Lam, C.H.; Tran, F.; Young, T.M. Potential Impacts on Treated Water Quality of Recycling Dewatered Sludge Supernatant during Harmful Cyanobacterial Blooms. Toxins 2021, 13, 99. [Google Scholar] [CrossRef]
  64. Coral, L.A.; Zamyadi, A.; Barbeau, B.; Bassetti, F.J.; Lapolli, F.R.; Prévost, M. Oxidation of Microcystis aeruginosa and Anabaena flos-aquae by ozone: Impacts on cell integrity and chlorination by-product formation. Water Res. 2013, 47, 2983–2994. [Google Scholar] [CrossRef]
  65. Zamyadi, A.; Coral, L.A.; Barbeau, B.; Dorner, S.; Lapolli, F.R.; Prévost, M. Fate of toxic cyanobacterial genera from natural bloom events during ozonation. Water Res. 2015, 73, 204–215. [Google Scholar] [CrossRef]
  66. Ding, J.; Shi, H.; Timmons, T.; Adams, C. Release and removal of microcystins from microcystis during oxidative-, physical-, and UV-based disinfection. J. Environ. Eng. 2010, 136, 2–11. [Google Scholar] [CrossRef]
  67. Zamyadi, A.; Ho, L.; Newcombe, G.; Daly, R.I.; Burch, M.; Baker, P.; Prévost, M. Release and oxidation of cell-bound saxitoxins during chlorination of Anabaena circinalis cells. Environ. Sci. Technol. 2010, 44, 9055–9061. [Google Scholar] [CrossRef]
  68. Fan, J.J.; Hobson, P.; Ho, L.; Daly, R.; Brookes, J. The effects of various control and water treatment processes on the membrane integrity and toxin fate of cyanobacteria. J. Hazard. Mater. 2014, 264, 313–322. [Google Scholar] [CrossRef] [PubMed]
  69. Zhang, H.; Dan, Y.; Adams, C.D.; Shi, H.; Ma, Y.; Eichholz, T. Effect of oxidant demand on the release and degradation of microcystin-LR from Microcystis aeruginosa during oxidation. Chemosphere 2017, 181, 562–568. [Google Scholar] [CrossRef] [PubMed]
  70. Qi, J.; Lan, H.; Liu, R.; Miao, S.; Liu, H.; Qu, J. Prechlorination of algae-laden water: The effects of transportation time on cell integrity, algal organic matter release, and chlorinated disinfection byproduct formation. Water Res. 2016, 102, 221–228. [Google Scholar] [CrossRef] [PubMed]
  71. Li, X.; Chen, S.; Zeng, J.; Song, W.; Yu, X. Comparing the effects of chlorination on membrane integrity and toxin fate of high-and low-viability cyanobacteria. Water Res. 2020, 177, 115769. [Google Scholar] [CrossRef]
  72. Li, X.; Chen, S.; Zeng, J.; Chabi, K.; Song, W.; Xian, X.; Yu, X. Impact of chlorination on cell inactivation, toxin release and degradation of cyanobacteria of development and maintenance stage. Chem. Eng. J. 2020, 397, 125378. [Google Scholar] [CrossRef]
  73. Fan, J.; Rao, L.; Chiu, Y.T.; Lin, T.F. Impact of chlorine on the cell integrity and toxin release and degradation of colonial Microcystis. Water Res. 2016, 102, 394–404. [Google Scholar] [CrossRef]
  74. He, X.; Wert, E.C. Colonial cell disaggregation and intracellular microcystin release following chlorination of naturally occurring Microcystis. Water Res. 2016, 101, 10–16. [Google Scholar] [CrossRef]
  75. Greenstein, K.E.; Zamyadi, A.; Glover, C.M.; Adams, C.; Rosenfeldt, E.; Wert, E.C. Delayed Release of Intracellular Microcystin Following Partial Oxidation of Cultured and Naturally Occurring Cyanobacteria. Toxins 2020, 12, 335. [Google Scholar] [CrossRef]
  76. Moradinejad, S. Characterization of Phenomena Governing Oxidation of Cyanobacteria through Metagenomics. Ph.D Thesis, Polytechnique Montréal, Montreal, QC, Canada, 2021. [Google Scholar]
  77. Zamyadi, A.; Dorner, S.; Ndong, M.; Ellis, D.; Bolduc, A.; Bastien, C.; Prévost, M. Breakthrough and accumulation of toxic cyanobacteria in full scale clarification and filtration processes. In Proceedings of the American Water Works Association-Water Quality Technology Conference and Exposition (WQTC), Toronto, ON, Canada, 4–8 November 2012; p. 27. [Google Scholar]
  78. Wert, E.C.; Dong, M.M.; Rosario-Ortiz, F.L. Using digital flow cytometry to assess the degradation of three cyanobacteria species after oxidation processes. Water Res. 2013, 47, 3752–3761. [Google Scholar] [CrossRef]
  79. Moradinejad, S.; Glover, C.; Mailly, J.; Zadfatollah-Seighalani, T.; Peldszus, S.; Barbeau, B.; Dorner, S.; Prévost, M.; Zamyadi, A. Using advanced spectroscopy and organic matter characterization to evaluate the impact of oxidation on cyanobacteria. Toxins 2019, 11, 278. [Google Scholar] [CrossRef] [Green Version]
  80. Moradinejad, S.; Glover, C.; Dorner, S.; Barbeau, B.; Prévost, M.; Zamyadi, A. Using advanced spectroscopy and visualization techniques to assess the impact of oxidation on cyanobacteria cell morphology. In Proceedings of the WQTC, Dallas, TX, USA, 3–7 November 2019. [Google Scholar]
  81. Piezer, K.; Li, L.; Jeon, Y.; Kadudula, A.; Seo, Y. The Application of Potassium Permanganate to Treat Cyanobacteria-Laden Water. Process Saf. Environ. Prot. 2020, 148, 400–414. [Google Scholar] [CrossRef]
  82. Zhou, S.; Shao, Y.; Gao, N.; Deng, Y.; Qiao, J.; Ou, H.; Deng, J. Effects of different algaecides on the photosynthetic capacity, cell integrity and microcystin-LR release of Microcystis aeruginosa. Sci. Total Environ. 2013, 463, 111–119. [Google Scholar] [CrossRef] [PubMed]
  83. Xu, H.; Brookes, J.; Hobson, P.; Pei, H. Impact of copper sulphate, potassium permanganate, and hydrogen peroxide on Pseudanabaena galeata cell integrity, release and degradation of 2-methylisoborneol. Water Res. 2019, 157, 64–73. [Google Scholar] [CrossRef] [PubMed]
  84. Zhou, T.; Cao, H.; Zheng, J.; Teng, F.; Wang, X.; Lou, K.; Zhang, X.; Tao, Y. Suppression of water-bloom cyanobacterium Microcystis aeruginosa by algaecide hydrogen peroxide maximized through programmed cell death. J. Hazard. Mater. 2020, 393, 122394. [Google Scholar] [CrossRef] [PubMed]
  85. Daly, R.I.; Ho, L.; Brookes, J.D. Effect of chlorination on Microcystis aeruginosa cell integrity and subsequent microcystin release and degradation. Environ. Sci. Technol. 2007, 41, 4447–4453. [Google Scholar] [CrossRef]
  86. Matthijs, H.C.P.; Visser, P.M.; Reeze, B.; Meeuse, J.; Slot, P.C.; Wijn, G.; Talens, R.; Huisman, J. Selective suppression of harmful cyanobacteria in an entire lake with hydrogen peroxide. Water Res. 2012, 46, 1460–1472. [Google Scholar] [CrossRef]
  87. Foo, S.C.; Chapman, I.J.; Hartnell, D.M.; Turner, A.D.; Franklin, D.J. Effects of H2O2 on growth, metabolic activity and membrane integrity in three strains of Microcystis aeruginosa. Environ. Sci. Pollut. Res. 2020, 27, 38916–38927. [Google Scholar] [CrossRef]
  88. Luukkonen, T.; Teeriniemi, J.; Prokkola, H.; Rämö, J.; Lassi, U. Chemical aspects of peracetic acid based wastewater disinfection. Water Sa 2014, 40, 73–80. [Google Scholar] [CrossRef] [Green Version]
  89. Almuhtaram, H.; Hofmann, R. Evaluation of ultraviolet/peracetic acid to degrade M. aeruginosa and microcystins-LR and-RR. J. Hazard. Mater. 2022, 424, 127357. [Google Scholar] [CrossRef]
  90. Wang, H.-Q.; Mao, T.-G.; Xi, B.-D.; Zhang, L.-Y.; Zhou, Q.-H. KMnO4 pre-oxidation for Microcystis aeruginosa removal by a low dosage of flocculant. Ecol. Eng. 2015, 81, 298–300. [Google Scholar] [CrossRef]
  91. Barešová, M.; Načeradská, J.; Novotná, K.; Čermáková, L.; Pivokonský, M. The impact of preozonation on the coagulation of cellular organic matter produced by Microcystis aeruginosa and its toxin degradation. J. Environ. Sci. 2020, 98, 124–133. [Google Scholar] [CrossRef] [PubMed]
  92. Kormas, K.A.; Lymperopoulou, D.S. Cyanobacterial toxin degrading bacteria: Who are they? Biomed. Res. Int. 2013, 2013, 463894. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Min, H.; Golden, S.S. A new circadian class 2 gene, opcA, whose product is important for reductant production at night in Synechococcus elongatus PCC 7942. J. Bacteriol. 2000, 182, 6214–6221. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Sun, J.; Xu, H.; Pei, H.; Jin, Y.; Li, H.; Ma, C. Worse than cell lysis: The resilience of Oscillatoria sp. during sludge storage in drinking water treatment. Water Res. 2018, 142, 405–414. [Google Scholar] [CrossRef]
  95. Pimentel, J.S.; Giani, A. Microcystin production and regulation under nutrient stress conditions in toxic microcystis strains. Appl. Environ. Microbiol. 2014, 80, 5836–5843. [Google Scholar] [CrossRef] [Green Version]
  96. Schatz, D.; Keren, Y.; Vardi, A.; Sukenik, A.; Carmeli, S.; Börner, T.; Dittmann, E.; Kaplan, A. Towards clarification of the biological role of microcystins, a family of cyanobacterial toxins. Environ. Microbiol. 2007, 9, 965–970. [Google Scholar] [CrossRef]
  97. Chow, C.W.K.; Drikas, M.; House, J.; Burch, M.D.; Velzeboer, R.M.A. The impact of conventional water treatment processes on cells of the cyanobacterium Microcystis aeruginosa. Water Res. 1999, 33, 3253–3262. [Google Scholar] [CrossRef]
  98. United States Environmental Protection Agency (USEPA). Drinking Water Treatment Plant Residuals Management Technical Report, Summary of Residuals Generation, Treatment, and Disposal at Large Community Water Systems; United States Environmental Protection Agency (USEPA): Washington, DC, USA, 2011; p. 378.
  99. Turner, T.; Wheeler, R.; Stone, A.; Oliver, I. Potential Alternative Reuse Pathways for Water Treatment Residuals: Remaining Barriers and Questions—A Review. Water Air Soil Pollut. 2019, 230, 227. [Google Scholar] [CrossRef] [Green Version]
  100. Qrenawi, L.I.; Rabah, F.K.J. Sludge management in water treatment plants: Literature review. Int. J. Environ. Waste Manag. 2021, 27, 93. [Google Scholar] [CrossRef]
  101. Ahmad, T.; Ahmad, K.; Alam, M. Characterization of Water Treatment Plant’s Sludge and its Safe Disposal Options. Procedia Environ. Sci. 2016, 35, 950–955. [Google Scholar] [CrossRef]
  102. Jung, K.W.; Hwang, M.J.; Park, D.S.; Ahn, K.H. Comprehensive reuse of drinking water treatment residuals in coagulation and adsorption processes. J. Environ. Manag. 2016, 181, 425–434. [Google Scholar] [CrossRef] [PubMed]
  103. Xu, Y.; Chen, T.; Xu, R.; He, L.; Cui, F. Impact of recycling alum sludge on coagulation of low-turbidity source waters. Desalination Water Treat. 2015, 57, 6732–6739. [Google Scholar] [CrossRef]
  104. Suman, A.; Ahmad, T.; Ahmad, K. Dairy wastewater treatment using water treatment sludge as coagulant: A novel treatment approach. Environ. Dev. Sustain. 2017, 20, 1615–1625. [Google Scholar] [CrossRef]
  105. Chen, W.; Song, L.; Gan, N.; Li, L. Sorption, degradation and mobility of microcystins in Chinese agriculture soils: Risk assessment for groundwater protection. Environ. Pollut. 2006, 144, 752–758. [Google Scholar] [CrossRef] [PubMed]
  106. Clemente, A.; Wilson, A.; Oliveira, S.; Menezes, I.; Gois, A.; Capelo-Neto, J. The role of hydraulic conditions of coagulation and flocculation on the damage of cyanobacteria. Sci. Total Environ. 2020, 740, 139737. [Google Scholar] [CrossRef] [PubMed]
  107. Pestana, C.J.; Capelo-Neto, J.; Lawton, L.; Oliveira, S.; Carloto, I.; Linhares, H.P. The effect of water treatment unit processes on cyanobacterial trichome integrity. Sci. Total Environ. 2019, 659, 1403–1414. [Google Scholar] [CrossRef] [PubMed]
  108. Pei, H.; Xu, H.; Wang, J.; Jin, Y.; Xiao, H.; Ma, C.; Sun, J.; Li, H. 16S rRNA Gene amplicon sequencing reveals significant changes in microbial compositions during cyanobacteria-laden drinking water sludge storage. Environ. Sci. Technol. 2017, 51, 12774–12783. [Google Scholar] [CrossRef]
  109. Chorus, I.; Welker, M. Toxic Cyanobacteria in Water: A Guide to Their Public Health Consequences, Monitoring and Management, 2nd ed.; CRC Press: Boca Raton, FL, USA, 2021; p. 859. [Google Scholar]
  110. Chorus, I.; Bartram, J. Toxic Cyanobacteria in Water: A Guide to Their Public Health Consequences, Monitoring, and Management; WHO: London, UK; New York, NY, USA, 1999. [Google Scholar]
  111. Ellis, D. Guide D’intervention Pour les Propriétaires, les Exploitants ou les Concepteurs de Stations de Production D’eau Potablemunicipales aux Prises avec une Problématique de Fleurs D’eau de Cyanobactéries; Ministère du Développement durable, Environnement et Parcs: Québec, QC, Canada, 2009; p. 51.
  112. America Water Works Association (AWWA). Algae Source to Treatment. Manual of Water Supply Practices—M57, 1st ed.; America Water Works Association (AWWA): Denver, CO, USA, 2010; p. 481. [Google Scholar]
  113. Hawkins, P.R.; Holliday, J.; Kathuria, A.; Bowling, L. Change in cyanobacterial biovolume due to preservation by Lugol’s Iodine. Harmful Algae 2005, 4, 1033–1043. [Google Scholar] [CrossRef]
  114. Park, J.; Kim, Y.; Kim, M.; Lee, W.H.; Park, J.; Kim, Y.; Kim, M.; Lee, W.H. A novel method for cell counting of Microcystis colonies in water resources using a digital imaging flow cytometer and microscope. Environ. Eng. Res. 2018, 24, 397–403. [Google Scholar] [CrossRef]
  115. Zamyadi, A.; McQuaid, N.; Prévost, M.; Dorner, S. Monitoring of potentially toxic cyanobacteria using an online multi-probe in drinking water sources. J. Environ. Monit. 2012, 14, 579–588. [Google Scholar] [CrossRef]
  116. McQuaid, N.; Zamyadi, A.; Prévost, M.; Bird, D.F.; Dorner, S. Use of in vivo phycocyanin fluorescence to monitor potential microcystin-producing cyanobacterial biovolume in a drinking water source. J. Environ. Monit. 2011, 13, 455–463. [Google Scholar] [CrossRef] [PubMed]
  117. Gregor, J.; Marsalek, B.; Sipkova, H. Detection and estimation of potentially toxic cyanobacteria in raw water at the drinking water treatment plant by in vivo fluorescence method. Water Res. 2007, 41, 228–234. [Google Scholar] [CrossRef] [PubMed]
  118. Zamyadi, A.; Choo, F.; Newcombe, G.; Stuetz, R.; Henderson, R.K. A review of monitoring technologies for real-time management of cyanobacteria: Recent advances and future direction. TrAC Trends Anal. Chem. 2016, 85, 83–96. [Google Scholar] [CrossRef]
  119. Bowling, L.C.; Zamyadi, A.; Henderson, R.K. Assessment of in situ fluorometry to measure cyanobacterial presence in water bodies with diverse cyanobacterial populations. Water Res. 2016, 105, 22–33. [Google Scholar] [CrossRef] [PubMed]
  120. Thomson-Laing, G.; Puddick, J.; Wood, S.A. Predicting cyanobacterial biovolumes from phycocyanin fluorescence using a handheld fluorometer in the field. Harmful Algae 2020, 97, 101869. [Google Scholar] [CrossRef] [PubMed]
  121. Cotterill, V.; Hamilton, D.P.; Puddick, J.; Suren, A.; Wood, S.A. Phycocyanin sensors as an early warning system for cyanobacteria blooms concentrations: A case study in the Rotorua lakes. N. Z. J. Mar. Freshw. Res. 2019, 53, 555–570. [Google Scholar] [CrossRef]
  122. World Health Organization (WHO). Cyanobacterial Toxins: Microcystin-LR in Drinking-Water (Background Document for Development of WHO Guidelines for Drinking Water Quality); WHO/SDE/WSH/03.04/57; 2nd Edition Addendum to Volume 2; Health Critiria and Other Supporting Information; WHO: Geneva, Switzerland, 1998; p. 18. [Google Scholar]
  123. WHO. Cyanobacterial Toxins: Microcystins. Background Document for Development of WHO Guidelines for Drinking-Water Quality and Guidelines for Safe Recreational Water Environments; WHO/HEP/ECH/WSH/2020.6; World Health Organization: Geneva, Switzerland, 2020. [Google Scholar]
  124. Health Canada; Federal-Provincial-Territorial Commitee on Drinking Water. Cyanobacterial toxins in drinking water. Document for public consultation. Guidelines for Canadian Drinking Water Quality: Guideline Technical Document. 2016; 177. [Google Scholar]
  125. Ridal, J.; Brownlee, B.; McKenna, G.; Levac, N. Removal of taste and odour compounds by conventional granular activated carbon filtration. Water Qual. Res. J. Can. 2001, 36, 43–54. [Google Scholar] [CrossRef]
  126. Smith, K.M. Characterization of Activated Carbon for Taste and Odour Control. Master’s Thesis, University of Toronto, Toronto, ON, Canada, 2011. [Google Scholar]
  127. Hobson, P.; Fazekas, C.; House, J.; Daly, R.I.; Kildea, T.; Giglio, S.; Burch, M.; Lin, T.-F.; Chen, Y.-M. Tastes and Odours in Reservoirs; Water Quality Research Australia: Adelaide, Australia, 2010; p. 98. [Google Scholar]
  128. Kim, K.T.; Park, Y.-G. Geosmin and 2-MIB Removal by Full-Scale Drinking Water Treatment Processes in the Republic of Korea. Water 2021, 13, 628. [Google Scholar] [CrossRef]
  129. Zamyadi, A.; Henderson, R.; Stuetz, R.; Hofmann, R.; Ho, L.; Newcombe, G. Fate of geosmin and 2-methylisoborneol in full-scale water treatment plants. Water Res. 2015, 83, 171–183. [Google Scholar] [CrossRef]
  130. Sanseverino, I.; Conduto António, D.; Loos, R.; Lettieri, T. Cyanotoxins: Methods and Approaches for Their Analysis and Detection; EUR 28624; European Commission: Brussels, Belgium, 2017; p. 70. [Google Scholar]
  131. Picardo, M.; Filatova, D.; Nuñez, O.; Farré, M. Recent advances in the detection of natural toxins in freshwater environments. TrAC Trends Anal. Chem. 2019, 112, 75–86. [Google Scholar] [CrossRef]
  132. United States Environmental Protection Agency (USEPA). Method 546: Determination of Total Microcystins and Nodularins in Drinking Water and Ambient Water by Adda Enzyme-Linked Immunosorbent Assay; United States Environmental Protection Agency (USEPA): Washington, DC, USA, 2016. [Google Scholar]
  133. Young, W.F.; Horth, H.; Crane, R.; Ogden, T.; Arnott, M. Taste and odour threshold concentrations of potential potable water contaminants. Water Res. 1996, 30, 331–340. [Google Scholar] [CrossRef]
  134. Watson, S.B. Aquatic taste and odor: A primary signal of drinking-water integrity. J. Toxicol. Environ. Health Part A 2004, 67, 1779–1795. [Google Scholar] [CrossRef] [PubMed]
  135. Wert, E.C.; Korak, J.A.; Trenholm, R.A.; Rosario-Ortiz, F.L. Effect of oxidant exposure on the release of intracellular microcystin, MIB, and geosmin from three cyanobacteria species. Water Res. 2014, 52, 251–259. [Google Scholar] [CrossRef] [PubMed]
  136. Chong, S.; Lee, H.; An, K.-G. Predicting Taste and Odor Compounds in a Shallow Reservoir Using a Three–Dimensional Hydrodynamic Ecological Model. Water 2018, 10, 1396. [Google Scholar] [CrossRef] [Green Version]
  137. Bernhardt, H.; Clasen, J. Eutrophication control as an essential condition for an optimum disinfection. Water Supply 1993, 11, 89–108. [Google Scholar]
  138. Agence Française de Sécurité Sanitaire des Aliments (AFSSA); Agence Française de Sécurité Sanitaire de l’Environnement et du Travail (AFSSET). Avis relatif à l’évaluation des risques sanitaires liés à la présence de cyanobactéries dans les plans et cours d’eau destinés à la baignade et/ou à d’autres usages, Saisine AFSSET no 2004/008; France, Juillet. 2006; 2.
  139. Pietsch, J.; Bornmann, K.; Schmidt, W. Relevance of intra- and extracellular cyanotoxins for drinking water treatment. Acta Hydrochim. Hydrobiol. 2002, 30, 7–15. [Google Scholar] [CrossRef]
  140. Newcombe, G.; Brooke, S.; Cullum, P.; Nicholson, B.; Slyman, N. Oxidation, adsorption and biological treatment for algal toxin removal. In Proceedings of the American Water Works Association-Water Quality Technology Conference Seattle, Washington, DC, USA, 10–14 November 2002; p. 5. [Google Scholar]
  141. Newcombe, G. Removal of Algal Toxins from Drinking Water Using Ozone and GAC; 90904; American Water Works Association Research Foundation and American Water Works Association: Denver, CO, USA, 2002; p. 133. [Google Scholar]
  142. Newcombe, G.; Drikas, M. MIB removal: Adsorption capacity and kinetics of eight activated carbons. In Proceedings of the Australian Water and Wastewater Association WaterTECH Conference, Sydney, Australia, 27–28 May 1996; pp. 615–620. [Google Scholar]
  143. United States Environmental Protection Agency (USEPA). Cyanobacteria and Cyanotoxins: Information for Drinking Water Systems; United States Environmental Protection Agency (USEPA): Washington, DC, USA, 2019; p. 11.
  144. Lusty, M.W.; Gobler, C.J. The efficacy of hydrogen peroxide in mitigating cyanobacterial blooms and altering microbial communities across four lakes in NY, USA. Toxins 2020, 12, 428. [Google Scholar] [CrossRef]
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Figure 1. Cyanobacterial accumulation in two low risk DWTPs (maximum influx cell: <500 cells/mL). Only DWTP1 had pre-ozonation. The average values are from August to October 2011, adapted from [43].
Figure 1. Cyanobacterial accumulation in two low risk DWTPs (maximum influx cell: <500 cells/mL). Only DWTP1 had pre-ozonation. The average values are from August to October 2011, adapted from [43].
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Figure 2. Comparison of cell count reduction following ozonation (2 mg/L) in the (a) cultured Dolichospermum, Microcystis [64], (b) Natural bloom [65].
Figure 2. Comparison of cell count reduction following ozonation (2 mg/L) in the (a) cultured Dolichospermum, Microcystis [64], (b) Natural bloom [65].
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Figure 3. (a) Comparison of the cell viability results of cultured-based Microcystis and Dolichospermum [37,68,71,72,85] and natural cyanobacterial blooms [74,76,77] following pre-chlorination. [74] used Chl-a as a proxy for cell viability. (b) Cell viability experimental data and fitted model of unicellular [37,68,71,72,85] and colonial Microcystis [73] following pre-chlorination. (c) Cell viability experimental data and fitted model for three different cyanobacterial blooms following pre-chlorination.
Figure 3. (a) Comparison of the cell viability results of cultured-based Microcystis and Dolichospermum [37,68,71,72,85] and natural cyanobacterial blooms [74,76,77] following pre-chlorination. [74] used Chl-a as a proxy for cell viability. (b) Cell viability experimental data and fitted model of unicellular [37,68,71,72,85] and colonial Microcystis [73] following pre-chlorination. (c) Cell viability experimental data and fitted model for three different cyanobacterial blooms following pre-chlorination.
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Figure 4. (a) Comparison of the cell viability results of cultured-based Microcystis, Dolichospermum, Oscillatoria, Lyngby asp. [37,64,68,78,79], and natural blooms [65,76] following pre-ozonation. (b) Cell viability experimental data and fitted model for cultured-based and natural bloom samples following pre-ozonation.
Figure 4. (a) Comparison of the cell viability results of cultured-based Microcystis, Dolichospermum, Oscillatoria, Lyngby asp. [37,64,68,78,79], and natural blooms [65,76] following pre-ozonation. (b) Cell viability experimental data and fitted model for cultured-based and natural bloom samples following pre-ozonation.
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Figure 5. (a) Comparison of the cell viability results of cultured-based and natural bloom samples for different studies following potassium permanganate peroxidation: Microcystis [37,68], Microcystis, Dolichospermum [66,79], and natural blooms [76]. (b) Comparison of the cell viability results of cultured-based samples (Microcystis) following potassium permanganate peroxidation adapted from [81]. (c) Comparison of the cell viability results of natural bloom samples (from Lake Erie) adapted from [81].
Figure 5. (a) Comparison of the cell viability results of cultured-based and natural bloom samples for different studies following potassium permanganate peroxidation: Microcystis [37,68], Microcystis, Dolichospermum [66,79], and natural blooms [76]. (b) Comparison of the cell viability results of cultured-based samples (Microcystis) following potassium permanganate peroxidation adapted from [81]. (c) Comparison of the cell viability results of natural bloom samples (from Lake Erie) adapted from [81].
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Figure 6. Comparison of the cell viability results of cultured-based cyanobacterial cells (Microcystis, Pseudanabaena) [37,68,83,84] and natural bloom cells [76,86] after oxidation by hydrogen peroxide.
Figure 6. Comparison of the cell viability results of cultured-based cyanobacterial cells (Microcystis, Pseudanabaena) [37,68,83,84] and natural bloom cells [76,86] after oxidation by hydrogen peroxide.
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Figure 7. Summary of pre-oxidation (low and medium-high CT) impact on cyanobacteria/cyanotoxins and downflow processes (#: very low impact, +: increase, and -: decrease). Low CT for Cl2 = CT < 4 mg min/L, low CT for O3 = CT < 1 mg min/L, low CT for KMnO4 = CT < 50 mg min/L, and low CT for H2O2 = CT < 50 mg h/L.
Figure 7. Summary of pre-oxidation (low and medium-high CT) impact on cyanobacteria/cyanotoxins and downflow processes (#: very low impact, +: increase, and -: decrease). Low CT for Cl2 = CT < 4 mg min/L, low CT for O3 = CT < 1 mg min/L, low CT for KMnO4 = CT < 50 mg min/L, and low CT for H2O2 = CT < 50 mg h/L.
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Figure 8. Decision framework for cyanobacterial bloom management in drinking water treatment utilities.
Figure 8. Decision framework for cyanobacterial bloom management in drinking water treatment utilities.
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Table
Table 6. Data of cyanobacteria-laden sludge treatment. MIB: 2-Methylisoborneol.
Table 6. Data of cyanobacteria-laden sludge treatment. MIB: 2-Methylisoborneol.
Source of SludgeScaleTreatment Agent/DosageContact TimeInitial ConditionsCell Count ReductionMetabolite ReductionReference
Sludge thickenerLaboratory3 mg/L KMnO42 h5.0 × 104 cells/mL Pseudanabaena>95%-[57]
Laboratory10–100 mg/L PAC1 h100/L MIB-42–100% MIB
Full-scale10 mg/L KMnO415 h (max.)4.3 × 105 cells/mL
(natural blooms)		13–98% total and Pseudanabaena cell counts-
Full-scale10 mg/L KMnO4
20 mg/L PAC		KMnO4: 24–72 h
PAC: 1 h 		3.7 × 105 cells/mL
120 ng/L MIB
(natural blooms)		40–52% in total and Pseudanabaena cell counts20–22% MIB
Sludge holding tankLaboratory5 mg/L
KMnO4		60 min2.3–2.7 × 106 cells/mL
63–161 ng/L MCs
(natural blooms)		46–55% total cell counts0.3–24% MCs[59]
10 mg/L KMnO459–62% total cell counts2–32% MCs
10 mg/L
H2O2		24 h58% total cell counts27% MCs
20 mg/L
H2O2		77% total cell counts41% MCs
Full-scale
(shock oxidation)		10 mg/L KMnO424–72 h2.4 × 106 cells/mL
88–1083 ng/L MCs
(natural blooms)		24–43% total cell counts
(31% cell count increase after 48 h in one sample)		MCs:
3–25% decrease in one sample
37–589% increase in one sample
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Jalili, F.; Moradinejad, S.; Zamyadi, A.; Dorner, S.; Sauvé, S.; Prévost, M. Evidence-Based Framework to Manage Cyanobacteria and Cyanotoxins in Water and Sludge from Drinking Water Treatment Plants. Toxins 2022, 14, 410. https://doi.org/10.3390/toxins14060410

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Jalili F, Moradinejad S, Zamyadi A, Dorner S, Sauvé S, Prévost M. Evidence-Based Framework to Manage Cyanobacteria and Cyanotoxins in Water and Sludge from Drinking Water Treatment Plants. Toxins. 2022; 14(6):410. https://doi.org/10.3390/toxins14060410

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Jalili, Farhad, Saber Moradinejad, Arash Zamyadi, Sarah Dorner, Sébastien Sauvé, and Michèle Prévost. 2022. "Evidence-Based Framework to Manage Cyanobacteria and Cyanotoxins in Water and Sludge from Drinking Water Treatment Plants" Toxins 14, no. 6: 410. https://doi.org/10.3390/toxins14060410

APA Style

Jalili, F., Moradinejad, S., Zamyadi, A., Dorner, S., Sauvé, S., & Prévost, M. (2022). Evidence-Based Framework to Manage Cyanobacteria and Cyanotoxins in Water and Sludge from Drinking Water Treatment Plants. Toxins, 14(6), 410. https://doi.org/10.3390/toxins14060410

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