# Experimental testing of two urban stressors on freshwater biofilms

Romain Vrba, Isabelle Lavoie, Nicolas Creusot, Mélissa Eon, Débora Millan-Navarro, Agnès Feurtet-Mazel, Nicolas Mazzella, Aurélie Moreira, Dolors Planas, Soizic Morin

► To cite this version:

Romain Vrba, Isabelle Lavoie, Nicolas Creusot, Mélissa Eon, Débora Millan-Navarro, et al. Experimental testing of two urban stressors on freshwater biofilms. Aquatic Toxicology, 2024, 272, pp.106972. 10.1016/j.aquatox.2024.106972. hal-04613841

HAL Id: hal-04613841

https://hal.inrae.fr/hal-04613841

Submitted on 17 Jun 2024

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<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# 1 Experimental testing of two urban stressors on freshwater biofilms

2 Romain VRBA¹,², Isabelle LAVOIE², Nicolas CREUSOT¹, Mélissa EON¹, Débora MILLAN-NAVARRO¹,

3 Agnès FEURTET-MAZEL³, Nicolas MAZZELLA¹, Aurélie MOREIRA¹, Dolors PLANAS⁴, Soizic MORIN¹*

5 ¹ INRAE, UR EABX, 50 avenue de Verdun, 33612 Cestas cedex, France

6 ² INRS-ETE, 490 rue de la Couronne, Québec, QC G1K 9A9, Canada

7 ³ Univ. Bordeaux, UMR CNRS 5805 EPOC-OASU, F-336120 Arcachon, France

8 ⁴ UQAM, GRIL-Département des sciences biologiques, 141 Avenue du Président-Kennedy, Montréal,

9 QC H2X 1Y4, Canada

10 *Corresponding author at: INRAE, UR EABX, 50 avenue de Verdun, 33612 Cestas cedex, France.

11 Tel. : +33 6 95 43 84 28

12 E-mail address: soizic.morin@inrae.fr

# 14 Graphical abstract

<figure><img src="image_1.png" /></figure>

# 17 Abstract

18 Aquatic ecosystems and their communities are exposed to numerous stressors of various 19 natures (chemical and physical), whose impacts are often poorly documented. In urban areas, 20 the use of biocides such as dodecyldimethylbenzylammonium chloride (DDBAC) and their 21 subsequent release in wastewater result in their transfer to urban aquatic ecosystems. DDBAC 22 is known to be toxic to most aquatic organisms. Artificial light at night (ALAN) is another 23 stressor that is increasing globally, especially in urban areas. ALAN may have a negative impact 24 on photosynthetic cycles of periphytic biofilms, which in turn may result in changes in their 25 metabolic functioning. Moreover, studies suggest that exposure to artificial light could 26 increase the biocidal effect of DDBAC on biofilms. The present study investigates the 27 individual and combined effects of DDBAC and/or ALAN on the functioning and structure of 28 photosynthetic biofilms. We exposed biofilms in artificial channels to a nominal concentration 29 of 30 mg.L⁻¹ of DDBAC and/or ALAN for 10 days. ALAN modified DDBAC exposure, decreasing 30 concentrations in the water but not accumulation in biofilms. DDBAC had negative impacts on 31 biofilm functioning and structure. Photosynthetic activity was inhibited by > 90% after 2 days 32 of exposure, compared to the controls, and did not recover over the duration of the 33 experiment. Biofilm composition was also impacted, with a marked decrease in green algae 34 and the disappearance of microfauna under DDBAC exposure. The integrity of algal cells was 35 compromised where DDBAC exposure altered the chloroplasts and chlorophyll content.
# 1 Experimental testing of two urban stressors on freshwater biofilms

2 Romain VRBA¹,², Isabelle LAVOIE², Nicolas CREUSOT¹, Mélissa EON¹, Débora MILLAN-NAVARRO¹,

3 Agnès FEURTET-MAZEL³, Nicolas MAZZELLA¹, Aurélie MOREIRA¹, Dolors PLANAS⁴, Soizic MORIN¹*

5 ¹ INRAE, UR EABX, 50 avenue de Verdun, 33612 Cestas cedex, France

6 ² INRS-ETE, 490 rue de la Couronne, Québec, QC G1K 9A9, Canada

7 ³ Univ. Bordeaux, UMR CNRS 5805 EPOC-OASU, F-336120 Arcachon, France

8 ⁴ UQAM, GRIL-Département des sciences biologiques, 141 Avenue du Président-Kennedy, Montréal,

9 QC H2X 1Y4, Canada

10 *Corresponding author at: INRAE, UR EABX, 50 avenue de Verdun, 33612 Cestas cedex, France.

11 Tel. : +33 6 95 43 84 28

12 E-mail address: soizic.morin@inrae.fr

# 14 Graphical abstract

<figure><img src="image_2.png" /></figure>

# 17 Abstract

18 Aquatic ecosystems and their communities are exposed to numerous stressors of various 19 natures (chemical and physical), whose impacts are often poorly documented. In urban areas, 20 the use of biocides such as dodecyldimethylbenzylammonium chloride (DDBAC) and their 21 subsequent release in wastewater result in their transfer to urban aquatic ecosystems. DDBAC 22 is known to be toxic to most aquatic organisms. Artificial light at night (ALAN) is another 23 stressor that is increasing globally, especially in urban areas. ALAN may have a negative impact 24 on photosynthetic cycles of periphytic biofilms, which in turn may result in changes in their 25 metabolic functioning. Moreover, studies suggest that exposure to artificial light could 26 increase the biocidal effect of DDBAC on biofilms. The present study investigates the 27 individual and combined effects of DDBAC and/or ALAN on the functioning and structure of 28 photosynthetic biofilms. We exposed biofilms in artificial channels to a nominal concentration 29 of 30 mg.L⁻¹ of DDBAC and/or ALAN for 10 days. ALAN modified DDBAC exposure, decreasing 30 concentrations in the water but not accumulation in biofilms. DDBAC had negative impacts on 31 biofilm functioning and structure. Photosynthetic activity was inhibited by > 90% after 2 days 32 of exposure, compared to the controls, and did not recover over the duration of the 33 experiment. Biofilm composition was also impacted, with a marked decrease in green algae 34 and the disappearance of microfauna under DDBAC exposure. The integrity of algal cells was 35 compromised where DDBAC exposure altered the chloroplasts and chlorophyll content.
36

37

38

Impacts on autotrophs were also observed through a shift in lipid profiles, in particular a strong decrease in glycolipid content was noted. We found no significant interactive effect of ALAN and DDBAC on the studied endpoints.

39

**Keywords:** Biocide, ALAN, biofilm, photosynthetic efficiency, bioaccumulation, lipids.

40

# 1- Introduction

41

42

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56

Biocides are widely used in different areas of activity including agriculture and industry, as well as in a multitude of domestic products (e.g. pharmaceutical, personal care and household products) (Abbott et al., 2020). Among these biocides, dodecyldimethylbenzylammonium chloride (DDBAC) is commonly used as an active substance in medical disinfection products, as well as in household products such as detergents. Recently, following the regulation of some broadly used disinfectant agents such as triclosan, quaternary ammonium compounds like DDBAC have become popular substitutes (Sreevidya et al., 2018; Arnold et al., 2023). Moreover, DDBAC and all other benzalkonium chloride derivatives have been shown to be very effective against the SARS-CoV virus (Rabenau et al., 2005), resulting in their widespread use as disinfectants during the Covid 19 pandemic (US EPA, 2020). This multi-purpose product eventually reaches wastewaters, where its concentration varies greatly depending on the land use, population and wastewater treatment facilities, among other factors. For example, a concentration of 6 mg.L⁻¹ was recorded in hospital effluents (Kümmerer et al., 1997) due to the intensive use of DDBAC as a disinfectant. In municipal wastewater effluents, DDBAC concentrations were observed to range from just a few ng per liter up to 170 µg.L⁻¹ (Clara et al., 2007; Martínez-Carballo et al., 2007; Zhang et al., 2015).

57

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Once in the aquatic environment, DDBAC is relatively stable to photodegradation and its half-life in surface waters can reach 180 days, although the presence of photosensitizers (e.g. natural organic matter, Hora & Arnold 2020, Mohapatra et al., 2024) can reduce this half-life to a week (US EPA, 2006). The organic carbon-water partition coefficient ($K_{oc}$) of DDBAC is 5.43 and, therefore, it has a high adsorption ratio on sewage sludge, sediments or humic substances (van Wijk et al., 2009). Although the transfer of DDBAC to urban aquatic ecosystems has been recognized, the ecotoxicity of DDBAC is poorly documented. The toxicity of DDBAC to cells is due to its quaternary ammonium polar head, which has the capacity to bind to the surface membrane, while the alkyl lipophilic chain alters the phospholipid bilayer. This alteration can rapidly lead to membrane disruption and progressive lysis of the cell (Eich et al., 2000). Regarding aquatic organisms, there is redundancy in the biological models used for toxicity assessment where most studies have focused on *Daphnia magna* to determine the acute toxicity of DDBAC (Kreuzinger et al., 2007; Leal et al., 1994; Chen et al., 2014; Lavorgna et al., 2016). To date, *D. magna* is the most sensitive organism to DDBAC, where the concentration needed to immobilize 50% of organisms ($EC_{50}$) was of 5.9 µg.L⁻¹ (US EPA, 2006). As a result, the US EPA established a non-observed adverse effect concentration (NOAEC) of 4.15 µg.L⁻¹ for aquatic invertebrates. Phytoplanktonic organisms seem to be less sensitive to DDBAC than aquatic invertebrates. Indeed, other studies investigated the effects of DDBAC on various microalgae species and the range of $EC_{50}$ based on growth was found to vary between
36

37

38

Impacts on autotrophs were also observed through a shift in lipid profiles, in particular a strong decrease in glycolipid content was noted. We found no significant interactive effect of ALAN and DDBAC on the studied endpoints.

39

**Keywords:** Biocide, ALAN, biofilm, photosynthetic efficiency, bioaccumulation, lipids.

40

# 1- Introduction

41

42

43

44

45

46

47

48

49

50

51

52

53

54

55

56

Biocides are widely used in different areas of activity including agriculture and industry, as well as in a multitude of domestic products (e.g. pharmaceutical, personal care and household products) (Abbott et al., 2020). Among these biocides, dodecyldimethylbenzylammonium chloride (DDBAC) is commonly used as an active substance in medical disinfection products, as well as in household products such as detergents. Recently, following the regulation of some broadly used disinfectant agents such as triclosan, quaternary ammonium compounds like DDBAC have become popular substitutes (Sreevidya et al., 2018; Arnold et al., 2023). Moreover, DDBAC and all other benzalkonium chloride derivatives have been shown to be very effective against the SARS-CoV virus (Rabenau et al., 2005), resulting in their widespread use as disinfectants during the Covid 19 pandemic (US EPA, 2020). This multi-purpose product eventually reaches wastewaters, where its concentration varies greatly depending on the land use, population and wastewater treatment facilities, among other factors. For example, a concentration of 6 mg.L⁻¹ was recorded in hospital effluents (Kümmerer et al., 1997) due to the intensive use of DDBAC as a disinfectant. In municipal wastewater effluents, DDBAC concentrations were observed to range from just a few ng per liter up to 170 µg.L⁻¹ (Clara et al., 2007; Martínez-Carballo et al., 2007; Zhang et al., 2015).

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Once in the aquatic environment, DDBAC is relatively stable to photodegradation and its half-life in surface waters can reach 180 days, although the presence of photosensitizers (e.g. natural organic matter, Hora & Arnold 2020, Mohapatra et al., 2024) can reduce this half-life to a week (US EPA, 2006). The organic carbon-water partition coefficient ($K_{oc}$) of DDBAC is 5.43 and, therefore, it has a high adsorption ratio on sewage sludge, sediments or humic substances (van Wijk et al., 2009). Although the transfer of DDBAC to urban aquatic ecosystems has been recognized, the ecotoxicity of DDBAC is poorly documented. The toxicity of DDBAC to cells is due to its quaternary ammonium polar head, which has the capacity to bind to the surface membrane, while the alkyl lipophilic chain alters the phospholipid bilayer. This alteration can rapidly lead to membrane disruption and progressive lysis of the cell (Eich et al., 2000). Regarding aquatic organisms, there is redundancy in the biological models used for toxicity assessment where most studies have focused on *Daphnia magna* to determine the acute toxicity of DDBAC (Kreuzinger et al., 2007; Leal et al., 1994; Chen et al., 2014; Lavorgna et al., 2016). To date, *D. magna* is the most sensitive organism to DDBAC, where the concentration needed to immobilize 50% of organisms ($EC_{50}$) was of 5.9 µg.L⁻¹ (US EPA, 2006). As a result, the US EPA established a non-observed adverse effect concentration (NOAEC) of 4.15 µg.L⁻¹ for aquatic invertebrates. Phytoplanktonic organisms seem to be less sensitive to DDBAC than aquatic invertebrates. Indeed, other studies investigated the effects of DDBAC on various microalgae species and the range of $EC_{50}$ based on growth was found to vary between
77 58 µg.L⁻¹ for *Skeletonema costatum* (Kreuzinger et al., 2007) to 203 µg.L⁻¹ for *Chlorella vulgaris* (Sütterlin et al., 2008). However, to our knowledge, the chronic effects of DDBAC exposure on natural aquatic communities has not been considered.

80

81 In this study, we focused on stream biofilms (periphyton), which are complex structures
82 housing autotrophic microorganisms (e.g. cyanobacteria, green algae, diatoms), bacteria,
83 fungi, and other heterotrophic organisms such as microfauna (Battin et al., 2016). Biofilms are
84 ubiquitous in aquatic environments, including urban streams and ponds. They form the basis
85 of aquatic food webs, providing a source of energy for primary producers, including essential
86 fatty acids (Brett and Müller-Navarra, 1997). Biofilms can also be found downstream of
87 wastewater treatment plants (WWTPs) and have been used as ecological indicators of stress
88 (Tamminen et al., 2022; Tlili et al., 2020). The early detection of ecosystem impairment using
89 autotrophic biofilms can be based on functional (e.g., photosynthesis) and structural
90 endpoints describing biodiversity based on taxonomy as well as pigments or lipid profiles
91 (Sabater et al., 2007; Morin & Artigas, in press). Biofilms are also efficient bioaccumulators of
92 organic substances carried by waters under dissolved or particulate forms (Bonnineau et al.,
93 2021). Given the toxic mode of action of DDBAC, impacts on biofilm fatty acids, particularly on
94 membrane phospholipids, are likely to occur. The attack on phospholipid membrane may also
95 result in the disruption of key functions of biofilm such as photosynthesis. For example, Pozo-
96 Antonio and Sanmartin (2018) showed a significant decrease in chlorophyll *a* and
97 photosynthetic efficiency of phototrophic biofilms from church walls after a DDBAC
98 treatment. The effect of DDBAC as a biofouling removal agent was significantly enhanced
99 when combined with artificial light or UV irradiation.

100

101 Artificial light at night (ALAN) has become a global pollution concern and, as of 2014, more
102 than 20% of the world's land surfaces between 75°N and 60°S were exposed to a light-polluted
103 sky (Falchi et al., 2016). Most urban areas have developed along rivers and coastlines,
104 increasing the exposure of urban aquatic environments to ALAN. Concerns about the impact
105 of ALAN on aquatic ecosystems and research on this topic are quite recent (Perkin et al., 2011).
106 Indeed, ALAN was only recognized as harmful for freshwater ecosystems in the early 2000s
107 (Longcore and Rich, 2004). ALAN could lead to disruptions in nycthemeral cycles of
108 autotrophic organisms in biofilms by inducing variability in the maximum efficiency of
109 photosynthesis (Maggi and Serôdio, 2020). ALAN can also alter taxonomic composition by
110 favouring certain autotrophic groups over others, thereby exerting differential selection, as
111 shown in a study where cyanobacteria proportions in biofilm decreased under ALAN (Grubisic
112 et al., 2017).

113

114 The objective of this study was to determine the individual and combined effects of two
115 anthropogenic stressors typical from urban aquatic ecosystems (DDBAC and ALAN) on
116 autotrophic organisms within biofilms, using structural and functional descriptors of impact.
117 To this aim, we performed a 10-day experiment in which natural autotrophic biofilms were
118 exposed in laboratory channels to DDBAC and/or ALAN. We hypothesized that, given the Koc
77

58 µg.L⁻¹ for *Skeletonema costatum* (Kreuzinger et al., 2007) to 203 µg.L⁻¹ for *Chlorella vulgaris*

78

(Sütterlin et al., 2008). However, to our knowledge, the chronic effects of DDBAC exposure on
natural aquatic communities has not been considered.

79

80

81

In this study, we focused on stream biofilms (periphyton), which are complex structures housing autotrophic microorganisms (e.g. cyanobacteria, green algae, diatoms), bacteria, fungi, and other heterotrophic organisms such as microfauna (Battin et al., 2016). Biofilms are ubiquitous in aquatic environments, including urban streams and ponds. They form the basis of aquatic food webs, providing a source of energy for primary producers, including essential fatty acids (Brett and Müller-Navarra, 1997). Biofilms can also be found downstream of wastewater treatment plants (WWTPs) and have been used as ecological indicators of stress (Tamminen et al., 2022; Tlili et al., 2020). The early detection of ecosystem impairment using autotrophic biofilms can be based on functional (e.g., photosynthesis) and structural endpoints describing biodiversity based on taxonomy as well as pigments or lipid profiles (Sabater et al., 2007; Morin & Artigas, in press). Biofilms are also efficient bioaccumulators of organic substances carried by waters under dissolved or particulate forms (Bonnineau et al., 2021). Given the toxic mode of action of DDBAC, impacts on biofilm fatty acids, particularly on membrane phospholipids, are likely to occur. The attack on phospholipid membrane may also result in the disruption of key functions of biofilm such as photosynthesis. For example, Pozo-Antonio and Sanmartin (2018) showed a significant decrease in chlorophyll *a* and photosynthetic efficiency of phototrophic biofilms from church walls after a DDBAC treatment. The effect of DDBAC as a biofouling removal agent was significantly enhanced when combined with artificial light or UV irradiation.

82

83

84

85

86

87

88

89

90

91

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101

Artificial light at night (ALAN) has become a global pollution concern and, as of 2014, more
than 20% of the world's land surfaces between 75°N and 60°S were exposed to a light-polluted
sky (Falchi et al., 2016). Most urban areas have developed along rivers and coastlines,
increasing the exposure of urban aquatic environments to ALAN. Concerns about the impact
of ALAN on aquatic ecosystems and research on this topic are quite recent (Perkin et al., 2011).
Indeed, ALAN was only recognized as harmful for freshwater ecosystems in the early 2000s
(Longcore and Rich, 2004). ALAN could lead to disruptions in nycthemeral cycles of
autotrophic organisms in biofilms by inducing variability in the maximum efficiency of
photosynthesis (Maggi and Serôdio, 2020). ALAN can also alter taxonomic composition by
favouring certain autotrophic groups over others, thereby exerting differential selection, as
shown in a study where cyanobacteria proportions in biofilm decreased under ALAN (Grubisic
et al., 2017).

102

103

104

105

106

107

108

109

110

111

112

113

114

The objective of this study was to determine the individual and combined effects of two
anthropogenic stressors typical from urban aquatic ecosystems (DDBAC and ALAN) on
autotrophic organisms within biofilms, using structural and functional descriptors of impact.
To this aim, we performed a 10-day experiment in which natural autotrophic biofilms were
exposed in laboratory channels to DDBAC and/or ALAN. We hypothesized that, given the Koc

115

116

117

118
119 of the substance, DDBAC would bioaccumulate in the biofilms, generating toxicity to 120 microorganisms. Functional (photosynthetic efficiency) and structural (taxonomic 121 composition, pigment and lipid profiles) descriptors were predicted to be impacted by DDBAC 122 exposure. ALAN was expected to modify the development of microalgae, thus to impact the 123 proportions of algal groups, to alter photosynthetic efficiency to a certain extent, and to 124 increase the sensitivity of microalgae to additional stress. As a consequence, we hypothesized 125 that co-exposure to ALAN and DDBAC would enhance the toxic effects of the biocide. 126

126

127 **2- Material and methods**

128 **2.1- Experimental design**

129 Biofilms were grown in a DDBAC-free pond in Cestas, near Bordeaux, France. A previous study 130 classified this small water body as a hypereutrophic pond (Chaumet et al., 2019; Neury- 131 Ormanni et al., 2020). Glass slides (~200 cm²) were immersed in the photic zone, at a depth 132 of 30-50 cm. Colonization took place in winter (December 2020 to April 2021), during five 133 months to collect sufficient amounts of biomass. At this period, daylight irradiance on the top 134 surface on the water (5 cm deep) was below 100 µmol.s⁻¹.m⁻². At the end of the colonization 135 period, 36 glass slides covered by mature biofilms were randomly distributed among four 136 series of experimental units (EU) for the experiment. The EUs consisted of experimental 137 channels of about 30 L filled with pond water that had been previously filtered (20 µm) to 138 remove suspended material and most planktonic organisms. To specifically assess the impacts 139 of urban factors (DDBAC and ALAN), while avoiding possible inter-channel variability likely to 140 influence biofilm responses, three parallel channels were set up and connected to individual 141 10-L tanks (one per EU, making the slides pseudoreplicates for each condition). Biofilms 142 corresponding to day 0 (d0, 5 slides) were collected immediately after their recovery from the 143 pond. 144

144 On d0, two EUs were contaminated with a solution of dodecyldimethylbenzylammonium 145 chloride (DDBAC; Sigma Aldrich, France; CAS: 139-07-1, purity: >99%) to reach a concentration 146 of 30 mg.L⁻¹. This concentration was selected to elicit an effect on photochemical efficiency 147 while using as low a concentration as possible, based on the observed EC₅₀ of preliminary dose- 148 response experiments (Appendix A). In order to assess only the effect of altered photoperiod, 149 not light level, a non-stressful incident light intensity of 20 µmol.s⁻¹.m⁻² was selected. Light was 150 kept on overnight and then from day 1 (d1), one control series and one DDBAC series were 151 exposed to an alternating 14 h day / 10 h night photoperiod (alternated photoperiod, AP) and 152 the other two series (one control, one DDBAC) were exposed to a 24 h day photoperiod 153 (continuous photoperiod, CP) throughout the course of the experiment (Figure 1). Room 154 temperature was maintained constant at 20.5 ± 0.1°C, while water temperature was kept at 155 18.7 ± 0.2°C. The tanks were refilled with 3 L of filtered pond water contaminated at 30 mg.L⁻¹ 156 DDBAC on day 4 (d4) to compensate for water evaporation. 157
of the substance, DDBAC would bioaccumulate in the biofilms, generating toxicity to microorganisms. Functional (photosynthetic efficiency) and structural (taxonomic composition, pigment and lipid profiles) descriptors were predicted to be impacted by DDBAC exposure. ALAN was expected to modify the development of microalgae, thus to impact the proportions of algal groups, to alter photosynthetic efficiency to a certain extent, and to increase the sensitivity of microalgae to additional stress. As a consequence, we hypothesized that co-exposure to ALAN and DDBAC would enhance the toxic effects of the biocide.

126

## 2- Material and methods

### 2.1- Experimental design

Biofilms were grown in a DDBAC-free pond in Cestas, near Bordeaux, France. A previous study classified this small water body as a hypereutrophic pond (Chaumet et al., 2019; Neury-Ormanni et al., 2020). Glass slides (~200 cm²) were immersed in the photic zone, at a depth of 30-50 cm. Colonization took place in winter (December 2020 to April 2021), during five months to collect sufficient amounts of biomass. At this period, daylight irradiance on the top surface on the water (5 cm deep) was below 100 µmol·s⁻¹·m⁻². At the end of the colonization period, 36 glass slides covered by mature biofilms were randomly distributed among four series of experimental units (EU) for the experiment. The EUs consisted of experimental channels of about 30 L filled with pond water that had been previously filtered (20 µm) to remove suspended material and most planktonic organisms. To specifically assess the impacts of urban factors (DDBAC and ALAN), while avoiding possible inter-channel variability likely to influence biofilm responses, three parallel channels were set up and connected to individual 10-L tanks (one per EU, making the slides pseudoreplicates for each condition). Biofilms corresponding to day 0 (d0, 5 slides) were collected immediately after their recovery from the pond.

On d0, two EUs were contaminated with a solution of dodecyldimethylbenzylammonium chloride (DDBAC; Sigma Aldrich, France; CAS: 139-07-1, purity: >99%) to reach a concentration of 30 mg·L⁻¹. This concentration was selected to elicit an effect on photochemical efficiency while using as low a concentration as possible, based on the observed EC₅₀ of preliminary dose-response experiments (Appendix A). In order to assess only the effect of altered photoperiod, not light level, a non-stressful incident light intensity of 20 µmol·s⁻¹·m⁻² was selected. Light was kept on overnight and then from day 1 (d1), one control series and one DDBAC series were exposed to an alternating 14 h day / 10 h night photoperiod (alternated photoperiod, AP) and the other two series (one control, one DDBAC) were exposed to a 24 h day photoperiod (continuous photoperiod, CP) throughout the course of the experiment (Figure 1). Room temperature was maintained constant at 20.5 ± 0.1°C, while water temperature was kept at 18.7 ± 0.2°C. The tanks were refilled with 3 L of filtered pond water contaminated at 30 mg·L⁻¹ DDBAC on day 4 (d4) to compensate for water evaporation.
<figure><img src="image_2.png" /><figcaption>Figure 1: Experimental design and sampling strategy.</figcaption></figure>

<table><thead><tr><th rowspan="3"></th><th rowspan="3"></th><th colspan="6">DDBAC addition</th></tr><tr><th colspan="2">↓</th><th colspan="2">↓</th><th rowspan="2">d10</th></tr><tr><th>d0</th><th>d1</th><th>d2</th><th>d4</th><th>d7</th></tr></thead><tbody><tr><td rowspan="2"><strong>Water analyses</strong></td><td>Nutrients</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td>DDBAC</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td rowspan="2"><strong>Biofilm analyses</strong></td><td>DDBAC</td><td>✓</td><td>✓</td><td>✓</td><td></td><td></td><td>✓</td></tr><tr><td>Photosynthetic efficiency</td><td>✓</td><td></td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td>Microscopy</td><td></td><td>✓</td><td></td><td>✓</td><td></td><td></td><td>✓</td></tr><tr><td>Pigments*</td><td></td><td>✓</td><td>✓</td><td>✓</td><td></td><td></td><td>✓</td></tr><tr><td>Lipids</td><td></td><td>✓</td><td></td><td></td><td></td><td></td><td>✓</td></tr></tbody></table>

*Only performed on one replicate per treatment

## 2.2- Water chemistry measurements

On d0, d1, d2, d4, d7 and d10, 20 mL water samples were taken from all channels, filtered on 1-µm PTFE filters and stored at 4°C in the dark until analysis (performed within 48 h of collection). Nutrients and mineral salts were analysed as described in Chaumet et al. (2019), using a Metrohm 881 Compact Ionic Chromatograph pro (Metrohm). Anion analysis (PO₄⁻, NO₃⁻, NO₂⁻, Cl⁻ and SO₄²⁻) was performed using a Supp 4/5 Guard/4.0 precolumn followed by a Metrosep A Supp5 – 250/4.0 column. The mobile phase was a mixture of a solution of 3.2 mmol.L⁻¹ Na₂CO₃ and a solution of 1 mmol.L⁻¹ NaHCO₃. Cation analysis (Na⁺, K⁺, Ca²⁺, Mg²⁺ and NH₄⁺) was performed using a C4 Guard/4.0 precolumn followed by a Metrosep C6 - 250/4.0 column). All precolumns and columns were provided by Metrohm. The eluent used was a mixture of 2.5 mmol.L⁻¹ HNO₃ and a solution of 1.7 mmol.L⁻¹ 10,12-Pentacosadynoic acid (PCDA). The limits of quantification (LOQ) of the different ions analysed are shown in Table 1.

DDBAC concentrations in the water were monitored frequently over the experiment, at d0, d1, d2, d4, d7 and d10. Three samples of 20 mL were collected from each channel and stored at -20°C together with the stock solution until analysis. The samples were analysed using an Ultimate 3000 HPLC coupled with an API 2000 triple quadrupole mass spectrometer. We used a Gemini® NX-C18 column from Phenomenex as a stationary phase. The mobile phase was 90:10 5 mM ammonium acetate/acetonitrile. We worked in isocratic mode, so the composition of the mobile phase was constant during the analysis. The flow rate was set at 0.6 mL.min⁻¹ and the injection volume was set at 20 µL. An internal standard of benzyl-2,3,4,5,6-d5-dimethyl-n-dodecylammonium chloride (DDBAC-d5) was used (Cluzeau, France; CAS: 139-07-1, purity: >98%) at an equivalent of 100 µg.L⁻¹ in both samples and standard solutions for calibration. The mass spectrometer was operated in multiple reaction monitoring (MRM) mode with a total cycle time of 0.8 seconds. The MRM transitions were 304>91 (quantitation) and 304>212 (confirmation) for DDBAC. For DDBAC-d5, the MRM transitions were 309>96 and 309>212. The declustering potential was set to 65 V and the collision
<figure><img src="image_3.png" /><figcaption>Figure 1: Experimental design and sampling strategy.</figcaption></figure>

<table><thead><tr><th rowspan="3"></th><th rowspan="3"></th><th colspan="6">DDBAC addition</th></tr><tr><th colspan="2">↓</th><th colspan="2">↓</th><th rowspan="2">d10</th></tr><tr><th>d0</th><th>d1</th><th>d2</th><th>d4</th><th>d7</th></tr></thead><tbody><tr><td rowspan="2">Water analyses</td><td>Nutrients</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td>DDBAC</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td rowspan="2">Biofilm analyses</td><td>DDBAC</td><td>✓</td><td>✓</td><td>✓</td><td></td><td></td><td>✓</td></tr><tr><td>Photosynthetic efficiency</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td><td>✓</td></tr><tr><td>Microscopy</td><td></td><td>✓</td><td>✓</td><td></td><td></td><td></td><td>✓</td></tr><tr><td>Pigments*</td><td></td><td>✓</td><td>✓</td><td>✓</td><td></td><td></td><td>✓</td></tr><tr><td>Lipids</td><td></td><td>✓</td><td></td><td></td><td></td><td></td><td>✓</td></tr></tbody></table>

*Only performed on one replicate per treatment

## 2.2- Water chemistry measurements

On d0, d1, d2, d4, d7 and d10, 20 mL water samples were taken from all channels, filtered on 1-µm PTFE filters and stored at 4°C in the dark until analysis (performed within 48 h of collection). Nutrients and mineral salts were analysed as described in Chaumet et al. (2019), using a Metrohm 881 Compact Ionic Chromatograph pro (Metrohm). Anion analysis (PO₄³⁻, NO₃⁻, NO₂⁻, Cl⁻ and SO₄²⁻) was performed using a Supp 4/5 Guard/4.0 precolumn followed by a Metrosep A Supp5 – 250/4.0 column. The mobile phase was a mixture of a solution of 3.2 mmol.L⁻¹ Na₂CO₃ and a solution of 1 mmol.L⁻¹ NaHCO₃. Cation analysis (Na⁺, K⁺, Ca²⁺, Mg²⁺ and NH₄⁺) was performed using a C4 Guard/4.0 precolumn followed by a Metrosep C6 - 250/4.0 column). All precolumns and columns were provided by Metrohm. The eluent used was a mixture of 2.5 mmol.L⁻¹ HNO₃ and a solution of 1.7 mmol.L⁻¹ 10,12-Pentacosadynoic acid (PCDA). The limits of quantification (LOQ) of the different ions analysed are shown in Table 1.

DDBAC concentrations in the water were monitored frequently over the experiment, at d0, d1, d2, d4, d7 and d10. Three samples of 20 mL were collected from each channel and stored at -20°C together with the stock solution until analysis. The samples were analysed using an Ultimate 3000 HPLC coupled with an API 2000 triple quadrupole mass spectrometer. We used a Gemini® NX-C18 column from Phenomenex as a stationary phase. The mobile phase was 90:10 5 mM ammonium acetate/acetonitrile. We worked in isocratic mode, so the composition of the mobile phase was constant during the analysis. The flow rate was set at 0.6 mL.min⁻¹ and the injection volume was set at 20 µL. An internal standard of benzyl-2,3,4,5,6-d5-dimethyl-n-dodecylammonium chloride (DDBAC-d5) was used (Cluzeau, France; CAS: 139-07-1, purity: >98%) at an equivalent of 100 µg.L⁻¹ in both samples and standard solutions for calibration. The mass spectrometer was operated in multiple reaction monitoring (MRM) mode with a total cycle time of 0.8 seconds. The MRM transitions were 304>91 (quantitation) and 304>212 (confirmation) for DDBAC. For DDBAC-d5, the MRM transitions were 309>96 and 309>212. The declustering potential was set to 65 V and the collision
energies were set either to 45 or 30 V for the quantitation or confirmation transition, respectively. Samples were diluted and the calibration range was from 1 to 200 µg.L⁻¹. Quality controls were regularly injected at concentrations of 5 and 25 µg.L⁻¹, as well as analytical blanks.

190

## 2.3- Biofilm biological endpoints

### 2.3.1- Photosynthetic efficiency

We sampled 2 cm² of biofilm from three different glass slides for each treatment on d0, d2, d4, d7, and d10, corresponding to three pseudoreplicates per treatment per sampling. Each biofilm sample was then suspended in 3 mL of channel water and gently shaken for homogenization. Photosynthetic activity (effective photosystem II quantum yield) was assessed on suspensions of biofilm (Genty et al. 1989) within one hour after collection using a Pulse Amplitude Modulation fluorimeter (Phyto-PAM, Heinz Walz GmbH, Germany) in quartz cuvettes under continuous agitation (Emitter-Detector Unit PHYTO-ED). Biofilm suspensions were adapted for 15 min to light (164 µmol.m⁻².s⁻¹) before fluorescence measurements at the same active radiation, in Actinic Light mode. The quantum yield values reported are mean values of five measurements per replicate, averaged for the three pigment groups using the device's reference spectra. After photochemical efficiency measurements, samples were preserved for further microscopic analyses by adding a few drops of Lugol solution then stored in the dark at 4°C.

### 2.3.2- Microscopic analyses

For microscopic observations, a Nageotte counting slide (Marienfeld, Germany) was used with 125 µL biofilm suspension samples collected on d0, d2 and d10 to determine initial taxonomic composition as well as to assess early structural changes and chronic effects of ALAN and DDBAC. Observations were made at x400 magnification under an optical microscope (Olympus BX51) equipped with a digital camera as described in Morin et al. (2010). Ten fields of view were scanned for enumeration of the main taxonomic groups observable on preserved material: diatoms, green algae, cyanobacteria and microfauna. Solitary and colonial algae and cyanobacteria were recorded and counted as cell numbers to allow for density comparisons. Microfauna organisms were recorded as individuals. All densities were expressed as a function of colonized biofilm surface (i.e., per cm²). Live (intact cell content) and dead (empty frustules) diatoms were counted separately to estimate mortality (Morin et al. 2010).

### 2.3.3- Analyses from freeze-dried biofilms

Biofilms were scraped from glass slides on d0, d1, d2 and d10 (4 slides per treatment) and were frozen in liquid nitrogen to prevent lipid degradation. The samples were then lyophilized for 24 hours, and used to determine lipid content, DDBAC bioaccumulation, and concentrations of photosynthetic pigments. Analyses focused on neutral lipids (storage lipids: triacylglycerides) and on polar lipids. Polar lipids correspond to phospholipids (lipids typical from cytoplasmic membranes: phosphatidylcholine-PC, phosphatidylethanolamine-PE, and phosphatidylglycerol-PG) and to glycolipids (characteristic from chloroplastic membranes:
energies were set either to 45 or 30 V for the quantitation or confirmation transition, respectively. Samples were diluted and the calibration range was from 1 to 200 µg.L⁻¹. Quality controls were regularly injected at concentrations of 5 and 25 µg.L⁻¹, as well as analytical blanks.

190

## 2.3- Biofilm biological endpoints

### 2.3.1- Photosynthetic efficiency

We sampled 2 cm² of biofilm from three different glass slides for each treatment on d0, d2, d4, d7, and d10, corresponding to three pseudoreplicates per treatment per sampling. Each biofilm sample was then suspended in 3 mL of channel water and gently shaken for homogenization. Photosynthetic activity (effective photosystem II quantum yield) was assessed on suspensions of biofilm (Genty et al. 1989) within one hour after collection using a Pulse Amplitude Modulation fluorimeter (Phyto-PAM, Heinz Walz GmbH, Germany) in quartz cuvettes under continuous agitation (Emitter-Detector Unit PHYTO-ED). Biofilm suspensions were adapted for 15 min to light (164 µmol.m⁻².s⁻¹) before fluorescence measurements at the same active radiation, in Actinic Light mode. The quantum yield values reported are mean values of five measurements per replicate, averaged for the three pigment groups using the device's reference spectra. After photochemical efficiency measurements, samples were preserved for further microscopic analyses by adding a few drops of Lugol solution then stored in the dark at 4°C.

### 2.3.2- Microscopic analyses

For microscopic observations, a Nageotte counting slide (Marienfeld, Germany) was used with 125 µL biofilm suspension samples collected on d0, d2 and d10 to determine initial taxonomic composition as well as to assess early structural changes and chronic effects of ALAN and DDBAC. Observations were made at x400 magnification under an optical microscope (Olympus BX51) equipped with a digital camera as described in Morin et al. (2010). Ten fields of view were scanned for enumeration of the main taxonomic groups observable on preserved material: diatoms, green algae, cyanobacteria and microfauna. Solitary and colonial algae and cyanobacteria were recorded and counted as cell numbers to allow for density comparisons. Microfauna organisms were recorded as individuals. All densities were expressed as a function of colonized biofilm surface (i.e., per cm²). Live (intact cell content) and dead (empty frustules) diatoms were counted separately to estimate mortality (Morin et al. 2010).

### 2.3.3- Analyses from freeze-dried biofilms

Biofilms were scraped from glass slides on d0, d1, d2 and d10 (4 slides per treatment) and were frozen in liquid nitrogen to prevent lipid degradation. The samples were then lyophilized for 24 hours, and used to determine lipid content, DDBAC bioaccumulation, and concentrations of photosynthetic pigments. Analyses focused on neutral lipids (storage lipids: triacylglycerides) and on polar lipids. Polar lipids correspond to phospholipids (lipids typical from cytoplasmic membranes: phosphatidylcholine-PC, phosphatidylethanolamine-PE, and phosphatidylglycerol-PG) and to glycolipids (characteristic from chloroplastic membranes:
226 digalactosyldiacylglycerol-DGDG, monogalactosyldiacylglycerol-MGDG,
227 sulfoquinovosyldiacylglycerol-SQDG). Lipids were extracted from the biofilms (20 mg dry
228 weight) using a mixture of MTBE-methanol (3:1 %v/v) and UPW-methanol (3:1 %v/v) with 150
229 mg of microbeads. Samples were then homogenized using a FastPrep (MP Biomedicals). The
230 upper organic fraction of the samples (i.e. MTBE) was recovered by centrifugation for lipid
231 analysis, while the lower hydrophilic fraction (mixture of UPW and methanol) was used to
232 determine DDBAC bioaccumulation.

233 **2.3.3.1- Lipid quantification**

234 For lipid analysis, samples from d0 and d10 were evaporated and injection solvent was added
235 before analysis by HPLC-MS/MS following the protocol detailed in Mazzella et al. (2023a).
236 Briefly, different stationary and mobile phases were used for the analysis of
237 phospholipids/glycolipids and triglycerides. For the phospholipid and glycolipid analyses, a
238 LUNA® NH2 HILIC column (100 x 2 mm, 3 µm) from Phenomenex was used as the stationary
239 phase, and a mixture of acetonitrile and 40 mM ammonium acetate buffer as the mobile
240 phase. The flow rate was set at 400 µL.min⁻¹. The proportions of these two solutions are given
241 in Appendix C. For the analysis of triglycerides and betaine lipids (i.e.
242 diacylglyceryltrimethylhomo-Ser, DGTS), a KINETEX® C8 column (100 x 2.1 mm, 2.6 µm) from
243 Phenomenex was used as the stationary phase, and the mobile phase was a mix of a solution
244 of acetonitrile/water/40 mM ammonium acetate buffer (600/390/10, v/v/v) and a solution of
245 isopropanol/acetonitrile/1 M ammonium acetate buffer (900/90/10, v/v/v). The flow rate was
246 set at 300 µL.min⁻¹. The proportions of these two solutions are given in Appendix C. Results
247 were then pre-treated with ANALYST® 1.6.2 software from Sciex. For polar lipids (i.e.
248 glycolipids and phospholipids), the limits of quantification ranged from 0.1 to 0.5 nmol.mg⁻¹,
249 depending on the lipid classes. For triglycerides and DGTS, the limits of quantification reached
250 were 0.01 and 0.1 nmol.mg⁻¹, respectively. Results were expressed as nmol.g⁻¹ freeze-dried
251 biofilm.

252 **2.3.3.2- DDBAC accumulation in biofilms**

253 The same analytical method used for water samples was used to determine bioaccumulated
254 DDBAC in the previously extracted hydrophilic fraction. It was generally necessary to perform
255 significant dilutions (i.e. 10,000-fold) in order to stay within the calibration range. Results were
256 then expressed in the log10 value of the bioconcentration factor (i.e. log(BCF)). The BCF was
257 calculated according to the following formula:

$$
258 \quad BCF = \frac{\text{Concentration of DDBAC in biofilm (mg.kg}^{-1}\text{)}}{\text{Concentration of DDBAC in water (mg.L}^{-1}\text{)}} \quad \text{(Eq.1)}
$$

259 **2.3.3.3- Photosynthetic pigments**

260 On selected freeze-dried samples (one replicate per treatment), pigment analyses were also
261 performed following the French standard NF T90-117 (1999). Ten mg of dry biofilm were put
262 in solution using 10 mL of acetone. After 20 minutes of ultrasonication, the mix was filtered
263 on a Büchner filter to remove the solid phase. Absorption was measured at the wavelengths
264 630 nm, 647 nm, 664 nm, 665 nm and 750 nm with a UV-1800 (Shimadzu) spectrophotometer,
265 and was then remeasured at the same wavelengths after acidification of the samples. The
226 digalactosyldiacylglycerol-DGDG, monogalactosyldiacylglycerol-MGDG,
227 sulfoquinovosyldiacylglycerol-SQDG). Lipids were extracted from the biofilms (20 mg dry
228 weight) using a mixture of MTBE-methanol (3:1 %v/v) and UPW-methanol (3:1 %v/v) with 150
229 mg of microbeads. Samples were then homogenized using a FastPrep (MP Biomedicals). The
230 upper organic fraction of the samples (i.e. MTBE) was recovered by centrifugation for lipid
231 analysis, while the lower hydrophilic fraction (mixture of UPW and methanol) was used to
232 determine DDBAC bioaccumulation.

233 **2.3.3.1- Lipid quantification**

234 For lipid analysis, samples from d0 and d10 were evaporated and injection solvent was added
235 before analysis by HPLC-MS/MS following the protocol detailed in Mazzella et al. (2023a).
236 Briefly, different stationary and mobile phases were used for the analysis of
237 phospholipids/glycolipids and triglycerides. For the phospholipid and glycolipid analyses, a
238 LUNA® NH2 HILIC column (100 x 2 mm, 3 µm) from Phenomenex was used as the stationary
239 phase, and a mixture of acetonitrile and 40 mM ammonium acetate buffer as the mobile
240 phase. The flow rate was set at 400 µL.min⁻¹. The proportions of these two solutions are given
241 in Appendix C. For the analysis of triglycerides and betaine lipids (i.e.
242 diacylglyceryltrimethylhomo-Ser, DGTS), a KINETEX® C8 column (100 x 2.1 mm, 2.6 µm) from
243 Phenomenex was used as the stationary phase, and the mobile phase was a mix of a solution
244 of acetonitrile/water/40 mM ammonium acetate buffer (600/390/10, v/v/v) and a solution of
245 isopropanol/acetonitrile/1 M ammonium acetate buffer (900/90/10, v/v/v). The flow rate was
246 set at 300 µL.min⁻¹. The proportions of these two solutions are given in Appendix C. Results
247 were then pre-treated with ANALYST® 1.6.2 software from Sciex. For polar lipids (i.e.
248 glycolipids and phospholipids), the limits of quantification ranged from 0.1 to 0.5 nmol.mg⁻¹,
249 depending on the lipid classes. For triglycerides and DGTS, the limits of quantification reached
250 were 0.01 and 0.1 nmol.mg⁻¹, respectively. Results were expressed as nmol.g⁻¹ freeze-dried
251 biofilm.

252 **2.3.3.2- DDBAC accumulation in biofilms**

253 The same analytical method used for water samples was used to determine bioaccumulated
254 DDBAC in the previously extracted hydrophilic fraction. It was generally necessary to perform
255 significant dilutions (i.e. 10,000-fold) in order to stay within the calibration range. Results were
256 then expressed in the log10 value of the bioconcentration factor (i.e. log(BCF)). The BCF was
257 calculated according to the following formula:

$$
258 \quad BCF = \frac{\text{Concentration of DDBAC in biofilm (mg.kg}^{-1})}{\text{Concentration of DDBAC in water (mg.L}^{-1})} \quad (\text{Eq.1})
$$

259 **2.3.3.3- Photosynthetic pigments**

260 On selected freeze-dried samples (one replicate per treatment), pigment analyses were also
261 performed following the French standard NF T90-117 (1999). Ten mg of dry biofilm were put
262 in solution using 10 mL of acetone. After 20 minutes of ultrasonication, the mix was filtered
263 on a Büchner filter to remove the solid phase. Absorption was measured at the wavelengths
264 630 nm, 647 nm, 664 nm, 665 nm and 750 nm with a UV-1800 (Shimadzu) spectrophotometer,
265 and was then remeasured at the same wavelengths after acidification of the samples. The
concentrations of chlorophyll pigments and phaeopigments were determined following the equations of Lorenzen (1967) and expressed as µg pigments.mg⁻¹ freeze-dried biofilm.

## 2.4- Data analyses

ANOVAs were performed after verification that homogeneity of variances and normality were met. A one-way ANOVA was used to test the effect of ALAN on DDBAC degradation, and two-way ANOVAs were used to assess the individual effect of ALAN and DDBAC, as well as their interaction, on nutrient concentrations. Two-way ANOVAs with sampling dates as repeated measures were conducted to assess the individual and interactive effects of ALAN and DDBAC (categorical variables with 2 levels: yes/no) on biofilm endpoints. Data were processed using R software (R Core Team, 2022).
concentrations of chlorophyll pigments and phaeopigments were determined following the equations of Lorenzen (1967) and expressed as µg pigments.mg⁻¹ freeze-dried biofilm.

## 2.4- Data analyses

ANOVAs were performed after verification that homogeneity of variances and normality were met. A one-way ANOVA was used to test the effect of ALAN on DDBAC degradation, and two-way ANOVAs were used to assess the individual effect of ALAN and DDBAC, as well as their interaction, on nutrient concentrations. Two-way ANOVAs with sampling dates as repeated measures were conducted to assess the individual and interactive effects of ALAN and DDBAC (categorical variables with 2 levels: yes/no) on biofilm endpoints. Data were processed using R software (R Core Team, 2022).
277 **3- Results**

278 **3.1- Water chemistry**

<table><caption>279 Table 1. Principal ions and DDBAC concentrations (mg.L⁻¹) in the four experimental treatments along the ten days of exposure. Results are expressed as the mean and
280 standard error for the whole experiment. Two-way ANOVAs were conducted, with ALAN and DDBAC as the two factors, except for DDBAC concentrations (One-way
281 ANOVA assessing the effect of ALAN). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = alternated photoperiod; CP = continuous photoperiod; LOQ =
282 Limit of quantification.</caption><thead><tr><th>Treatment</th><th>DDBAC</th><th>NO<sub>2</sub><sup>-</sup></th><th>NO<sub>3</sub><sup>-</sup></th><th>PO<sub>4</sub><sup>3-</sup></th><th>SO<sub>4</sub><sup>2-</sup></th><th>Cl<sup>-</sup></th><th>NH<sub>4</sub><sup>+</sup></th><th>Na<sup>+</sup></th><th>K<sup>+</sup></th><th>Ca<sup>2+</sup></th><th>Mg<sup>2+</sup></th></tr></thead><tbody><tr><td>Control-AP</td><td>&lt;LOQ</td><td>0.04 ± 0.03</td><td>1.69 ± 0.27</td><td>0.03 ± 0.02</td><td>21.0 ± 1.8</td><td>36.4 ± 2.7</td><td>0.09 ± 0.05</td><td>22.0 ± 1.9</td><td>4.80 ± 0.43</td><td>57.8 ± 5.1</td><td>4.90 ± 0.42</td></tr><tr><td>DDBAC-AP</td><td>26.6 ± 2.5</td><td>0.02 ± 0.00</td><td>0.66 ± 0.17</td><td>0.03 ± 0.02</td><td>18.9 ± 0.8</td><td>35.5 ± 1.4</td><td>0.11 ± 0.05</td><td>19.7 ± 0.9</td><td>4.34 ± 0.20</td><td>52.2 ± 2.5</td><td>4.41 ± 0.20</td></tr><tr><td>Control-CP</td><td>&lt;LOQ</td><td>0.02 ± 0.01</td><td>1.35 ± 0.23</td><td>0.01 ± 0.00</td><td>17.5 ± 0.3</td><td>30.3 ± 0.4</td><td>0.09 ± 0.06</td><td>18.5 ± 0.3</td><td>4.12 ± 0.06</td><td>48.4 ± 0.6</td><td>4.09 ± 0.06</td></tr><tr><td>DDBAC-CP</td><td>18.5 ± 2.0</td><td>0.02 ± 0.00</td><td>0.75 ± 0.10</td><td>0.03 ± 0.02</td><td>17.8 ± 0.3</td><td>33.0 ± 0.6</td><td>0.10 ± 0.06</td><td>18.8 ± 0.9</td><td>4.21 ± 0.11</td><td>49.5 ± 1.2</td><td>4.18 ± 0.09</td></tr><tr><td>LOQ</td><td>0.001</td><td>0.005</td><td>0.01</td><td>0.01</td><td>0.005</td><td>0.01</td><td>0.005</td><td>0.01</td><td>0.0025</td><td>0.15</td><td>0.15</td></tr><tr><td>Significant factor(s) and interaction(s)</td><td>ALAN</td><td>None</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>DDBAC</td></tr><tr><td>p</td><td>&lt;0.001</td><td>&gt;0.05</td><td>&lt;0.001</td><td>NA</td><td>&lt;0.05</td><td>&lt;0.01</td><td>NA</td><td>&lt;0.05</td><td>NA</td><td>&lt;0.05</td><td>&lt;0.05</td></tr></tbody></table>
277 **3- Results**

278 **3.1- Water chemistry**

<table><caption>279 Table 1. Principal ions and DDBAC concentrations (mg.L⁻¹) in the four experimental treatments along the ten days of exposure. Results are expressed as the mean and
280 standard error for the whole experiment. Two-way ANOVAs were conducted, with ALAN and DDBAC as the two factors, except for DDBAC concentrations (One-way
281 ANOVA assessing the effect of ALAN). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = alternated photoperiod; CP = continuous photoperiod; LOQ =
282 Limit of quantification.</caption><thead><tr><th>Treatment</th><th>DDBAC</th><th>NO<sub>2</sub><sup>-</sup></th><th>NO<sub>3</sub><sup>-</sup></th><th>PO<sub>4</sub><sup>3-</sup></th><th>SO<sub>4</sub><sup>2-</sup></th><th>Cl<sup>-</sup></th><th>NH<sub>4</sub><sup>+</sup></th><th>Na<sup>+</sup></th><th>K<sup>+</sup></th><th>Ca<sup>2+</sup></th><th>Mg<sup>2+</sup></th></tr></thead><tbody><tr><td>Control-AP</td><td>&lt;LOQ</td><td>0.04 ± 0.03</td><td>1.69 ± 0.27</td><td>0.03 ± 0.02</td><td>21.0 ± 1.8</td><td>36.4 ± 2.7</td><td>0.09 ± 0.05</td><td>22.0 ± 1.9</td><td>4.80 ± 0.43</td><td>57.8 ± 5.1</td><td>4.90 ± 0.42</td></tr><tr><td>DDBAC-AP</td><td>26.6 ± 2.5</td><td>0.02 ± 0.00</td><td>0.66 ± 0.17</td><td>0.03 ± 0.02</td><td>18.9 ± 0.8</td><td>35.5 ± 1.4</td><td>0.11 ± 0.05</td><td>19.7 ± 0.9</td><td>4.34 ± 0.20</td><td>52.2 ± 2.5</td><td>4.41 ± 0.20</td></tr><tr><td>Control-CP</td><td>&lt;LOQ</td><td>0.02 ± 0.01</td><td>1.35 ± 0.23</td><td>0.01 ± 0.00</td><td>17.5 ± 0.3</td><td>30.3 ± 0.4</td><td>0.09 ± 0.06</td><td>18.5 ± 0.3</td><td>4.12 ± 0.06</td><td>48.4 ± 0.6</td><td>4.09 ± 0.06</td></tr><tr><td>DDBAC-CP</td><td>18.5 ± 2.0</td><td>0.02 ± 0.00</td><td>0.75 ± 0.10</td><td>0.03 ± 0.02</td><td>17.8 ± 0.3</td><td>33.0 ± 0.6</td><td>0.10 ± 0.06</td><td>18.8 ± 0.9</td><td>4.21 ± 0.11</td><td>49.5 ± 1.2</td><td>4.18 ± 0.09</td></tr><tr><td>LOQ</td><td>0.001</td><td>0.005</td><td>0.01</td><td>0.01</td><td>0.005</td><td>0.01</td><td>0.005</td><td>0.01</td><td>0.0025</td><td>0.15</td><td>0.15</td></tr><tr><td>Significant factor(s) and interaction(s)</td><td>ALAN</td><td>None</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>NA</td><td>DDBAC</td><td>DDBAC</td></tr><tr><td>p</td><td>&lt;0.001</td><td>&gt;0.05</td><td>&lt;0.001</td><td>NA</td><td>&lt;0.05</td><td>&lt;0.01</td><td>NA</td><td>&lt;0.05</td><td>NA</td><td>&lt;0.05</td><td>&lt;0.05</td></tr></tbody></table>
In the control channels, DDBAC concentrations were always below quantification limit (Table 1), confirming that no cross-contamination occurred. Contaminant concentrations averaged 26.6 ± 2.5 mg.L⁻¹ (AP) and 18.5 ± 2.0 mg.L⁻¹ (CP) in the DDBAC treatments. Differences in DDBAC concentrations were observed between the DDBAC treatment channels exposed to an alternated photoperiod and those exposed to a continuous photoperiod (F[1,10] = 232.72; p < 0.001, also see Figure 1), suggesting that degradation occurred under continuous photoperiod. Nutrient concentrations were also modified by DDBAC exposure (Table 1). Indeed, DDBAC addition resulted in lower concentrations of nitrate, sulfate, Cl⁻, Na⁺, Ca²⁺ and Mg⁺ than in the control (Table 1), while ALAN showed no effect on nutrient concentrations.

293

## 3.2- Bioaccumulation of DDBAC in biofilms

<figure><img src="image_3.png" /><figcaption>Figure 2. Evolution of DDBAC bioaccumulated in biofilms (expressed in log(BCF)) between the different ALAN treatments (n=3). AP = alternated photoperiod CP = continuous photoperiod</figcaption></figure>

297

Figure 2 shows the log(BCF) for the hydrophilic fraction of biofilm samples at d1, d2 and d10 for AP and CP treatments. DDBAC was recovered from biofilms for every treatment and every sampling time, showing that this compound bioaccumulates well in autotrophic biofilms, with a mean log(BCF) of 3.22 ± 0.25. The amount of DDBAC bioaccumulated in the biofilms showed no significant changes between sampling times (F[2,28]= 2.38; p = 0.11) nor ALAN treatments (F[1,28] = 0.28; p = 0.60), and no Time x ALAN interaction (F[2,28] = 0.025; p = 0.98).

304

## 3.3- Photosynthetic efficiency

306
In the control channels, DDBAC concentrations were always below quantification limit (Table 1), confirming that no cross-contamination occurred. Contaminant concentrations averaged 26.6 ± 2.5 mg.L⁻¹ (AP) and 18.5 ± 2.0 mg.L⁻¹ (CP) in the DDBAC treatments. Differences in DDBAC concentrations were observed between the DDBAC treatment channels exposed to an alternated photoperiod and those exposed to a continuous photoperiod (F[1,10] = 232.72; p < 0.001, also see Figure 1), suggesting that degradation occurred under continuous photoperiod. Nutrient concentrations were also modified by DDBAC exposure (Table 1). Indeed, DDBAC addition resulted in lower concentrations of nitrate, sulfate, Cl⁻, Na⁺, Ca²⁺ and Mg⁺ than in the control (Table 1), while ALAN showed no effect on nutrient concentrations.

293

## 3.2- Bioaccumulation of DDBAC in biofilms

<figure><img src="image_4.png" /><figcaption>Figure 2. Evolution of DDBAC bioaccumulated in biofilms (expressed in log(BCF)) between the different ALAN treatments (n=3). AP = alternated photoperiod CP = continuous photoperiod</figcaption></figure>

297

Figure 2 shows the log(BCF) for the hydrophilic fraction of biofilm samples at d1, d2 and d10 for AP and CP treatments. DDBAC was recovered from biofilms for every treatment and every sampling time, showing that this compound bioaccumulates well in autotrophic biofilms, with a mean log(BCF) of 3.22 ± 0.25. The amount of DDBAC bioaccumulated in the biofilms showed no significant changes between sampling times (F[2,28]= 2.38; p = 0.11) nor ALAN treatments (F[1,28] = 0.28; p = 0.60), and no Time x ALAN interaction (F[2,28] = 0.025; p = 0.98).

304

## 3.3- Photosynthetic efficiency

306
<figure><img src="image_4.png" /><figcaption>Figure 3. Evolution of photosynthesis yield in biofilms along the ten days of the experiment (d0: n=5, d2 and d10: n=3). Control = non-exposed biofilm; DDBAC = contaminated biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod</figcaption></figure>

Figure 3 shows that photosynthetic activity was stable over the time of the experiment, with a mean value of 0.42 ± 0.01 in the control AP and CP channels, regardless of the ALAN treatment (F[1,42] = 2.65; p > 0.05). Biofilms exposed to DDBAC showed an almost complete inhibition of photosynthetic yield starting from d2 and lasting until the end of the experiment (F[1,42] = 1279; p < 0.001). In the same way as for control channels, photosynthetic activity showed no difference between AP and CP for contaminated channels (F[1,42] = 0.18; p = 0.67).

## 3.4- Photosynthetic pigments

<figure><img src="image_5.png" /><figcaption>Figure 4. Changes in photosynthetic pigments content in biofilms along the ten days of the experiment (n = 1). Control = non-exposed biofilm; DDBAC = contaminated biofilm; AP = alternated photoperiod; CP = continuous photoperiod.</figcaption></figure>

Chlorophyll *a* concentrations decreased over time in the DDBAC contaminated samples, compared to the control samples (Figure 4). At d2, no chlorophyll *a* was detected in the DDBAC-AP samples, and only phaeopigments were measured. At the end of the experiment, pigment concentrations in DDBAC samples were very low compared to the controls. Unfortunately, the lack of replication does not allow for any conclusion on the effect of the tested treatments on chlorophyll *a* or phaeopigments.
<figure><img src="image_5.png" /><figcaption>Figure 3. Evolution of photosynthesis yield in biofilms along the ten days of the experiment (d0: n=5, d2 and d10: n=3). Control = non-exposed biofilm; DDBAC = contaminated biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod</figcaption></figure>

Figure 3 shows that photosynthetic activity was stable over the time of the experiment, with a mean value of 0.42 ± 0.01 in the control AP and CP channels, regardless of the ALAN treatment (F<sub>[1,42]</sub> = 2.65; p > 0.05). Biofilms exposed to DDBAC showed an almost complete inhibition of photosynthetic yield starting from d2 and lasting until the end of the experiment (F<sub>[1,42]</sub> = 1279; p < 0.001). In the same way as for control channels, photosynthetic activity showed no difference between AP and CP for contaminated channels (F<sub>[1,42]</sub> = 0.18; p = 0.67).

### 3.4- Photosynthetic pigments

<figure><img src="image_6.png" /><figcaption>Figure 4. Changes in photosynthetic pigments content in biofilms along the ten days of the experiment (n = 1). Control = non-exposed biofilm; DDBAC = contaminated biofilm; AP = alternated photoperiod; CP = continuous photoperiod.</figcaption></figure>

Chlorophyll *a* concentrations decreased over time in the DDBAC contaminated samples, compared to the control samples (Figure 4). At d2, no chlorophyll *a* was detected in the DDBAC-AP samples, and only phaeopigments were measured. At the end of the experiment, pigment concentrations in DDBAC samples were very low compared to the controls. Unfortunately, the lack of replication does not allow for any conclusion on the effect of the tested treatments on chlorophyll *a* or phaeopigments.
## 3.5- Community composition

<table><caption>Table 2. Evolution of the taxonomic composition of biofilms during the experiment. Densities are expressed as individuals×10³ / cm², i.e. number of cells for microalgae, and number of organisms for microfauna. Diatom mortality is expressed in %. Results are expressed as mean and standard error for each date and treatment (d0: n=5, d2 and d10: n=3). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod</caption><thead><tr><th>Date</th><th>Treatment</th><th>Diatom density</th><th>Diatom mortality</th><th>Green algae density</th><th>Cyanobacteria density</th><th>Microfauna density</th></tr></thead><tbody><tr><td>d0</td><td>Control</td><td>81.3 ± 33.7</td><td>7 ± 1</td><td>4.1 ± 2.3</td><td>3.8 ± 2.0</td><td>0.3 ± 0.0</td></tr><tr><td>d2</td><td>Control-AP</td><td>141.3 ± 9.1</td><td>3 ± 0</td><td>26.7 ± 4.4</td><td>1.6 ± 2.0</td><td>0.6 ± 0.3</td></tr><tr><td>d2</td><td>DDBAC-AP</td><td>113.2 ± 11.5</td><td>3 ± 1</td><td>11.8 ± 2.8</td><td>0</td><td>0.2 ± 0.2</td></tr><tr><td>d2</td><td>Control-CP</td><td>132.1 ± 68.8</td><td>3 ± 1</td><td>51.7 ± 46.0</td><td>0</td><td>0.3 ± 0.3</td></tr><tr><td>d2</td><td>DDBAC-CP</td><td>145.0 ± 26.1</td><td>2 ± 1</td><td>6.2 ± 1.7</td><td>0</td><td>0</td></tr><tr><td>d10</td><td>Control-AP</td><td>98.2 ± 11.7</td><td>5 ± 2</td><td>225.6 ± 133.5</td><td>11.2 ± 7.8</td><td>0.9 ± 0.5</td></tr><tr><td>d10</td><td>DDBAC-AP</td><td>128.5 ± 26.0</td><td>2 ± 0</td><td>18.9 ± 15.0</td><td>4.8 ± 3.0</td><td>0</td></tr><tr><td>d10</td><td>Control-CP</td><td>91.3 ± 14.2</td><td>6 ± 1</td><td>354.2 ± 83.1</td><td>0</td><td>1.0 ± 0.2</td></tr><tr><td>d10</td><td>DDBAC-CP</td><td>101.0 ± 27.0</td><td>1 ± 0</td><td>30.7 ± 8.2</td><td>0</td><td>0</td></tr><tr><td colspan="2">Significant factor(s) and interaction(s)</td><td>None</td><td>DDBAC</td><td>DDBAC × Time</td><td>None</td><td>DDBAC × Time</td></tr><tr><td colspan="2">p</td><td>&gt;0.05</td><td>&lt;0.05</td><td>&lt;0.001</td><td>&gt;0.05</td><td>&lt;0.01</td></tr></tbody></table>

Diatoms (91%) dominated the biofilm community at d0, whereas low proportions of green algae (5%), cyanobacteria (4%) and microfauna (1%) were observed (Table 2). In the control channels, green algae developed markedly and became the main algal group in terms of density by d10, with no significant effect of the ALAN treatment (e.g. 67% in AP channels and 79% in CP). The density of green algae slightly increased from d0 to d10 in the DDBAC contaminated channels, but not to the same extent as in the control channels. Indeed, a significant Time × DDBAC interaction was highlighted for green algae densities (p < 0.001;
## 3.5- Community composition

<table><caption>Table 2. Evolution of the taxonomic composition of biofilms during the experiment. Densities are expressed as individuals×10³ / cm², i.e. number of cells for microalgae, and number of organisms for microfauna. Diatom mortality is expressed in %. Results are expressed as mean and standard error for each date and treatment (d0: n=5, d2 and d10: n=3). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod</caption><thead><tr><th>Date</th><th>Treatment</th><th>Diatom density</th><th>Diatom mortality</th><th>Green algae density</th><th>Cyanobacteria density</th><th>Microfauna density</th></tr></thead><tbody><tr><td>d0</td><td>Control</td><td>81.3 ± 33.7</td><td>7 ± 1</td><td>4.1 ± 2.3</td><td>3.8 ± 2.0</td><td>0.3 ± 0.0</td></tr><tr><td>d2</td><td>Control-AP</td><td>141.3 ± 9.1</td><td>3 ± 0</td><td>26.7 ± 4.4</td><td>1.6 ± 2.0</td><td>0.6 ± 0.3</td></tr><tr><td>d2</td><td>DDBAC-AP</td><td>113.2 ± 11.5</td><td>3 ± 1</td><td>11.8 ± 2.8</td><td>0</td><td>0.2 ± 0.2</td></tr><tr><td>d2</td><td>Control-CP</td><td>132.1 ± 68.8</td><td>3 ± 1</td><td>51.7 ± 46.0</td><td>0</td><td>0.3 ± 0.3</td></tr><tr><td>d2</td><td>DDBAC-CP</td><td>145.0 ± 26.1</td><td>2 ± 1</td><td>6.2 ± 1.7</td><td>0</td><td>0</td></tr><tr><td>d10</td><td>Control-AP</td><td>98.2 ± 11.7</td><td>5 ± 2</td><td>225.6 ± 133.5</td><td>11.2 ± 7.8</td><td>0.9 ± 0.5</td></tr><tr><td>d10</td><td>DDBAC-AP</td><td>128.5 ± 26.0</td><td>2 ± 0</td><td>18.9 ± 15.0</td><td>4.8 ± 3.0</td><td>0</td></tr><tr><td>d10</td><td>Control-CP</td><td>91.3 ± 14.2</td><td>6 ± 1</td><td>354.2 ± 83.1</td><td>0</td><td>1.0 ± 0.2</td></tr><tr><td>d10</td><td>DDBAC-CP</td><td>101.0 ± 27.0</td><td>1 ± 0</td><td>30.7 ± 8.2</td><td>0</td><td>0</td></tr><tr><td colspan="2">Significant factor(s) and interaction(s)</td><td>None</td><td>DDBAC</td><td>DDBAC × Time</td><td>None</td><td>DDBAC × Time</td></tr><tr><td colspan="2">p</td><td>&gt;0.05</td><td>&lt;0.05</td><td>&lt;0.001</td><td>&gt;0.05</td><td>&lt;0.01</td></tr></tbody></table>

Diatoms (91%) dominated the biofilm community at d0, whereas low proportions of green algae (5%), cyanobacteria (4%) and microfauna (1%) were observed (Table 2). In the control channels, green algae developed markedly and became the main algal group in terms of density by d10, with no significant effect of the ALAN treatment (e.g. 67% in AP channels and 79% in CP). The density of green algae slightly increased from d0 to d10 in the DDBAC contaminated channels, but not to the same extent as in the control channels. Indeed, a significant Time × DDBAC interaction was highlighted for green algae densities ($p < 0.001$;
344

Table 2). Microfauna increased in control channels, but had completely disappeared from biofilms exposed to DDBAC by d10. For this component, a significant Time × DDBAC interaction was also noted (p < 0.01; Table 2). ALAN treatments did not have a significant impact on the biofilm taxonomic composition.

345

346

347

348

Diatom mortality values (based on the ratio between frustules without cell content and total diatom frustules) significantly decreased under DDBAC exposure (Table 2). In addition, microscopic observations highlighted differences in the aspect (shape, colour) of chloroplasts in diatoms and chlorophytes exposed to DDBAC, becoming highly granular and darker (Figure B.2). These were, however, not considered as 'dead' cells in estimating diatom mortality.

349

350

351

352

353

354

## 3.6- Lipid content

355

<figure><img src="image_6.png" /></figure>

356

357

<caption>Figure 5. Effects of DDBAC and continuous photoperiod on the proportion of total lipids and on lipid classes (n=4). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = alternated photoperiod; CP = continuous photoperiod; PE = phosphatidylethanolamine; PC = phosphatidylcholine; PG = phosphatidylglycerol; DGDG = digalactosyldiacylglycerol; MGDG = monogalactosyldiacylglycerol; SQDG = sulfoquinovosyldiacylglycerol; TAG = triacylglycerides.</caption>

358

359

360

361

362

363

On d0, total lipid concentration reached 42.6 nmol.mg⁻¹ of biofilm dry weight. Almost half of the lipids were phospholipids, and the other half were glycolipids, while neutral lipids (TAG) represented a minor portion (Table 3). By d10, total lipid concentration had decreased in all treatments down to an average of 23.6 ± 7.53 nmol.mg⁻¹ of biofilm dry weight (Table 3). No significant differences were found between AP and CP, while DDBAC significantly affected

364

365

366

367
<figure><img src="image_7.png" /></figure>

Table 2). Microfauna increased in control channels, but had completely disappeared from biofilms exposed to DDBAC by d10. For this component, a significant Time × DDBAC interaction was also noted (p < 0.01; Table 2). ALAN treatments did not have a significant impact on the biofilm taxonomic composition.

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /></figure>

<figure><img src="image_17.png" /></figure>

<figure><img src="image_18.png" /></figure>

Diatom mortality values (based on the ratio between frustules without cell content and total diatom frustules) significantly decreased under DDBAC exposure (Table 2). In addition, microscopic observations highlighted differences in the aspect (shape, colour) of chloroplasts in diatoms and chlorophytes exposed to DDBAC, becoming highly granular and darker (Figure B.2). These were, however, not considered as 'dead' cells in estimating diatom mortality.

## 3.6- Lipid content

<figure><img src="image_19.png" /></figure>

356

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /></figure>

<figure><img src="image_22.png" /></figure>

<figure><img src="image_23.png" /></figure>

<figure><img src="image_24.png" /><figcaption>Figure 5. Effects of DDBAC and continuous photoperiod on the proportion of total lipids and on lipid classes (n=4). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = alternated photoperiod; CP = continuous photoperiod; PE = phosphatidylethanolamine; PC = phosphatidylcholine; PG = phosphatidylglycerol; DGDG = digalactosyldiacylglycerol; MGDG = monogalactosyldiacylglycerol; SQDG = sulfoquinovosyldiacylglycerol; TAG = triacylglycerides.</figcaption></figure>

362

<figure><img src="image_25.png" /></figure>

<figure><img src="image_26.png" /></figure>

<figure><img src="image_27.png" /></figure>

<figure><img src="image_28.png" /></figure>

<figure><img src="image_29.png" /></figure>

On d0, total lipid concentration reached 42.6 nmol.mg⁻¹ of biofilm dry weight. Almost half of the lipids were phospholipids, and the other half were glycolipids, while neutral lipids (TAG) represented a minor portion (Table 3). By d10, total lipid concentration had decreased in all treatments down to an average of 23.6 ± 7.53 nmol.mg⁻¹ of biofilm dry weight (Table 3). No significant differences were found between AP and CP, while DDBAC significantly affected
368 glycolipids over time (significant Time × DDBAC interaction; p < 0.01). Figure 5 shows lipid
369 contents under DDBAC and ALAN exposure on d0 and d10. At the beginning of the experiment,
370 phosphatidylglycerol (PG) was the predominant lipid in the biofilms, while phospholipids and
371 glycolipids in the biofilms were evenly distributed. Total lipid amounts decreased over time in
372 the controls, but the proportions of phospholipids and glycolipids remained similar to those
373 at d0, all the main classes of lipids being present. Total phospholipids remained stable over
374 time, whatever the treatment (p > 0.05), but phosphatidylethanolamine (PE) and PG were
375 higher in biofilms exposed to DDBAC compared to controls (Figure 5, Table 3). In the samples
376 exposed to DDBAC, glycolipids such as monogalactosyldiacylglycerol (MGDG) and
377 digalactosyldiacylglycerol (DGDG) decreased significantly, or even became barely detectable,
378 with significant Time × DDBAC interactions (respectively, p < 0.01 and 0.001).
379 Diacylglyceryltrimethylhomo-Ser (DGTS) was never detected and, therefore, is not mentioned
380 further.
368 glycolipids over time (significant Time × DDBAC interaction; p < 0.01). Figure 5 shows lipid
369 contents under DDBAC and ALAN exposure on d0 and d10. At the beginning of the experiment,
370 phosphatidylglycerol (PG) was the predominant lipid in the biofilms, while phospholipids and
371 glycolipids in the biofilms were evenly distributed. Total lipid amounts decreased over time in
372 the controls, but the proportions of phospholipids and glycolipids remained similar to those
373 at d0, all the main classes of lipids being present. Total phospholipids remained stable over
374 time, whatever the treatment (p > 0.05), but phosphatidylethanolamine (PE) and PG were
375 higher in biofilms exposed to DDBAC compared to controls (Figure 5, Table 3). In the samples
376 exposed to DDBAC, glycolipids such as monogalactosyldiacylglycerol (MGDG) and
377 digalactosyldiacylglycerol (DGDG) decreased significantly, or even became barely detectable,
378 with significant Time × DDBAC interactions (respectively, p < 0.01 and 0.001).
379 Diacylglyceryltrimethylhomo-Ser (DGTS) was never detected and, therefore, is not mentioned
380 further.
<table><caption>Table 3: Evolution of lipid content of biofilms (in nmol.g-1 freeze-dried biofilm) between the beginning and the end of the experiment. Results are expressed as mean and standard error for each date and treatment (n=4). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod; PE = phosphatidylethanolamine; PC = phosphatidylcholine; PG = phosphatidylglycerol; DGDG = digalactosyldiacylglycerol; MGDG = monogalactosyldiacylglycerol; SQDG = sulfoquinovosyldiacylglycerol; TAG = triacylglycerides. Two-ways ANOVAs on repeated measures were applied on data.</caption><thead><tr><th>Date</th><th>Treatment</th><th>Neutral lipids (TAG)</th><th>PG</th><th>PE</th><th>PC</th><th>Σ Phospholipids</th><th>DGDG</th><th>MGDG</th><th>SQDG</th><th>Σ Glycolipids</th><th>Total lipids</th></tr></thead><tbody><tr><td>d0</td><td>Control</td><td>4.2 ± 2.5</td><td>13.7 ± 3.1</td><td>2.8 ± 0.5</td><td>4.6 ± 0.2</td><td>21.0 ± 3.3</td><td>5.6 ± 1.2</td><td>6.4 ± 1.6</td><td>5.3 ± 1.1</td><td>17.3 ± 2.4</td><td>42.6 ± 3.2</td></tr><tr><td>d10</td><td>Control-AP</td><td>3.9 ± 2.0</td><td>6.4 ± 1.0</td><td>1.3 ± 0.5</td><td>3.5 ± 1.5</td><td>11.2 ± 2.4</td><td>4.8 ± 0.8</td><td>2.9 ± 0.5</td><td>4.2 ± 0.7</td><td>12.0 ± 0.4</td><td>27.1 ± 4.1</td></tr><tr><td>d10</td><td>DDBAC-AP</td><td>0.2 ± 0.3</td><td>14.3 ± 2.6</td><td>3.0 ± 0.4</td><td>0.9 ± 0.4</td><td>18.2 ± 2.6</td><td>0</td><td>0</td><td>0.7 ± 0.2</td><td>0.7 ± 0.2</td><td>19.1 ± 2.6</td></tr><tr><td>d10</td><td>Control-CP</td><td>6.6 ± 2.9</td><td>6.9 ± 6.7</td><td>0.7 ± 0.1</td><td>2.6 ± 1.1</td><td>10.1 ± 7.8</td><td>5.3 ± 0.2</td><td>3.3 ± 1.1</td><td>5.8 ± 1.2</td><td>14.4 ± 2.2</td><td>31.1 ± 8.1</td></tr><tr><td>d10</td><td>DDBAC-CP</td><td>0.1 ± 0.1</td><td>13.3 ± 3.5</td><td>2.3 ± 1.0</td><td>1.0 ± 1.0</td><td>16.6 ± 5.2</td><td>0</td><td>0</td><td>0.4 ± 0.2</td><td>0.4 ± 0.2</td><td>17.1 ± 5.3</td></tr><tr><td></td><td>Significant factor(s)</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>None</td><td>DDBAC 12× Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td></tr><tr><td></td><td>p</td><td>0.01</td><td>&lt;0.05</td><td>&lt;0.01</td><td>&lt;0.01</td><td>&gt;0.05</td><td>&lt;0.001</td><td>&lt;0.01</td><td>&lt;0.001</td><td>&lt;0.001</td><td>&lt;0.001</td></tr></tbody></table>

382
<table><caption>Table 3: Evolution of lipid content of biofilms (in nmol.g-1 freeze-dried biofilm) between the beginning and the end of the experiment. Results are expressed as mean and standard error for each date and treatment (n=4). DDBAC = contaminated biofilm; Control = non-exposed biofilm; AP = Alternated Photoperiod; CP = Continuous Photoperiod; PE = phosphatidylethanolamine; PC = phosphatidylcholine; PG = phosphatidylglycerol; DGDG = digalactosyldiacylglycerol; MGDG = monogalactosyldiacylglycerol; SQDG = sulfoquinovosyldiacylglycerol; TAG = triacylglycerides. Two-ways ANOVAs on repeated measures were applied on data.</caption><caption>381</caption><thead><tr><th>Date</th><th>Treatment</th><th>Neutral lipids (TAG)</th><th>PG</th><th>PE</th><th>PC</th><th>Σ Phospholipids</th><th>DGDG</th><th>MGDG</th><th>SQDG</th><th>Σ Glycolipids</th><th>Total lipids</th></tr></thead><tbody><tr><td>d0</td><td>Control</td><td>4.2 ± 2.5</td><td>13.7 ± 3.1</td><td>2.8 ± 0.5</td><td>4.6 ± 0.2</td><td>21.0 ± 3.3</td><td>5.6 ± 1.2</td><td>6.4 ± 1.6</td><td>5.3 ± 1.1</td><td>17.3 ± 2.4</td><td>42.6 ± 3.2</td></tr><tr><td>d10</td><td>Control-AP</td><td>3.9 ± 2.0</td><td>6.4 ± 1.0</td><td>1.3 ± 0.5</td><td>3.5 ± 1.5</td><td>11.2 ± 2.4</td><td>4.8 ± 0.8</td><td>2.9 ± 0.5</td><td>4.2 ± 0.7</td><td>12.0 ± 0.4</td><td>27.1 ± 4.1</td></tr><tr><td>d10</td><td>DDBAC-AP</td><td>0.2 ± 0.3</td><td>14.3 ± 2.6</td><td>3.0 ± 0.4</td><td>0.9 ± 0.4</td><td>18.2 ± 2.6</td><td>0</td><td>0</td><td>0.7 ± 0.2</td><td>0.7 ± 0.2</td><td>19.1 ± 2.6</td></tr><tr><td>d10</td><td>Control-CP</td><td>6.6 ± 2.9</td><td>6.9 ± 6.7</td><td>0.7 ± 0.1</td><td>2.6 ± 1.1</td><td>10.1 ± 7.8</td><td>5.3 ± 0.2</td><td>3.3 ± 1.1</td><td>5.8 ± 1.2</td><td>14.4 ± 2.2</td><td>31.1 ± 8.1</td></tr><tr><td>d10</td><td>DDBAC-CP</td><td>0.1 ± 0.1</td><td>13.3 ± 3.5</td><td>2.3 ± 1.0</td><td>1.0 ± 1.0</td><td>16.6 ± 5.2</td><td>0</td><td>0</td><td>0.4 ± 0.2</td><td>0.4 ± 0.2</td><td>17.1 ± 5.3</td></tr><tr><td></td><td>Significant factor(s)</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>None</td><td>DDBAC 12× Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td><td>DDBAC × Time</td></tr><tr><td></td><td>p</td><td>0.01</td><td>&lt;0.05</td><td>&lt;0.01</td><td>&lt;0.01</td><td>&gt;0.05</td><td>&lt;0.001</td><td>&lt;0.01</td><td>&lt;0.001</td><td>&lt;0.001</td><td>&lt;0.001</td></tr></tbody></table>

382
# 4. Discussion

## 4.1- Experimental conditions and DDBAC exposure

In this experiment, natural biofilms were transferred to controlled laboratory conditions under ALAN and DDBAC treatments. Changes in environmental conditions were likely to affect biofilm characteristics; control conditions were however favourable to the development of autotrophic biofilms, as highlighted by continuous growth from d0 to d10 and by stable physiological state (assessed through photosynthetic efficiency). During the 10 days, most physicochemical parameters remained stable, and no nutrient depletion was observed at the end of the experiment in any treatment. However, DDBAC seemed to have an indirect effect on the concentrations of certain nutrients over the 10-day exposure period of our experiment. For example, under DDBAC exposure, concentrations of $NO_3^-$ were lower than in the control (Table 1).

Contrary to the hypothesis of Pozo-Antonio and Sanmartin (2018), who suggested that artificial light could lead to an increase in internalization of the contaminant, we found no significant difference in DDBAC concentrations in biofilm between ALAN treatments (Figure 2). Thus, our results do not support the hypothesis of a higher uptake of the substance from the water through increased bioaccumulation under ALAN exposure. Lower concentrations of DDBAC under CP compared to AP (Figure 1) might be linked to the differential development of microorganisms exposed to ALAN. Some bacterial strains such as *Aeromonas hydrophila* can biodegrade DDBAC under N-depleted conditions and use the degradation product as a source of nitrogen (Patrauchan and Oriel, 2003). The above-mentioned decrease in certain nutrient concentrations in the water under DDBAC treatment may have provided favourable conditions for such strains able to grow consuming DDBAC as a nutrient source. Moreover, the low light intensity used in the experiment may have enhanced the growth of such heterotrophs in the absence of a strong competition with autotroph organisms from the biofilm.

The potential for DDBAC exposure on biofilm consumers (related to bioaccumulation in the biofilm) was similar between ALAN treatments, independently of differences in DDBAC concentrations in the water. DDBAC accumulation rapidly stabilized to average $log(BCF)$ of 3.22 (Figure 2), falling in the range measured for organochlorine substances in biofilms, and greater than previous observations for other pesticides or pharmaceuticals (Bonnineau et al. 2021). Our results demonstrate the retention capacity of biofilms for this compound, suggesting a risk of transfer to primary consumers.

## 4.2- Effects of DDBAC on the biofilm

The first effect of DDBAC observed was the marked and rapid decrease in photosynthetic efficiency (more than 90% compared with the initial yield; Figure 3). This sudden inhibition of photosynthesis was not expected because the exposure concentration was based on the $EC_5$ for photosynthesis inhibition after four hours of exposure determined in a preliminary experiment ($EC_5$ = 30 mg.L$^{-1}$; Figure A). However, this result is in line with the high ecotoxicity of DDBAC to freshwater and estuarine/marine algae reported by Arnold et al. (2023), with $EC_{50}$ values as low as 14 µg.L$^{-1}$. Based on our microscopic observations, diatom mortality was low in all samples (<10%). It should be noted that, following Morin et al. (2010), only diatoms that
# 4. Discussion

## 4.1- Experimental conditions and DDBAC exposure

In this experiment, natural biofilms were transferred to controlled laboratory conditions under ALAN and DDBAC treatments. Changes in environmental conditions were likely to affect biofilm characteristics; control conditions were however favourable to the development of autotrophic biofilms, as highlighted by continuous growth from d0 to d10 and by stable physiological state (assessed through photosynthetic efficiency). During the 10 days, most physicochemical parameters remained stable, and no nutrient depletion was observed at the end of the experiment in any treatment. However, DDBAC seemed to have an indirect effect on the concentrations of certain nutrients over the 10-day exposure period of our experiment. For example, under DDBAC exposure, concentrations of $NO_3^-$ were lower than in the control (Table 1).

Contrary to the hypothesis of Pozo-Antonio and Sanmartin (2018), who suggested that artificial light could lead to an increase in internalization of the contaminant, we found no significant difference in DDBAC concentrations in biofilm between ALAN treatments (Figure 2). Thus, our results do not support the hypothesis of a higher uptake of the substance from the water through increased bioaccumulation under ALAN exposure. Lower concentrations of DDBAC under CP compared to AP (Figure 1) might be linked to the differential development of microorganisms exposed to ALAN. Some bacterial strains such as *Aeromonas hydrophila* can biodegrade DDBAC under N-depleted conditions and use the degradation product as a source of nitrogen (Patrauchan and Oriel, 2003). The above-mentioned decrease in certain nutrient concentrations in the water under DDBAC treatment may have provided favourable conditions for such strains able to grow consuming DDBAC as a nutrient source. Moreover, the low light intensity used in the experiment may have enhanced the growth of such heterotrophs in the absence of a strong competition with autotroph organisms from the biofilm.

The potential for DDBAC exposure on biofilm consumers (related to bioaccumulation in the biofilm) was similar between ALAN treatments, independently of differences in DDBAC concentrations in the water. DDBAC accumulation rapidly stabilized to average $log(BCF)$ of 3.22 (Figure 2), falling in the range measured for organochlorine substances in biofilms, and greater than previous observations for other pesticides or pharmaceuticals (Bonnineau et al. 2021). Our results demonstrate the retention capacity of biofilms for this compound, suggesting a risk of transfer to primary consumers.

## 4.2- Effects of DDBAC on the biofilm

The first effect of DDBAC observed was the marked and rapid decrease in photosynthetic efficiency (more than 90% compared with the initial yield; Figure 3). This sudden inhibition of photosynthesis was not expected because the exposure concentration was based on the $EC_5$ for photosynthesis inhibition after four hours of exposure determined in a preliminary experiment ($EC_5$ = 30 mg.L$^{-1}$; Figure A). However, this result is in line with the high ecotoxicity of DDBAC to freshwater and estuarine/marine algae reported by Arnold et al. (2023), with $EC_{50}$ values as low as 14 µg.L$^{-1}$. Based on our microscopic observations, diatom mortality was low in all samples (<10%). It should be noted that, following Morin et al. (2010), only diatoms that
no longer had chlorophyll in their cell content were considered as dead cells, while cells with chloroplasts were counted as live diatoms. However, many of the cells where chloroplastic content was observed seemed to have suffered critical alteration of their photosynthetic equipment (Figure B.2). Their integrity was potentially severely affected, which could explain the loss of photosynthetic yield in biofilms exposed to DDBAC. This deterioration of cell integrity may be due to the fact that DDBAC is able to quickly penetrate the cell (Severina et al. 2001). Our results highlight, in agreement with Wood et al. (2014) and Pandey et al. (2017), the ecological relevance of microscopic observations of fresh material to better assess the ecotoxicity of biocides.

Marked differences in the autotrophic community composition were highlighted by microscopy counts between d2 and d10 under DDBAC exposure. Indeed, densities of green algae were 10 times lower than under control conditions (Table 2). Despite the lack of replication for pigment analyses, the degradation of chlorophyll pigments in DDBAC treatments (Figure 4) is in line with the impairment of the algal component of the biofilms as highlighted through photosynthetic inhibition and chloroplast alteration. The striking decrease in lipid content in biofilms exposed to DDBAC (Table 3), particularly glycolipids found mainly in thylakoid membranes within photosynthetic cells (Zulu et al., 2018), supports the idea that the contaminant is internalized within phototrophic cells, with deleterious impacts on the autotrophic component. Diatom-rich biofilms are generally source of essential polyunsaturated fatty acids such as eicosapentaenoic acid (Zulu et al., 2018), ensuring a high nutritional quality of this food resource for primary consumers (Brett and Müller-Navarra, 1997). Demailly et al. (2019) also demonstrated that diatom fatty acid composition may be altered by organic substances such as pesticides. More recently, Mazzella et al. (2023b) highlighted a decrease in eicosapentaenoic acid content, and more generally in polyunsaturated fatty acids, in DDBAC-exposed biofilms collected from the present study. DDBAC-driven changes in biofilm community structure and impact on glycolipids could therefore lead to a decrease in the nutritional quality of biofilms (decrease in essential polyunsaturated fatty acids, such as certain omega-3 and omega-6) and subsequently affect the trophic chain (Brett and Müller-Navarra, 1997), worsened by DDBAC accumulation.

In parallel to the decrease in glycolipids noted under DDBAC contamination, we observed an increase in phospholipids. It is difficult to clearly explain this increase because the phospholipid groups analysed (i.e. PE, PC and PG) are not specific to any taxonomic group and can be found in both the plasma membranes of microalgae (Zulu et al., 2018) and in prokaryotic cells (Li-Beisson et al., 2013). One hypothesis to explain this increase could be the development of DDBAC-resistant bacteria, as mentioned above. Indeed, the mechanism of resistance to benzalkonium chloride identified in *Pseudomonas aeruginosa* consists of an increase in the percentage of phospholipids (Sakagami et al. 1989). Considering the impact of DDBAC on biofilm bacteria would be required to determine if the increase in phospholipids observed in our study reflects the development of specific bacteria under DDBAC exposure.

Sensitivity to DDBAC differed among biofilm taxa, whose proportions changed over time, and with DDBAC concentration. This result suggests that the biofilm community acclimated to the chemical stress and that only resistant/tolerant species survived DDBAC exposure according
no longer had chlorophyll in their cell content were considered as dead cells, while cells with chloroplasts were counted as live diatoms. However, many of the cells where chloroplastic content was observed seemed to have suffered critical alteration of their photosynthetic equipment (Figure B.2). Their integrity was potentially severely affected, which could explain the loss of photosynthetic yield in biofilms exposed to DDBAC. This deterioration of cell integrity may be due to the fact that DDBAC is able to quickly penetrate the cell (Severina et al. 2001). Our results highlight, in agreement with Wood et al. (2014) and Pandey et al. (2017), the ecological relevance of microscopic observations of fresh material to better assess the ecotoxicity of biocides.

Marked differences in the autotrophic community composition were highlighted by microscopy counts between d2 and d10 under DDBAC exposure. Indeed, densities of green algae were 10 times lower than under control conditions (Table 2). Despite the lack of replication for pigment analyses, the degradation of chlorophyll pigments in DDBAC treatments (Figure 4) is in line with the impairment of the algal component of the biofilms as highlighted through photosynthetic inhibition and chloroplast alteration. The striking decrease in lipid content in biofilms exposed to DDBAC (Table 3), particularly glycolipids found mainly in thylakoid membranes within photosynthetic cells (Zulu et al., 2018), supports the idea that the contaminant is internalized within phototrophic cells, with deleterious impacts on the autotrophic component. Diatom-rich biofilms are generally source of essential polyunsaturated fatty acids such as eicosapentaenoic acid (Zulu et al., 2018), ensuring a high nutritional quality of this food resource for primary consumers (Brett and Müller-Navarra, 1997). Demailly et al. (2019) also demonstrated that diatom fatty acid composition may be altered by organic substances such as pesticides. More recently, Mazzella et al. (2023b) highlighted a decrease in eicosapentaenoic acid content, and more generally in polyunsaturated fatty acids, in DDBAC-exposed biofilms collected from the present study. DDBAC-driven changes in biofilm community structure and impact on glycolipids could therefore lead to a decrease in the nutritional quality of biofilms (decrease in essential polyunsaturated fatty acids, such as certain omega-3 and omega-6) and subsequently affect the trophic chain (Brett and Müller-Navarra, 1997), worsened by DDBAC accumulation.

In parallel to the decrease in glycolipids noted under DDBAC contamination, we observed an increase in phospholipids. It is difficult to clearly explain this increase because the phospholipid groups analysed (i.e. PE, PC and PG) are not specific to any taxonomic group and can be found in both the plasma membranes of microalgae (Zulu et al., 2018) and in prokaryotic cells (Li-Beisson et al., 2013). One hypothesis to explain this increase could be the development of DDBAC-resistant bacteria, as mentioned above. Indeed, the mechanism of resistance to benzalkonium chloride identified in *Pseudomonas aeruginosa* consists of an increase in the percentage of phospholipids (Sakagami et al. 1989). Considering the impact of DDBAC on biofilm bacteria would be required to determine if the increase in phospholipids observed in our study reflects the development of specific bacteria under DDBAC exposure.

Sensitivity to DDBAC differed among biofilm taxa, whose proportions changed over time, and with DDBAC concentration. This result suggests that the biofilm community acclimated to the chemical stress and that only resistant/tolerant species survived DDBAC exposure according
468 to the Pollution-Induced Community Tolerance concept (Blanck et al., 1988). Diatoms
469 appeared to be minimally affected based on growth and mortality, even though the alteration
470 of their photosynthetic structure and cell content suggests a marked impact (Figure B.2).
471 Green algae, which experienced rapid growth in non-contaminated channels, appeared to be
472 negatively affected by DDBAC at d10. DDBAC also had a strong effect on heterotrophic
473 microfauna naturally present in the biofilms (e.g. rotifers), suggesting that they are highly
474 sensitive to this contaminant. Our dataset does not allow to disentangle whether such impacts
475 derive from direct (through water contamination) or indirect (trophic transfer from
476 contaminated biofilms) effects. However, such results confirm the high sensitivity of aquatic
477 microfauna and invertebrates to DDBAC, as already demonstrated for acute exposure with
478 *Daphnia magna* (Kreuzinger et al., 2007; Leal et al., 1994; Chen et al., 2014; Lavorgna et al.,
479 2016).
480

480

481 **4.3- Effects of ALAN on the biofilm**

482 Our experimental conditions were based on constant light intensities over time, which does
483 not represent realistic conditions. In the field, daylight variations in irradiation are important
484 drivers of biofilm growth and photosynthetic performance (Laviale et al. 2009). Under our
485 exposure conditions, ALAN did not significantly modify the structure or physiology of the
486 autotrophic organisms in the biofilm based on the parameters that we investigated. The low
487 light intensity chosen, and the use of similar light levels during day and night, may have not
488 been enough to generate stress during continuous light exposure, at the timescale of the
489 experiment. Indeed, using biofilms originating from the same pond and exposed to higher
490 irradiance (~200 µmol.s⁻¹.m⁻²), Roux et al. (2024) highlighted than ALAN decreased the daily
491 photosynthetic yield after 14 and 30 days of exposure.
492

492 Concerning the polar lipid classes, an increase in MGDG or DGDG is often observed during low
493 light intensity exposures (Gushina and Harwood, 2009). In the present experiment, while light
494 was low, the duration of exposure differed, which could explain the absence of a significant
495 effect on these two glycolipids over time. Conversely, high light levels can lead to a decrease
496 in polar lipids, especially phospholipids, in favour of TAGs, especially in the case of filamentous
497 green algae such as *Cladophora* spp. (Gushina and Harwood, 2009). Changes in lipid content
498 with ALAN were not clearly observed here, contrarily to what was observed based on a more
499 detailed assessment of fatty acids conducted on the same samples (Mazzella et al. (2023b).
500 Amini Khoeyi et al. (2011) also showed that a prolonged photoperiod can decrease the content
501 of monounsaturated (MUFA) and polyunsaturated (PUFA) fatty acids in the microalgae
502 *Chlorella vulgaris*. Considering the constant low light conditions we used, it is likely that the
503 apical endpoints we measured did not allow to observe subtle changes occurring at the
504 individual or subindividual scales. This could be due to compensation phenomena or to the
505 fact that we averaged lipids across different organisms present within a natural biofilm, which
506 differs from the analysis of an isolated chlorophyte strain.
507

507

508 **4.4- Combined effects of DDBAC and ALAN on the biofilm**
468 to the Pollution-Induced Community Tolerance concept (Blanck et al., 1988). Diatoms
469 appeared to be minimally affected based on growth and mortality, even though the alteration
470 of their photosynthetic structure and cell content suggests a marked impact (Figure B.2).
471 Green algae, which experienced rapid growth in non-contaminated channels, appeared to be
472 negatively affected by DDBAC at d10. DDBAC also had a strong effect on heterotrophic
473 microfauna naturally present in the biofilms (e.g. rotifers), suggesting that they are highly
474 sensitive to this contaminant. Our dataset does not allow to disentangle whether such impacts
475 derive from direct (through water contamination) or indirect (trophic transfer from
476 contaminated biofilms) effects. However, such results confirm the high sensitivity of aquatic
477 microfauna and invertebrates to DDBAC, as already demonstrated for acute exposure with
478 *Daphnia magna* (Kreuzinger et al., 2007; Leal et al., 1994; Chen et al., 2014; Lavorgna et al.,
479 2016).
480

480

481 **4.3- Effects of ALAN on the biofilm**

482 Our experimental conditions were based on constant light intensities over time, which does
483 not represent realistic conditions. In the field, daylight variations in irradiation are important
484 drivers of biofilm growth and photosynthetic performance (Laviale et al. 2009). Under our
485 exposure conditions, ALAN did not significantly modify the structure or physiology of the
486 autotrophic organisms in the biofilm based on the parameters that we investigated. The low
487 light intensity chosen, and the use of similar light levels during day and night, may have not
488 been enough to generate stress during continuous light exposure, at the timescale of the
489 experiment. Indeed, using biofilms originating from the same pond and exposed to higher
490 irradiance (~200 µmol.s⁻¹.m⁻²), Roux et al. (2024) highlighted than ALAN decreased the daily
491 photosynthetic yield after 14 and 30 days of exposure.
492

492 Concerning the polar lipid classes, an increase in MGDG or DGDG is often observed during low
493 light intensity exposures (Gushina and Harwood, 2009). In the present experiment, while light
494 was low, the duration of exposure differed, which could explain the absence of a significant
495 effect on these two glycolipids over time. Conversely, high light levels can lead to a decrease
496 in polar lipids, especially phospholipids, in favour of TAGs, especially in the case of filamentous
497 green algae such as *Cladophora* spp. (Gushina and Harwood, 2009). Changes in lipid content
498 with ALAN were not clearly observed here, contrarily to what was observed based on a more
499 detailed assessment of fatty acids conducted on the same samples (Mazzella et al. (2023b).
500 Amini Khoeyi et al. (2011) also showed that a prolonged photoperiod can decrease the content
501 of monounsaturated (MUFA) and polyunsaturated (PUFA) fatty acids in the microalgae
502 *Chlorella vulgaris*. Considering the constant low light conditions we used, it is likely that the
503 apical endpoints we measured did not allow to observe subtle changes occurring at the
504 individual or subindividual scales. This could be due to compensation phenomena or to the
505 fact that we averaged lipids across different organisms present within a natural biofilm, which
506 differs from the analysis of an isolated chlorophyte strain.
507

507

508 **4.4- Combined effects of DDBAC and ALAN on the biofilm**
Contrary to our hypotheses, DDBAC and ALAN exposure did not show interaction effects on any of the biological endpoints monitored in this experiment. However, ALAN impacted the fate of DDBAC by decreasing exposure concentrations in the medium under our experimental conditions. This result can hardly be extrapolated to natural light regime conditions, calling for further investigations under more realistic light scenarios. DDBAC exposure caused significant impacts on biofilm function and structure, especially impacting autotrophs. Given the wide range of uses of DDBAC, our study highlights that its transfer to the aquatic environment may result in a significant impairment of the functioning of aquatic ecosystems. As the high concentration of DDBAC tested drove most of the changes observed in biofilm composition and physiology, we cannot exclude that DDBAC exposure masked possible interactions with ALAN. Further experiments combining lower concentrations of DDBAC with more realistic light regimes would be required to completely rule out any interaction effects on aquatic biofilms.

## 5- Conclusion

In this experiment, we demonstrated that DDBAC exposure strongly impacts river biofilms. DDBAC accumulated quickly in biofilms and damaged the photosynthetic material, in turn altering photosynthesis. There was also evidence for an impact on the structure of the biofilm as changes in its taxonomic composition were observed. Indeed, even though the mortality index did not show greater mortality in the exposed diatoms compared with the controls, microscopic observations suggest that a marked number of individuals were strongly impacted on a physiological level (degraded cell content) and were potentially not viable. This hypothesis is supported by the results obtained during the lipidomic analysis, highlighting a strong decrease in the lipid classes associated with thylakoid membranes specific to microalgae. At elevated concentrations, we highlighted striking effects of DDBAC on the aquatic biota. As DDBAC is a common contaminant with multiple domestic uses, complementary research is needed to characterize its impacts at environmentally relevant concentrations.

In contrast, ALAN did not lead to any significant change in biofilms, even though the ALAN treatment modified their exposure to DDBAC. In view of the likely increase in DDBAC concentration in urban waters (downstream of WWTPs) where ALAN is often common, it would also be necessary to investigate whether the effects identified in the present study are manifested at environmental concentrations, under more realistic light regimes, in order to reassess the risk posed by DDBAC to aquatic biodiversity.

## Acknowledgements

The authors acknowledge the financial support from the Institut National de Recherche pour l'Agriculture, l'alimentation et l'Environnement (INRAE) and the Groupe de Recherche Interuniversitaire en Limnologie (GRIL).
Contrary to our hypotheses, DDBAC and ALAN exposure did not show interaction effects on any of the biological endpoints monitored in this experiment. However, ALAN impacted the fate of DDBAC by decreasing exposure concentrations in the medium under our experimental conditions. This result can hardly be extrapolated to natural light regime conditions, calling for further investigations under more realistic light scenarios. DDBAC exposure caused significant impacts on biofilm function and structure, especially impacting autotrophs. Given the wide range of uses of DDBAC, our study highlights that its transfer to the aquatic environment may result in a significant impairment of the functioning of aquatic ecosystems. As the high concentration of DDBAC tested drove most of the changes observed in biofilm composition and physiology, we cannot exclude that DDBAC exposure masked possible interactions with ALAN. Further experiments combining lower concentrations of DDBAC with more realistic light regimes would be required to completely rule out any interaction effects on aquatic biofilms.

521

522

## 5- Conclusion

In this experiment, we demonstrated that DDBAC exposure strongly impacts river biofilms. DDBAC accumulated quickly in biofilms and damaged the photosynthetic material, in turn altering photosynthesis. There was also evidence for an impact on the structure of the biofilm as changes in its taxonomic composition were observed. Indeed, even though the mortality index did not show greater mortality in the exposed diatoms compared with the controls, microscopic observations suggest that a marked number of individuals were strongly impacted on a physiological level (degraded cell content) and were potentially not viable. This hypothesis is supported by the results obtained during the lipidomic analysis, highlighting a strong decrease in the lipid classes associated with thylakoid membranes specific to microalgae. At elevated concentrations, we highlighted striking effects of DDBAC on the aquatic biota. As DDBAC is a common contaminant with multiple domestic uses, complementary research is needed to characterize its impacts at environmentally relevant concentrations.

In contrast, ALAN did not lead to any significant change in biofilms, even though the ALAN treatment modified their exposure to DDBAC. In view of the likely increase in DDBAC concentration in urban waters (downstream of WWTPs) where ALAN is often common, it would also be necessary to investigate whether the effects identified in the present study are manifested at environmental concentrations, under more realistic light regimes, in order to reassess the risk posed by DDBAC to aquatic biodiversity.

543

544

## Acknowledgements

546

The authors acknowledge the financial support from the Institut National de Recherche pour l'Agriculture, l'alimentation et l'Environnement (INRAE) and the Groupe de Recherche Interuniversitaire en Limnologie (GRIL).

550
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600 Grubisic, M., Singer, G., Bruno, M.C., Grunsven, R.H.A. van, Manfrin, A., Monaghan, M.T., Hölker, F., 2017. Artificial light at night decreases biomass and alters community composition of benthic primary producers in a sub-alpine stream. Limnology and Oceanography 62, 2799-2810. https://doi.org/10.1002/lno.10607

604 Hora, P.I., Arnold, W.A., 2020. Photochemical fate of quaternary ammonium compounds in river water. Environmental Science: Processes & Impacts 22(6),1368-1381. https://pubs.rsc.org/en/content/articlelanding/2020/em/d0em00086h

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610 Kreuzinger, N., Fuerhacker, M., Grillitsch, B., Scharf, S., Uhl, M., Gans, O., 2007. Methodological approach towards the environmental significance of uncharacterized substances — quaternary ammonium compounds as an example. Desalination 215, 209-222. https://doi.org/10.1016/j.desal.2006.10.036

614 Laviale, M., Prygiel, J., Lemoine, Y., Courseaux, A., Créach, A. 2009. Stream periphyton photoacclimation response in field conditions: effect of community development and seasonal changes. Journal of Phycology 45, 1072-1082. https://doi.org/10.1111/j.1529-8817.2009.00747.x

617 Kahrilas, G.A., Blotevogel, J., Stewart, P.S., Borch, T., 2015. Biocides in Hydraulic Fracturing Fluids: A Critical Review of Their Usage, Mobility, Degradation, and Toxicity. Environ. Sci. Technol. 49, 16-32. https://doi.org/10.1021/es503724k

620 Kim, M., Hatt, J.K., Weigand, M.R., Krishnan, R., Pavlostathis, S.G., Konstantinidis, K.T., 2018. Genomic and Transcriptomic Insights into How Bacteria Withstand High Concentrations of Benzalkonium Chloride Biocides. Appl. Environ. Microbiol. 84. https://doi.org/10.1128/AEM.00197-18

624 Kreuzinger, N., Fuerhacker, M., Scharf, S., Uhl, M., Gans, O., Grillitsch, B., 2007. Methodological approach towards the environmental significance of uncharacterized substances — quaternary ammonium compounds as an example. Desalination 215, 209-222. https://doi.org/10.1016/j.desal.2006.10.036

628 Kümmerer, K., Eitel, A., Braun, U., Hubner, P., Daschner, F., Mascart, G., Milandri, M., Reinthaler, F., Verhoef, J., 1997. Analysis of benzalkonium chloride in the effluent from European hospitals by solid-phase extraction and high-performance liquid chromatography with post-column ion-pairing and fluorescence detection. Journal of Chromatography A 774, 281-286. https://doi.org/10.1016/S0021-9673(97)00242-2

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645 https://doi.org/10.1199/tab.0161

646 Longcore, T., Rich, C., 2004. Ecological light pollution. Frontiers in Ecology and the
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648 Lorenzen, C.J., 1967. Determination of chlorophyll and pheo-pigments: Spectrophotometric
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650 Maggi, E., Serôdio, J., 2020. Artificial Light at Night: A New Challenge in Microphytobenthos
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652 Martínez-Carballo, E., Sitka, A., González-Barreiro, C., Kreuzinger, N., Fürhacker, M., Scharf, S.,
653 Gans, O., 2007. Determination of selected quaternary ammonium compounds by liquid
654 chromatography with mass spectrometry. Part I. Application to surface, waste and indirect
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656 https://doi.org/10.1016/j.envpol.2006.04.033

657 Mazzella, N., Fadhlaoui, M., Moreira, A., Morin, S., 2023a. Molecular species composition of polar lipids
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659 Analytical Chemistry 5:e27. https://doi.org/10.7717/peerj-achem.27

660 Mazzella, N., Vrba, R., Moreira, A., Creusot, N., Eon, M., Millan-Navarro, D., et al. 2023b. Mixed light
661 photoperiod and biocide pollution affect lipid profiles of periphyton communities in freshwater
662 ecosystems. Journal of Hazardous Materials Advances 12,100378.
663 https://www.sciencedirect.com/science/article/pii/S2772416623001493

664 Mohapatra, S., Xian, J.L.L., Galvez-Rodriguez, A., Ekande, O.S., Drewes, J.E., Gin, K.Y.-H., 2024.
665 Photochemical fate of quaternary ammonium compounds (QACs) and degradation pathways
666 prediction through computational. Journal of Hazardous Materials 465, 133483.
667 https://www.sciencedirect.com/science/article/pii/S030438942400061X

668 Morin, S., Proia, L., Ricart, M., Bonnineau, C., Geiszinger, A., Ricciardi, F., Guasch, H., Romani,
669 A.M., Sabater, S., 2010. Effects of a bactericide on the structure and survival of benthic diatom
670 communities. Vie et Milieu / Life & Environment 60, 109.

671 Neury-Ormanni, J., Vedrenne, J., Wagner, M., Jan, G., Morin, S., 2020. Micro-meiofauna
672 morphofunctional traits linked to trophic activity. Hydrobiologia 847, 2725–2736.
673 https://doi.org/10.1007/s10750-019-04120-0

674 Pandey, L.K., Bergey, E.A., Lyu, J., Park, J., Choi, S., Lee, H., Depuydt, S., Oh, Y.T., Lee, S.M.,
675 Han, T. 2017. The use of diatoms in ecotoxicology and bioassessment: Insights, advances and
676 challenges. Water Research 118, 39-58. https://doi.org/10.1016/j.watres.2017.01.062.

677 Patrauchan, M.A., Oriel, P.J., 2003. Degradation of benzyldimethylalkylammonium chloride by
678 Aeromonas hydrophila sp. K. Journal of Applied Microbiology 94, 266–272.
679 https://doi.org/10.1046/j.1365-2672.2003.01829.x

680 Pérez, P., Fernández, E., Beiras, R., 2009. Toxicity of Benzalkonium Chloride on Monoalgal
681 Cultures and Natural Assemblages of Marine Phytoplankton. Water Air Soil Pollut 201, 319–
682 330. https://doi.org/10.1007/s11270-008-9947-x

683 Perkin, E.K., Hölker, F., Richardson, J.S., Sadler, J.P., Wolter, C., Tockner, K., 2011. The influence
684 of artificial light on stream and riparian ecosystems: questions, challenges, and perspectives.
685 Ecosphere 2, art122. https://doi.org/10.1890/ES11-00241.1

686 Pozo-Antonio, J.S., Sanmartín, P., 2018. Exposure to artificial daylight or UV irradiation (A, B
687 or C) prior to chemical cleaning: an effective combination for removing phototrophs from
688 granite. Biofouling 34, 851–869. https://doi.org/10.1080/08927014.2018.1512103
R Core Team (2020). — European Environment Agency [WWW Document], n.d. URL https://www.eea.europa.eu/data-and-maps/indicators/oxygen-consuming-substances-in-rivers/r-development-core-team-2006 (accessed 10.31.22).
Rabenau, H.F., Kampf, G., Cinatl, J., Doerr, H.W., 2005. Efficacy of various disinfectants against SARS coronavirus. Journal of Hospital Infection 61, 107–111. https://doi.org/10.1016/j.jhin.2004.12.023
Roux, C., Madru, C., Millan-Navarro, D., Jan, G., Mazzella, N., Moreira, A., Vedrenne, J., Carassou, L., Morin, S., 2024. Impact of urban pollution on freshwater biofilms: Oxidative stress, photosynthesis and lipid responses. Journal of Hazardous Materials 472:164523. https://doi.org/10.1016/j.jhazmat.2024.134523
Sakagami, Y., Yokoyama, H., Nishimura, H., Ose, Y., Tashima, T., 1989. Mechanism of resistance to benzalkonium chloride by Pseudomonas aeruginosa. Appl Environ Microbiol 55, 2036–2040.
Severina, I.I., Muntyan, M.S., Lewis, K., Skulachev, V.P., 2001. Transfer of Cationic Antibacterial Agents Berberine, Palmatine, and Benzalkonium Through Bimolecular Planar Phospholipid Film and Staphylococcus aureus Membrane. IUBMB Life 52, 321–324. https://doi.org/10.1080/152165401317291183
Singh, S.P., Singh, P., 2015. Effect of temperature and light on the growth of algae species: A review. Renew. Sustain. Energy Rev. 50, 431–444. https://doi.org/10.1016/j.rser.2015.05.024
Sreevidya, V.S., Lenz, K.A., Svoboda, K.R., Ma, H., 2018. Benzalkonium chloride, benzethonium chloride, and chloroxylenol - Three replacement antimicrobials are more toxic than triclosan and triclocarban in two model organisms. Environmental Pollution 235, 814–824. https://doi.org/10.1016/j.envpol.2017.12.108
Sütterlin, H., Alexy, R., Kümmerer, K., 2008. The toxicity of the quaternary ammonium compound benzalkonium chloride alone and in mixtures with other anionic compounds to bacteria in test systems with Vibrio fischeri and Pseudomonas putida. Ecotoxicology and Environmental Safety 71, 498–505. https://doi.org/10.1016/j.ecoenv.2007.12.015
Tamminen, M., Spaak, J., Tlili, A., Eggen, R., Stamm, C., Räsänen, K., 2022. Wastewater constituents impact biofilm microbial community in receiving streams. Science of The Total Environment 807, 151080. https://doi.org/10.1016/j.scitotenv.2021.151080
Tlili, A., Corcoll, N., Arrhenius, Å., Backhaus, T., Hollender, J., Creusot, N., Wagner, B., Behra, R., 2020. Tolerance patterns in stream biofilms link complex chemical pollution to ecological impacts. Environ. Sci. Technol. 54, 10745–10753. https://doi.org/10.1021/acs.est.0c02975
Ugwu, C.U., Aoyagi, H., Uchiyama, H., 2007. Influence of irradiance, dissolved oxygen concentration, and temperature on the growth of Chlorella sorokiniana. Photosynthetica 45, 309–311. https://doi.org/10.1007/s11099-007-0052-y
United States Environmental Protection Agency 2006. Reregistration eligibility decision for alkyl dimethyl benzyl ammonium chloride. (https://nepis.epa.gov). Accessed on 17 Feb. 2021.
US EPA, O., 2020. About List N: Disinfectants for Coronavirus (COVID-19) [WWW Document]. URL https://www.epa.gov/coronavirus/about-list-n-disinfectants-coronavirus-covid-19-0 (accessed 10.31.22).
van Wijk, D., Gyimesi-van den Bos, M., Garttener-Arends, I., Geurts, M., Kamstra, J., Thomas, P., 2009. Bioavailability and detoxification of cationics: I. Algal toxicity of alkyltrimethyl ammonium salts in the presence of suspended sediment and humic acid. Chemosphere 75, 303–309. https://doi.org/10.1016/j.chemosphere.2008.12.047
R Core Team (2020). — European Environment Agency [WWW Document], n.d. URL https://www.eea.europa.eu/data-and-maps/indicators/oxygen-consuming-substances-in-rivers/r-development-core-team-2006 (accessed 10.31.22).
Rabenau, H.F., Kampf, G., Cinatl, J., Doerr, H.W., 2005. Efficacy of various disinfectants against SARS coronavirus. Journal of Hospital Infection 61, 107–111. https://doi.org/10.1016/j.jhin.2004.12.023
Roux, C., Madru, C., Millan-Navarro, D., Jan, G., Mazzella, N., Moreira, A., Vedrenne, J., Carassou, L., Morin, S., 2024. Impact of urban pollution on freshwater biofilms: Oxidative stress, photosynthesis and lipid responses. Journal of Hazardous Materials 472:164523. https://doi.org/10.1016/j.jhazmat.2024.134523
Sakagami, Y., Yokoyama, H., Nishimura, H., Ose, Y., Tashima, T., 1989. Mechanism of resistance to benzalkonium chloride by Pseudomonas aeruginosa. Appl Environ Microbiol 55, 2036–2040.
Severina, I.I., Muntyan, M.S., Lewis, K., Skulachev, V.P., 2001. Transfer of Cationic Antibacterial Agents Berberine, Palmatine, and Benzalkonium Through Bimolecular Planar Phospholipid Film and Staphylococcus aureus Membrane. IUBMB Life 52, 321–324. https://doi.org/10.1080/152165401317291183
Singh, S.P., Singh, P., 2015. Effect of temperature and light on the growth of algae species: A review. Renew. Sustain. Energy Rev. 50, 431–444. https://doi.org/10.1016/j.rser.2015.05.024
Sreevidya, V.S., Lenz, K.A., Svoboda, K.R., Ma, H., 2018. Benzalkonium chloride, benzethonium chloride, and chloroxylenol - Three replacement antimicrobials are more toxic than triclosan and triclocarban in two model organisms. Environmental Pollution 235, 814–824. https://doi.org/10.1016/j.envpol.2017.12.108
Sütterlin, H., Alexy, R., Kümmerer, K., 2008. The toxicity of the quaternary ammonium compound benzalkonium chloride alone and in mixtures with other anionic compounds to bacteria in test systems with Vibrio fischeri and Pseudomonas putida. Ecotoxicology and Environmental Safety 71, 498–505. https://doi.org/10.1016/j.ecoenv.2007.12.015
Tamminen, M., Spaak, J., Tlili, A., Eggen, R., Stamm, C., Räsänen, K., 2022. Wastewater constituents impact biofilm microbial community in receiving streams. Science of The Total Environment 807, 151080. https://doi.org/10.1016/j.scitotenv.2021.151080
Tlili, A., Corcoll, N., Arrhenius, Å., Backhaus, T., Hollender, J., Creusot, N., Wagner, B., Behra, R., 2020. Tolerance patterns in stream biofilms link complex chemical pollution to ecological impacts. Environ. Sci. Technol. 54, 10745–10753. https://doi.org/10.1021/acs.est.0c02975
Ugwu, C.U., Aoyagi, H., Uchiyama, H., 2007. Influence of irradiance, dissolved oxygen concentration, and temperature on the growth of Chlorella sorokiniana. Photosynthetica 45, 309–311. https://doi.org/10.1007/s11099-007-0052-y
United States Environmental Protection Agency 2006. Reregistration eligibility decision for alkyl dimethyl benzyl ammonium chloride. (https://nepis.epa.gov). Accessed on 17 Feb. 2021.
US EPA, O., 2020. About List N: Disinfectants for Coronavirus (COVID-19) [WWW Document]. URL https://www.epa.gov/coronavirus/about-list-n-disinfectants-coronavirus-covid-19-0 (accessed 10.31.22).
van Wijk, D., Gyimesi-van den Bos, M., Garttener-Arends, I., Geurts, M., Kamstra, J., Thomas, P., 2009. Bioavailability and detoxification of cationics: I. Algal toxicity of alkyltrimethyl ammonium salts in the presence of suspended sediment and humic acid. Chemosphere 75, 303–309. https://doi.org/10.1016/j.chemosphere.2008.12.047
735 Wassenaar, T., Ussery, D., Nielsen, L., Ingmer, H., 2015. Review and phylogenetic analysis of
736 qac genes that reduce susceptibility to quaternary ammonium compounds in Staphylococcus
737 species. European Journal of Microbiology and Immunology 5, 44–61.
738 https://doi.org/10.1556/eujmi-d-14-00038

739 Wood, R.J., Mitrovic, S.M., Kefford, B.J. 2014 Determining the relative sensitivity of benthic
diatoms to atrazine using rapid toxicity testing: a novel method. Science of the Total
740 Environment 485-486,421-427. doi: 10.1016/j.scitotenv.2014.03.115.
741

742 Yang, Z., Geng, L., Wang, W., Zhang, J., 2012. Combined effects of temperature, light intensity, and nitrogen concentration on the growth and polysaccharide content of Microcystis aeruginosa in batch culture. Biochem. Syst. Ecol. 41, 130–135.
743
744 https://doi.org/10.1016/j.bse.2011.12.015

745 Yin, H., Wang, Lingling, Zeng, G., Wang, Longfei, Li, Y., 2022. The Roles of Different Fractions in Freshwater Biofilms in the Photodegradation of Methyl Orange and Bisphenol A in Aqueous Solutions. Int. J. Environ. Res. Public. Health 19, 12995.
746
747 https://doi.org/10.3390/ijerph192012995

750 Zhang, C., Cui, F., Zeng, G., Jiang, M., Yang, Z., Yu, Z., Zhu, M., Shen, L., 2015. Quaternary ammonium compounds (QACs): A review on occurrence, fate and toxicity in the environment. Science of The Total Environment 518-519, 352–362.
751
752
753 https://doi.org/10.1016/j.scitotenv.2015.03.007

754 Zulu, N.N., Zienkiewicz, K., Vollheyde, K., Feussner, I., 2018. Current trends to comprehend lipid metabolism in diatoms. Progress in Lipid Research 70, 1–16.
755
756 https://doi.org/10.1016/j.plipres.2018.03.001

757
758
759
735 Wassenaar, T., Ussery, D., Nielsen, L., Ingmer, H., 2015. Review and phylogenetic analysis of
736 qac genes that reduce susceptibility to quaternary ammonium compounds in Staphylococcus
737 species. European Journal of Microbiology and Immunology 5, 44–61.
738 https://doi.org/10.1556/eujmi-d-14-00038

739 Wood, R.J., Mitrovic, S.M., Kefford, B.J. 2014 Determining the relative sensitivity of benthic
diatoms to atrazine using rapid toxicity testing: a novel method. Science of the Total
740 Environment 485-486,421-427. doi: 10.1016/j.scitotenv.2014.03.115.
741

742 Yang, Z., Geng, L., Wang, W., Zhang, J., 2012. Combined effects of temperature, light intensity,
743 and nitrogen concentration on the growth and polysaccharide content of Microcystis
744 aeruginosa in batch culture. Biochem. Syst. Ecol. 41, 130–135.
745 https://doi.org/10.1016/j.bse.2011.12.015

746 Yin, H., Wang, Lingling, Zeng, G., Wang, Longfei, Li, Y., 2022. The Roles of Different Fractions
747 in Freshwater Biofilms in the Photodegradation of Methyl Orange and Bisphenol A in Aqueous
748 Solutions. Int. J. Environ. Res. Public. Health 19, 12995.
749 https://doi.org/10.3390/ijerph192012995

750 Zhang, C., Cui, F., Zeng, G., Jiang, M., Yang, Z., Yu, Z., Zhu, M., Shen, L., 2015. Quaternary
751 ammonium compounds (QACs): A review on occurrence, fate and toxicity in the environment.
752 Science of The Total Environment 518-519, 352–362.
753 https://doi.org/10.1016/j.scitotenv.2015.03.007

754 Zulu, N.N., Zienkiewicz, K., Vollheyde, K., Feussner, I., 2018. Current trends to comprehend
755 lipid metabolism in diatoms. Progress in Lipid Research 70, 1–16.
756 https://doi.org/10.1016/j.plipres.2018.03.001

757
758
759
760 **Appendices**

761 **A- Preliminary experiment**

762

763 This preliminary experiment aimed to test the sensitivity of our biofilm to
764 dodecyldimethylbenzylammonium (DDBAC) to select a sub-lethal concentration to be used in
765 the main experiment. We ran a dose-response experiment in which we tested nine
766 concentrations ranging between 17.5 µg.L⁻¹ and 175 mg.L⁻¹ (following a logarithmic increase)
767 for 4 hours. Each concentration was tested twice, one test at a mean light level of 16.86
768 µmol.s.m⁻² and the other kept in the dark.

769 Glass slides (26.5 cm × 6 cm) colonized by biofilms for three months were scraped and the
770 biofilms put into 500 mL of water. For each concentration, we contaminated 1.5 mL of the
771 biofilm solution. After four hours of contamination, we analysed the photosynthetic efficiency
772 of the samples with a Phyto-PAM (see section 2.3).

<figure><img src="image_7.png" /></figure>

773

<caption>Figure A. Dose-Response curve for photosynthetic efficiency in biofilms exposed to light or dark and increasing DDBAC concentration for 4 hours. Yeff = Photosynthetic efficiency.</caption>

776

777 The dose-response curve suggests an EC₅₀ of 112 ± 3 mg.L⁻¹ and a EC₅ of 34 ± 3 mg.L⁻¹ for
778 photosynthesis inhibition in the biofilm exposed to light. For the biofilm kept in the dark, we
779 found EC₅₀ of 151 ± 3 mg.L⁻¹ and a EC₅ of 58 ± 3 mg.L⁻¹. This result allowed us to choose the
780 concentration of 30 mg.L⁻¹ for the 10 days biofilm exposure which is the lowest concentration
781 with a minimum effect on photosynthetic efficiency.

782

783

784

785
760 **Appendices**

761 **A- Preliminary experiment**

762

763 This preliminary experiment aimed to test the sensitivity of our biofilm to
764 dodecyldimethylbenzylammonium (DDBAC) to select a sub-lethal concentration to be used in
765 the main experiment. We ran a dose-response experiment in which we tested nine
766 concentrations ranging between 17.5 µg.L⁻¹ and 175 mg.L⁻¹ (following a logarithmic increase)
767 for 4 hours. Each concentration was tested twice, one test at a mean light level of 16.86
768 µmol.s.m⁻² and the other kept in the dark.

769 Glass slides (26.5 cm × 6 cm) colonized by biofilms for three months were scraped and the
770 biofilms put into 500 mL of water. For each concentration, we contaminated 1.5 mL of the
771 biofilm solution. After four hours of contamination, we analysed the photosynthetic efficiency
772 of the samples with a Phyto-PAM (see section 2.3).

<figure><img src="image_30.png" /></figure>

773

<caption>Figure A. Dose-Response curve for photosynthetic efficiency in biofilms exposed to light or dark and increasing DDBAC concentration for 4 hours. Yeff = Photosynthetic efficiency.</caption>

776

777 The dose-response curve suggests an EC₅₀ of 112 ± 3 mg.L⁻¹ and a EC₅ of 34 ± 3 mg.L⁻¹ for
778 photosynthesis inhibition in the biofilm exposed to light. For the biofilm kept in the dark, we
779 found EC₅₀ of 151 ± 3 mg.L⁻¹ and a EC₅ of 58 ± 3 mg.L⁻¹. This result allowed us to choose the
780 concentration of 30 mg.L⁻¹ for the 10 days biofilm exposure which is the lowest concentration
781 with a minimum effect on photosynthetic efficiency.

782

783

784

785
786 B- Effects of DDBAC on chloroplasts

787

<figure><img src="image_8.png" /><figcaption>Figure B.1. Chloroplasts of diatoms in non-contaminated biofilm. x400 magnification</figcaption></figure>

788

789

790

<figure><img src="image_9.png" /><figcaption>Figure B.2. Chloroplasts of diatoms in DDBAC contaminated biofilm after 10 days of exposure. x400 magnification.</figcaption></figure>

791

792

793

794
786 B- Effects of DDBAC on chloroplasts

787

<figure><img src="image_31.png" /><figcaption>Figure B.1. Chloroplasts of diatoms in non-contaminated biofilm. x400 magnification</figcaption></figure>

788

789

790

<figure><img src="image_32.png" /><figcaption>Figure B.2. Chloroplasts of diatoms in DDBAC contaminated biofilm after 10 days of exposure. x400 magnification.</figcaption></figure>

791

792

793

794
795 C- HPLC gradients for the lipidomic analysis

796

<caption>Table C.1. HPLC gradients for phospholipid and glycolipid analysis.</caption>

798

<table><thead><tr><td>Time (min)</td><td>40 mmol.L<sup>-1</sup> Ammonium acetate buffer (%)</td><td>Acetonitrile (%)</td></tr></thead><tbody><tr><td>0</td><td>5</td><td>95</td></tr><tr><td>2</td><td>5</td><td>95</td></tr><tr><td>7</td><td>30</td><td>70</td></tr><tr><td>10</td><td>30</td><td>70</td></tr><tr><td>11</td><td>5</td><td>95</td></tr><tr><td>13.7</td><td>5</td><td>95</td></tr></tbody></table>

799

800

<caption>Table C.2. HPLC gradients for triglyceride analysis.</caption>

801

<table><thead><tr><td>Time (min)</td><td>Solvent A* (%)</td><td>Solvent B* (%)</td></tr></thead><tbody><tr><td>0</td><td>50</td><td>50</td></tr><tr><td>0.3</td><td>50</td><td>50</td></tr><tr><td>5.3</td><td>1</td><td>99</td></tr><tr><td>7.3</td><td>1</td><td>99</td></tr><tr><td>8.3</td><td>50</td><td>50</td></tr></tbody></table>
795 C- HPLC gradients for the lipidomic analysis

796

<caption>Table C.1. HPLC gradients for phospholipid and glycolipid analysis.</caption>

798

<table><thead><tr><td>Time (min)</td><td>40 mmol.L<sup>-1</sup> Ammonium acetate buffer (%)</td><td>Acetonitrile (%)</td></tr></thead><tbody><tr><td>0</td><td>5</td><td>95</td></tr><tr><td>2</td><td>5</td><td>95</td></tr><tr><td>7</td><td>30</td><td>70</td></tr><tr><td>10</td><td>30</td><td>70</td></tr><tr><td>11</td><td>5</td><td>95</td></tr><tr><td>13.7</td><td>5</td><td>95</td></tr></tbody></table>

799

800

<caption>Table C.2. HPLC gradients for triglyceride analysis.</caption>

801

<table><thead><tr><td>Time (min)</td><td>Solvent A* (%)</td><td>Solvent B* (%)</td></tr></thead><tbody><tr><td>0</td><td>50</td><td>50</td></tr><tr><td>0.3</td><td>50</td><td>50</td></tr><tr><td>5.3</td><td>1</td><td>99</td></tr><tr><td>7.3</td><td>1</td><td>99</td></tr><tr><td>8.3</td><td>50</td><td>50</td></tr></tbody></table>
<table><tr><td>9.8</td><td>50</td><td>50</td></tr></table>

802 * Solvent A: solution of acetonitrile/water/40 mmol.L⁻¹ ammonium acetate buffer
803 (600/390/10, v/v/v)

804 Solvent B: solution of isopropanol/acetonitrile/1 mol.L⁻¹ ammonium acetate buffer
805 (900/90/10, v/v/v)

806

807 D- Lipid standards

808 Polar lipid standards were purchased from Avanti Polar Lipids. Quantitations of
809 phosphatidylcholine (PC), phosphatidylethanolamine (PE) and phosphatidylglycerol (PG) were
810 respectively carried out with 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine or PC (16:0/18:1)
811 (850457), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine or PE (16:0/18:1)
812 (850757), and 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) or PG (16:0/18:1)
813 (840457).

814 For glycolipids, monogalactosyldiacylglycerol (840523), digalactosyldiacylglycerol (840524)
815 and sulfoquinovosyldiacylglycerol (840525) from plant extracts were used as standards.
816 Quantitation was performed with the following molecular species: MGDG (16:3_18:3) (63% of
817 the total MGDG standard), DGDG (18:3_18:3) (22% of the total MGDG standard), and SQDG
818 (34:3) (78% of the total MGDG standard).

819 1,2-diheptadecanoyl-sn-glycero-3-phosphocholine or PC (2x17:0) (850360) was used as
820 internal standard for PC phospholipids, 1,2-diheptadecanoyl-sn-glycero-3-
821 phosphoethanolamine or PE (2x17:0) (830756) was used as internal standard for PE
822 phospholipids, and 1,2-diheptadecanoyl-sn-glycero-3-phospho-(1'-rac-glycerol) or PG
823 (2x17:0) (830456) was used as internal standard for PG phospholipids, and both MGDG, DGDG
824 and SQDG glycolipids.

825 1,2-dipalmitoyl-sn-glycero-3-O-4'-(N,N,N-trimethyl)-homoserine or DGTS (2x16:0) (857464)
826 was used for the diacylglyceryltrimethylhomo-Ser (DGTS) lipids. 1,2-dipalmitoyl-sn-glycero-3-
827 O-4'-[N,N,N-trimethyl(d9)]-homoserine or DGTS-d9 (2x16:0) (857463) was used as internal
828 standard for DGTS lipids

829 Triglycerides were purchased from Sigma-Aldrich. Tristearin or TAG (3x18:0) (≥99%, T5016)
830 was used as the calibration standard while TAG (3x17:0) (≥99%, T2151) was used as the
831 internal standard.

832

833

834
<table><tr><td>9.8</td><td>50</td><td>50</td></tr></table>

802 * Solvent A: solution of acetonitrile/water/40 mmol.L⁻¹ ammonium acetate buffer
803 (600/390/10, v/v/v)

804 Solvent B: solution of isopropanol/acetonitrile/1 mol.L⁻¹ ammonium acetate buffer
805 (900/90/10, v/v/v)

806

807 D- Lipid standards

808 Polar lipid standards were purchased from Avanti Polar Lipids. Quantitations of
809 phosphatidylcholine (PC), phosphatidylethanolamine (PE) and phosphatidylglycerol (PG) were
810 respectively carried out with 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine or PC (16:0/18:1)
811 (850457), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine or PE (16:0/18:1)
812 (850757), and 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) or PG (16:0/18:1)
813 (840457).

814 For glycolipids, monogalactosyldiacylglycerol (840523), digalactosyldiacylglycerol (840524)
815 and sulfoquinovosyldiacylglycerol (840525) from plant extracts were used as standards.
816 Quantitation was performed with the following molecular species: MGDG (16:3_18:3) (63% of
817 the total MGDG standard), DGDG (18:3_18:3) (22% of the total MGDG standard), and SQDG
818 (34:3) (78% of the total MGDG standard).

819 1,2-diheptadecanoyl-sn-glycero-3-phosphocholine or PC (2x17:0) (850360) was used as
820 internal standard for PC phospholipids, 1,2-diheptadecanoyl-sn-glycero-3-
821 phosphoethanolamine or PE (2x17:0) (830756) was used as internal standard for PE
822 phospholipids, and 1,2-diheptadecanoyl-sn-glycero-3-phospho-(1'-rac-glycerol) or PG
823 (2x17:0) (830456) was used as internal standard for PG phospholipids, and both MGDG, DGDG
824 and SQDG glycolipids.

825 1,2-dipalmitoyl-sn-glycero-3-O-4'-(N,N,N-trimethyl)-homoserine or DGTS (2x16:0) (857464)
826 was used for the diacylglyceryltrimethylhomo-Ser (DGTS) lipids. 1,2-dipalmitoyl-sn-glycero-3-
827 O-4'-[N,N,N-trimethyl(d9)]-homoserine or DGTS-d9 (2x16:0) (857463) was used as internal
828 standard for DGTS lipids

829 Triglycerides were purchased from Sigma-Aldrich. Tristearin or TAG (3x18:0) (≥99%, T5016)
830 was used as the calibration standard while TAG (3x17:0) (≥99%, T2151) was used as the
831 internal standard.

832

833

834
835 E- Mass spectrometry parameters for lipid analysis

836

837 Table E.1. Mass spectrometry parameters for phospholipid and glycolipid analysis.

838

<table><thead><tr><th></th><th>Curtain gas</th><th>CAD</th><th>IonSpray</th><th>Temperature</th><th>Ion source gas 1</th><th>Ion source gas 2</th><th>Declustering potential</th><th>Collision energy</th></tr></thead><tbody><tr><td>MGDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-61 V</td><td>-28 V</td></tr><tr><td>DGDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-61 V</td><td>-28 V</td></tr><tr><td>SQDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-126 V</td><td>-66 V</td></tr><tr><td>PE</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-50 V</td><td>-50 V</td></tr><tr><td>PG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-100 V</td><td>-50 V</td></tr><tr><td>PC</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-100 V</td><td>-50 V</td></tr></tbody></table>

839

840

841 Table E.2. Mass spectrometry parameters for triglyceride analysis.

<table><thead><tr><th></th><th>Curtain gas</th><th>CAD</th><th>IonSpray</th><th>Temperature</th><th>Ion source gas 1</th><th>Ion source gas 2</th><th>Declustering potential</th><th>Collision energy</th></tr></thead><tbody><tr><td>Triglycerides</td><td>30 psi</td><td>3</td><td>+5000 V</td><td>450°C</td><td>30 psi</td><td>45 psi</td><td>50 V</td><td>38 V</td></tr></tbody></table>

842
835 E- Mass spectrometry parameters for lipid analysis

836

<caption>837 Table E.1. Mass spectrometry parameters for phospholipid and glycolipid analysis.</caption>

838

<table><thead><tr><th></th><th>Curtain gas</th><th>CAD</th><th>IonSpray</th><th>Temperature</th><th>Ion source gas 1</th><th>Ion source gas 2</th><th>Declustering potential</th><th>Collision energy</th></tr></thead><tbody><tr><td>MGDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-61 V</td><td>-28 V</td></tr><tr><td>DGDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-61 V</td><td>-28 V</td></tr><tr><td>SQDG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-126 V</td><td>-66 V</td></tr><tr><td>PE</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-50 V</td><td>-50 V</td></tr><tr><td>PG</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-100 V</td><td>-50 V</td></tr><tr><td>PC</td><td>30 psi</td><td>3</td><td>-4500 V</td><td>450°C</td><td>30 psi</td><td>60 psi</td><td>-100 V</td><td>-50 V</td></tr></tbody></table>

839

840

<table><caption>841 Table E.2. Mass spectrometry parameters for triglyceride analysis.</caption><thead><tr><th></th><th>Curtain gas</th><th>CAD</th><th>IonSpray</th><th>Temperature</th><th>Ion source gas 1</th><th>Ion source gas 2</th><th>Declustering potential</th><th>Collision energy</th></tr></thead><tbody><tr><td>Triglycerides</td><td>30 psi</td><td>3</td><td>+5000 V</td><td>450°C</td><td>30 psi</td><td>45 psi</td><td>50 V</td><td>38 V</td></tr></tbody></table>

842