About 5% of total annual wastewater produced worldwide is saline (Vyrides, 2015). Several industries produce high or variable salinity effluents, such as oil refineries, aquaculture, seafood processing and tanneries (Vyrides, 2015; Lefebvre and Moletta, 2006). Seawater is also increasingly being used to alleviate water shortages leading to increased salinity in municipal wastewater treatment systems (Fu et al., 2022). Variable salinity is challenging for biological wastewater treatment processes because it can affect the metabolism and activity of the microorganisms by causing osmotic stress (Madigan et al., 2018; Sleator and Hill, 2002). In particular, hyperosmotic (salinity increase) changes are more detrimental than hypoosmotic (salinity decrease) changes (Csonka, 1989). The most common strategy is to gradually adapt the microorganisms to the new salinity, but the adaptation period can be very long (weeks to months) (Vyrides, 2015; Bassin et al., 2012; Gonzalez-Silva, 2016). Moreover, in many systems, it is common to have great variations due to process fluctuations rather than a gradual increase in salinity (Lefebvre and Moletta, 2006) and salinity may be different during different periods (Vyrides, 2015). Inoculation with salt-adapted microorganisms or with salt-adapted sludge has been shown to improve salinity acclimation (Cui et al., 2016; Sudarno et al., 2010; Vyrides and Stuckey, 2017). However, this strategy may not work during sudden increases in salinity and suitable inocula can be expensive or difficult to procure (Vyrides, 2015). Therefore, a more effective strategy is required for rapid salinity adaptation in microorganisms.

In principle, microbial cells must maintain an intracellular osmotic pressure greater than that of the environment to enable growth and cell division (Sleator and Hill, 2002). Microorganisms use two main strategies to adapt to salinity increase–i) the “salt-in” strategy and ii) the compatible solute strategy (Sleator and Hill, 2002; Csonka, 1989). In the “salt-in” strategy, the microorganisms increase their intracellular ion concentration to balance the external osmolarity (Vyrides and Stuckey, 2017). Owing to the high intracellular ionic strength, extensive structural adaptions are required, thus making this strategy exclusive to strictly halophilic bacteria (Sleator and Hill, 2002). In the second strategy, the primary response is an increase in the concentration of K⁺ (and its counter-ion glutamate), followed by a secondary response of an increase in the cytoplasmic concentration of compatible solutes through synthesis or uptake (Sleator and Hill, 2002). This strategy offers a greater degree of flexibility and is commonly used by halotolerant bacteria. Halotolerant microorganisms have a bi-phasic response to salinity increase. Compatible solutes, also known as osmolytes, are highly soluble molecules that do not interact with proteins, thus enabling them to accumulate at high intracellular concentrations without interfering with the cell function (Sleator and Hill, 2002). Not just one, but several osmolytes may be associated with salinity adaption, depending on the magnitude of salinity change and exposure time (Vyrides and Stuckey, 2017; Saum and Müller, 2007). The synthesis of osmolytes depends not only on the salinity but also on the availability of nutrients (Schimel et al., 2007). A significant amount of carbon and nitrogen are consumed during osmolyte production, leading to a 2-3 fold increase in the energetic cost for the microorganisms (Schimel et al., 2007). Thus, the synthesis of osmolytes is energetically expensive, and the uptake of osmolytes from the environment is an energetically cheaper option (Oren, 2011). As opposed to osmolytes that accumulate inside the cells, osmoprotectants are compounds that stimulate bacterial growth at high osmolality when provided in the growth medium (Wood, 2007). Adding the osmoprotectant in the beginning of a salinity exposure is preferable to a delayed addition to prevent the irreversible inhibition effects of salinity stress on cells (Vyrides and Stuckey, 2017). Among the commonly used osmolytes and osmoprotectants are sugars (sucrose, trehalose, etc.) and amino acids (glycine betaine, carnitine, proline, ectoine, choline, etc.) (Sleator and Hill, 2002; Vyrides and Stuckey, 2017; Oren, 1999). Several studies have shown that the exogenous addition of osmolytes can alleviate salinity stress across a wide variety of microorganisms (Vyrides and Stuckey, 2017).

While several microbial communities such as methanogens, denitrifiers and anammox have been found capable of osmoprotectant uptake, little is known about the salinity acclimation mechanisms in nitrifying microorganisms (Vyrides and Stuckey, 2017). Nitrification is a two-step process consisting of ammonia oxidation by ammonia oxidizing bacteria (AOB) and archaea (AOA), followed by nitrite oxidation by nitrite oxidizing bacteria (NOB). Within the genus Nitrospira, some species can perform both the nitrification steps (comammox) (Daims et al., 2015). Compared to heterotrophs, relatively less energy is produced during the autotrophic oxidation by nitrifiers, and the energy produced may not be sufficient to sustain growth at very high salt concentrations (Oren, 2011). Nonetheless, there are several marine nitrifiers that have an obligatory requirement for saline environments. In particular, AOA are known to thrive in seawater. Moreover, nitrifiers are found at high abundance in estuaries that frequently encounter salinity fluctuations, suggesting that several genera of nitrifying microorganisms are halotolerant (Bernhard and Bollmann, 2010; Santos et al., 2018; Ward et al., 2007). This is also supported by studies on nitrifying bioreactors where the same taxa were present across different salinities (Gonzalez-Silva, 2016; Navada et al., 2020a). A proteomic study suggested that the osmolytes sucrose and glycine were produced in response to a salinity increase in Nitrosomonas europaea (Ilgrande et al., 2018).

We hypothesize that the uptake of osmolytes may alleviate salinity stress in nitrifying microorganisms as it is energetically more favorable than de novo synthesis. We are aware of only a few studies on the effect of external osmolytes on nitrifiers, and with mixed results. In one study, the addition of a cocktail of osmolytes (betaine, trehalose, proline, 3-[N-morpholino]propanesulfonic acid, taurine, and γ-amino-n-butyric acid) did not enhance microcolony formation of N. europaea at salt concentration >0.1 M NaCl (∼6‰ salinity) (Wood and Sørensen, 1998). However, the study used a solution of sodium chloride, and was thus lacking K⁺ ions present in seawater that play an important role in osmoregulation (Sleator and Hill, 2002; Vyrides and Stuckey, 2017). In another study, Nitrobacter was capable of trehalose production and uptake of glycine betaine and sucrose from the medium (Vyrides and Stuckey, 2017). In a third study, the concentration of glutamine and proline increased in aerobic nitrifying granules, but no other osmolytes were analyzed (Wan et al., 2014). Recent studies show that the addition of osmolytes, such as glycine betaine and mannitol, can alleviate the inhibitory effects of salinity on AOB in activated sludge (Fu et al., 2022; Zuo et al., 2021). However, as far as we know, there exist no studies on the exogenous addition of osmolytes on nitrification in aerobic microbial communities with both nitrifying and heterotrophic bacteria, such as nitrifying biofilms.

Nitrifying biofilms are widely used in wastewater treatment, and nitrification is an important water treatment process for removing ammonia and nitrite that can be toxic to aquatic life. But nitrifying biofilms can be sensitive to salinity changes (Gonzalez-Silva, 2016; Navada and Vadstein, 2022). Therefore, it is important to minimize the negative impact of salinity increase on these biofilms. One potential strategy could be through the addition of osmolytes during a salinity increase. This is relevant, for instance, in recirculating aquaculture systems for anadromous species, such as Atlantic salmon, where the salinity must be changed during specific life stages of the fish (Navada et al., 2021), or during salinity fluctuations in industrial or municipal wastewater systems. The genome of some nitrifying bacteria have been reported to contain genes encoding the transport of osmolytes such as proline, ectoine, glutamate and betaine, suggesting that these can potentially be taken up for salinity acclimation (Wu et al., 2024). Thus, if the added osmolytes are effectively taken up by the nitrifiers for osmoregulation, it would improve the functionality of biofilm reactors, thus rendering water treatment systems more robust to salinity changes.

The goal of this study was to investigate the effect of the exogenous addition of osmolytes on nitrifying biofilms undergoing salinity increase from fresh- to seawater. Important osmolytes involved in bacterial osmoadaptation include glycine betaine, carnitine and proline (Sleator and Hill, 2002). The intracellular concentration of proline was reportedly increased during salinity increase in a study on aerobic nitrifying granules (Vyrides and Stuckey, 2017). N. europaea and Nitrosomonas winogradskyi have genes for production of glycine betaine and sucrose (Ilgrande et al., 2018), and Nitrobacter was found to accumulate these osmolytes from the medium under high salinity (Vyrides and Stuckey, 2017). Trehalose and ectoine are well known common osmoprotectants in several organisms (Elbein et al., 2003; Jebbar et al., 1992). Thus, a cocktail of the following common osmoprotectants was chosen: trehalose, sucrose, glycine betaine proline, carnitine, and ectoine. We hypothesized that the exogenous addition of osmolytes would alleviate salinity stress and lead to a lower reduction in nitrification rate upon salinity increase.

In the freshwater phase (day 2), there was no significant difference in any of the nitrification performance indicators (AOR, NO2_PR and NO3_PR) between treatments (Figure 1). The average ammonia oxidation rate was 0.76 ± 0.05 g m^-2 d^-1, and considerable nitrite accumulation was observed (NO2_PR = 0.24 ± 0.03 g m^-2 d^-1). This is likely because the refrigerated carriers did not get sufficient time to revive and the nitrite oxidizers could not catch up with the rapid increase in ammonia oxidation rate. Future studies should measure both ammonia and nitrite oxidation rates during the revival period, as nitrite oxidizers can take longer to re-establish. The reactors were well-aerated with dissolved oxygen of 80%–90% and the total nitrogen concentration was stable over the duration of the capacity test (Supplementary Figure A.4). This indicates that the ammonia-N was mainly converted to nitrite-N and nitrate-N, and the contribution of other nitrogen removal processes (such as denitrification) was negligible in this study.

On day 3 (the first day after seawater transfer), the seawater treatments (S and O) had a ∼50 and 94% reduction in the NO2_PR and NO3_PR, respectively. There was no significant difference between the seawater treatments with or without osmolytes for NO2_PR (p = 0.74) and NO3_PR (p = 0.20). This indicates that the exogenous addition of the osmolyte cocktail did not significantly improve the performance in nitrifying biofilms immediately after the salinity increase. There were some problems with ammonia measurements on day 3 and day 5. However, as presented above, nitrification was the major nitrogen conversion process in the reactors, and as such, the nitrite and nitrate production rates can be considered as suitable indicators of nitrification activity.

Surprisingly, 2 days after the salinity increase (day 5), nitrification was completely inhibited in the osmolyte treatment, indicated by the negligible nitrite and nitrate production rates (Figure 1). The NO2_PR in the O treatment was −0.03 g m^-2 d^-1, as the residual nitrite concentration was consumed within the first 8 h (from ∼1 to <0.5 mgN L^-1). This was significantly different (p < 0.001) from the positive NO2_PR (0.18 g m^-2 d^-1) in the S treatment. The nitrate concentration in both the seawater treatments was negligible during the first 8 h after exchange of medium, indicating a complete inhibition of nitrite oxidation. It is remarkable that contrary to our hypothesis, not only did the addition of osmolytes not improve salinity acclimation, it actually resulted in a lower nitrification activity compared to the treatment without osmolytes. This was likely due to the complex effects of osmolytes on the microbial community in the biofilms, as described further below.

We also observed a higher turbidity in the osmolyte treatment than in the other treatments on day 5. On measuring, the medium in the osmolyte treatment had an optical density (OD600) that was an order of magnitude higher than that of the other two treatments (0.050 compared to 0.007). This suggests a higher growth of free-living/pelagic bacteria in O compared to the other treatments. This was confirmed by light microscopy that showed a significantly higher abundance of bacterial cells (putative heterotrophs) in O. The increase in OD may also be attributed to the increased growth of heterotrophic microorganisms from the biofilm that were suddenly exposed to a carbon-rich medium. As the osmolytes are easily degradable organic compounds, they could have been consumed as substrates by the heterotrophs. The ratio of heterotrophs to nitrifiers can be as high as 4:1 even in nitrifying biofilms grown without an organic carbon source (Navada et al., 2019; Navada et al., 2020b). Thus, the osmolytes could have been taken up by both the planktonic and biofilm-attached heterotrophs. Although (Vyrides and Stuckey, 2017) argue that biodegradation of compatible solutes is less pronounced at higher salinities, we observed significant growth of planktonic heterotrophs in our study that can only be attributed to the uptake of osmolytes as substrate. As a majority of osmolyte studies have been on model heterotrophic bacteria like E. Coli (Rojas et al., 2014; Wood, 2015), the heterotrophs could also have acted as superior competitors relative to the nitrifiers for the uptake of osmolytes for osmoregulation. Both scenarios would have promoted the growth and activity of heterotrophs and thereby suppressed nitrification through microbial competition for resources, such as oxygen (Rittma et al., 2001). Thus, the osmoregulatory impact of the osmolytes on nitrification activity was confounded with the effect of the increased competition between the heterotrophs and nitrifiers.

As an osmolyte cocktail was used, we cannot determine which of the osmolytes contributed the most to the suppression in nitrification activity. The response of the microbial community would have been influenced by the type and concentration of osmolyte. It is possible that the growth of the heterotrophs could have been avoided by reducing the concentration of the highly degradable osmolytes (such as trehalose and sucrose) or by providing only the amine-based osmolytes. Future studies should investigate if there exist unique osmoprotectants that can selectively aid osmoregulation in nitrifiers, while not promoting the growth of heterotrophs. Further, some studies suggest that glycine betaine may be inhibited by other osmolytes (Feeney et al., 2014; Mendum and Smith, 2002). Thus, future studies should investigate different osmolytes separately under different ranges of concentrations, possibly at different levels of nitrogen loading or salinity stress.

Investigating the genomes of nitrifying species may provide an insight into the potential pathways for salinity acclimation in these microorganisms. A recent study investigated transporter genes in the metagenome-assembled genomes (MAG) of nitrifying bacteria subjected to salinity stress (Wu et al., 2024). The authors found nitrifying MAGs with genes encoding proline/betaine/ectoine transport (ProP), ectoine transport (EhuA) and glycine betaine transport (OpuA). Glutamate transporter genes have been reported in Nitrosomonas (Wu et al., 2024) and Nitrosopumilus (Kamanda Ngugi et al., 2015), suggesting that it could be a feasible osmoprotectant. Glutamate was not included in this study, and should be investigated. In future studies, metagenomic analysis could be performed first on the biofilm to identify the nitrifying microorganisms and if they encode any transporter genes for osmolytes. Based on this information, the respective osmolytes may be added during salinity stress to study if they are taken up during salinity acclimation. Metatranscriptomic and metaproteomic studies may help identify the genes and metabolites involved during salinity changes, thus giving a better understanding of the salinity acclimation mechanisms in nitrifying microorganisms.

Finally, the effect of osmotic stress on microorganisms growing in a biofilm can be more complex than for single free-living cells. Biofilms can be highly resistant to many different stresses, and biofilm formation is a protection strategy against stresses (Rode et al., 2020). Extracellular polymeric substances (EPS) in the biofilm matrix can protect against several types of stresses. Studies show that the EPS in the biofilm matrix near a bacterial cell may act as osmoprotectants, thus protecting against salinity stress (Seminara et al., 2012; Yan et al., 2017). Further, cells within a biofilm may respond to different stressors individually or collectively (Rode et al., 2020), leading to a higher versatility in response mechanisms. The collective response of microorganisms in biofilms to osmotic pressure changes needs to be better understood (Yan et al., 2017). Future studies should investigate the role of osmotic stress on the communication and response of the bacteria, along with the effect on matrix structure and function.

This study tested the exogenous addition of a cocktail of osmolytes as a potential sustainable strategy for salinity acclimation in nitrifying biofilms. The addition of an osmolyte cocktail did not improve salinity acclimation in nitrifying biofilms transferred from freshwater to seawater. On the contrary, after 48 h of exposure to seawater, the presence of osmoprotectants significantly decreased the nitrification activity. This was likely due to the uptake of osmolytes by the heterotrophs (as a substrate or for osmoregulation), which consequently led to a reduction of nitrification activity due to the competition between the heterotrophs and nitrifiers for oxygen and/or osmolytes. Further research is required on the metagenomic and metaproteomic level to investigate the physiological response of biofilm microorganisms to a salinity increase. Future studies should investigate the impact of individual osmoprotectants at different concentrations to identify osmolytes that are preferentially taken up by nitrifying microorganisms for salinity acclimation. The role of the biofilm matrix on salinity acclimation in nitrifying biofilms also needs further research.

In conclusion, the addition of an osmoprotectant cocktail did not succeed in improving the salinity tolerance of nitrifying biofilms. Nonetheless, our study highlights the complex contradicting effects of osmoprotectants on multispecies biofilms for water treatment, and opens the door for future studies on salinity acclimation in biofilms using osmoprotectants.