© EDP Sciences, 2020

1 Introduction

The salinization of freshwater ecosystems is an increasing environmental problem worldwide, which is directly or indirectly associated with human activities. Intensive agricultural practices, modification of waterways, and the use of rock salt for road construction are examples of human activities that directly affect freshwater salinity (Kaushal et al., 2005; Lind et al., 2018; Nielsen et al., 2003). Moreover, sea-level rise (SLR) caused by climate change is increasingly leading to salinization of aquifers, estuaries, coastal lakes, wetlands, and rivers (Bhuiyan and Dutta, 2012; Mulamba et al., 2019; Schallenberg et al., 2003). The degree of SLR is expected to increase, and sea level is estimated to rise to 2 m in the worst-case scenarios (Bamber et al., 2019). The impact of SLR on freshwater salinization is especially relevant in lowland rivers, which will face more salt contamination than usual, sometimes up to many kilometers upstream (Bhuiyan and Dutta, 2012; Mulamba et al., 2019; Rice et al., 2014; Vu et al., 2018). Furthermore, reduced river flow and discharge during dry spells and droughts (climate change increases drought duration and severity) facilitate the intrusion of seawater (Chiogna et al., 2018).

Macrophytes play a crucial role in determining the characteristics and functions of aquatic ecosystems (Franklin et al., 2008). The abundance and distribution of macrophytes are closely associated with water body hydrology, morphology, nutrient availability, sediment characteristics, and physical and chemical parameters of water (color, turbidity, temperature, pH, and dissolved oxygen; Feldmann and Nõges, 2007; Hrivnák et al., 2009; Rameshkumar et al., 2019). Macrophytes occupy and colonize water bodies with favorable growth conditions, which vary according to the species (Cao et al., 2019). They respond to tolerate changes in environmental conditions, which may lead to alterations in macrophyte community composition (Barko et al., 1986; Søndergaard et al., 2010). When ambient conditions exceed tolerable levels, macrophyte growth is suppressed, and the plants eventually perish.

High salinity in freshwater bodies has adverse effects on freshwater ecosystems (Cañedo-Argüelles et al., 2019). This is especially true for aquatic plants, which, in contrast to many animals, are unable to migrate to unaffected sites. Plants respond to salinity changes by adjusting their morphology and physiology according to their salinity tolerance (Twilley and Barko, 1990) and salt concentration, which at extreme levels causes plant death. Therefore, the excessive and prolonged intrusion of saltwater into freshwater bodies is expected to cause a further increase in salinity, leading to the elimination of freshwater macrophytes; however, studies have shown that some levels of salinity are tolerable (Borgnis and Boyer, 2016; Thouvenot and Thiébaut, 2018). Future aquatic vegetation management strategies, especially in coastal areas, will have to consider the effects of altered salinity levels. Understanding the tolerance and responses of macrophytes to changes in salinity levels could assist with the monitoring and management of such ecosystems.

Myriophyllum spicatum L. and Ceratophyllum demersum L. are common submerged macrophytes species that are widely distributed in low-flow rivers, lakes, and ponds (Germ et al., 2006; Keskinkan et al., 2007). These two species have common characteristics, such as rapid growth, low light preference, and fragment distribution. However, M. spicatum is a rooted species, while C. demersum does not form roots and is found free-floating or anchored to the substrate. These two species also present distinct responses to environmental stresses, including different heavy metals absorption (Keskinkan et al., 2007), sensitivity to ultraviolet B radiation (Germ et al., 2006), and selenium toxicity (Mechora et al., 2011). Because of the characteristics referred to above, several studies have investigated the responses of M. spicatum and C. demersum to changes in environmental conditions, including salinity changes (Li et al., 2011; Thouvenot et al., 2012). However, little is known about the biochemical responses of these two species to salt stress, since most studies focused on morphological adjustments rather than biochemical changes under different salinity conditions (Borgnis and Boyer, 2016; Thouvenot et al., 2012; Thouvenot and Thiébaut, 2018).

Because little work has been done on the biochemical responses of M. spicatum and C. demersum to salt stress, we investigated the effects of exposing these two species to different salinity levels for a short time period as an initial study. We focused on changes in biochemical composition (hydrogen peroxide and antioxidant enzymes levels), pigmentation (chlorophyll a, chlorophyll b, carotenoids, and anthocyanins levels), and growth rate parameters of macrophytes maintained under controlled temperature, light, and nutrient conditions, but under different salinity conditions, for 5 days. Further, we investigated the relationship between parameters under salinity influence to evaluate the suitability of biochemical and pigmentation responses as biomarkers to manage these two macrophytes.

2 Materials and methods

2.1 Macrophyte stock culture

Macrophytes were collected in large quantities from river colonies of the Kanto region, Japan, and healthy plants were cultured in 20–30 L glass aquariums as the stock culture. Each stock tank was containing 4–5 cm layer of nutrient washed river sand and selected healthy plants were fixed to the sand with the help of a thumb forceps. Light was maintained at 80–85 μmol m⁻² s⁻¹ photosynthetically active radiation (PAR) intensity at plant depth provided by LED lamps with a color rendering index of 80 (Model LT-NLD85L-HN, Ohm Electric, Toshima, Japan). The light period was set at 12 h light, 12 h dark. Tanks were kept in a temperature-controlled experiment room at 25 ± 2 °C continuously. A commercial aquaponics solution (Hyponex concentrated nutrient solution, Hyponex, Osaka, Japan) at 5 ppm was used as a nutrient source.

2.2 Experimental plants and salinity treatments

Plant cuttings (∼8 cm) from healthy stock plants were obtained and washed with 5% Hoagland solution; any attached algae were removed. Cuttings were immediately planted in separate tanks filled with 5% Hoagland solution containing nutrients washed river sand layer (∼3 cm). Light conditions were the same as described for the stock tanks. Plant cuttings were kept in tanks for one week for acclimatization and then transferred to experimental tanks (25 × 15 × 15, height × width × length) containing nutrients washed river sand (∼3 cm) and 4 L of 5% Hoagland solution. In each experimental tank, 3 cuttings of C. demersum or 4 cuttings of M. spicatum were planted. Planting was done by inserting ∼2.5 cm of cutting into the sand with the help of a thumb forceps. Light conditions were maintained at 80–85 μmol m⁻² s⁻¹ PAR intensity at plant depth and 12:12 light cycle. Water temperature was maintained at 25 ± 1.0 °C. Each tank had a general-purpose aquarium air pump to maintain the CO₂ supply (Fig. 1). Plant cuttings were kept under these conditions for 3 days prior to the treatments. Any water loss due to evaporation was replenished with distilled water daily.

After 3 days, plants were exposed to different salinities using a mixture of salts at a fixed ratio to achieve the desired salinity conditions. The salt mixture contained: 2.08% NaCl, 0.94% MgCl₂⋅6H₂O, 0.132% CaCl₂⋅2H₂O, and 0.06% KCl (Rout and Shaw, 2001). The salts were dissolved in a beaker using nutrient solutions taken from each tank. Then, the solution was added back slowly into the respective tanks and slowly mixed with a plastic rod without disturbing the plants. The different salinity levels were: 0 ppt, 1.5 ± 0.1 ppt, 2.5 ±0.1 ppt, 5.0 ± 0.2 ppt, and 10 ± 0.5 ppt, which were confirmed using a portable salinity tester (Eutech Expert CTS Pocket Tester, Thermo Scientific, Waltham, MA, USA). All salinity exposure experiments were performed in triplicate. After 5 days of salinity exposure, plants had their length recorded and were put in resealable bags after blotting and stored in ice. Then samples were stored at −80 °C until further analysis.

Fig. 1 Representative images of experimental tanks containing the two species, Myriophyllum spicatum and Ceratophyllum demersum, after five days of continuous exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt). C. demersum plants exposed to 10 ppt salinity kept their leaves closed even when exposed to light; at 5 ppt leaves remained partially closed.

2.3 Elongation measurement

The length of each plant cutting was measured at the beginning of the experiment (when the salt mixture was added to the tank) and at the end of the experiment (on the 5th day of exposure). The length was measured using a millimeter-scale in relation to the sand layer in the experiment tank while plants were still inside the tank. The elongation of plants is expressed as percent change over initial length after 5 days exposure.

2.4 Pigments quantification

Photosynthetic pigment content was estimated after extracting pigments from samples using N, N-dimethylformamide. Pigments were extracted by incubating 150 mg of samples collected from the upper part of the plants with 5 mL of N, N-dimethylformamide for 24 h in the dark at room temperature (25–27 °C). Then, the optical absorptions were spectrophotometrically measured at wavelengths of 664, 647, and 480 nm (UV-1280, Shimadzu, Kyoto, Japan). The concentrations of chlorophyll a (Chl a), chlorophyll b (Chl b), and total carotenoids (Car) were calculated using the equation provided by Wellburn (1994) and are expressed as milligrams per gram of dry weight (mg/g DW) for each pigment.

Anthocyanin content was measured spectrophotometrically, as described in Nakata and Ohme-Takagi (2014) but with modifications. Approximately 50 mg of samples collected from the upper portion of the plants were pulverized in liquid nitrogen and mixed with 2 mL of extraction buffer containing 45% (v/v) methanol and 5% (v/v) acetic acid in distilled water. After centrifugation at 3,500 rpm for 15 min at 20 °C, the supernatant was collected, and optical absorption at 637 and 530 nm was measured (UV-1280, Shimadzu, Kyoto, Japan). Anthocyanin content was calculated, taking one anthocyanin unit as equivalent to one absorbance unit in 1 mL of extraction solution (Teng et al., 2005).

2.5 Hydrogen peroxide and antioxidant enzymes

The activities of antioxidant enzymes, guaiacol peroxidase (GPX), catalase (CAT), and ascorbate peroxidase (APX), as well as H₂O₂ levels, were measured spectrophotometrically (UV-1280, Shimadzu, Kyoto, Japan). For these measurements, approximately 200 mg of each sample were crushed in liquid nitrogen in the presence of polyvinylpyrrolidone. Extraction was done by adding 5 mL of 0.05 M phosphate buffer (pH 6.0) to the crushed samples. Then the extraction mixture was centrifuged at 3,500 rpm for 10 min at 4 °C, and the supernatant was collected and stored at −80 °C until further analysis.

Hydrogen peroxide content was measured by mixing 750 µL of the supernatant with 2.5 mL of 0.1% (w/v) titanium sulfate in 20% (v/v) H₂SO₄ (Satterfield and Bonnell, 1955) and incubating the assay mixture at room temperature (25–27 °C) for 30 min. Then, the assay mixture was read at 410 nm. Hydrogen peroxide concentration in samples was estimated by comparing absorbance values with those of an H₂O₂ standard curve and was expressed as µmol g⁻¹ FW.

Guaiacol peroxidase activity was measured by mixing 100 μL of extract supernatant with 40 µL of 30 mM H₂O₂ and 50 µL of 25 mM guaiacol (MacAdam et al., 1992). The reaction was initiated with the addition of supernatant into the reaction medium. The absorbance at 420 nm was recorded every 10 s for 3 min. Based on the rate of absorbance increase, GPX activity was expressed as μmol min⁻¹ g⁻¹ FW using the extinction coefficient of 26.6 mmol⁻¹ cm⁻¹.

Catalase activity was measured by reacting 100 μL of 10 mM H₂O₂, 2 mL of 100 mM potassium phosphate buffer (pH 7.0), and 500 μL of extract supernatant (Aebi, 1984). The reaction was initiated with the addition of supernatant into the reaction medium. The absorbance at 240 nm was recorded every 10 s for 3 min. Based on the rate of absorbance decrease, CAT activity was expressed as μmol min⁻¹ g⁻¹ FW using the extinction coefficient of 40 mmol⁻¹ cm⁻¹.

Ascorbate peroxidase was measured according to Nakano and Asada (1981). The reaction mixture contained 100 μL of extract supernatant, 200 μL of 0.5 mM ascorbic acid in 50 mM potassium phosphate buffer (pH 7.0), and 2 mL of 50 mM potassium phosphate buffer (pH 7.0). The assay was started with the addition of 60 μL of 1 mM H₂O₂. The absorbance at 290 nm was recorded every 10 s for 3 min. Based on the rate of absorbance decrease, APX activity was expressed in μmol min⁻¹ g⁻¹ FW using the extinction coefficient of 2.8 mmol⁻¹ cm⁻¹.

2.6 Data analysis

The levels of antioxidant enzymes, H₂O₂, and anthocyanin in plants were normalized relative to those in the control group (0 ppt salinity). This was done by dividing the results of each salinity exposure group by those of the corresponding 0 ppt exposure group for each replicate. Data were analyzed using ANOVA followed by the Duncan post hoc test. Significant differences between experimental groups were evaluated using Student's t-test assuming equality of variance. Regression relationship analysis were used to evaluate relationships between parameters (H₂O₂, antioxidative enzymes, photosynthetic pigments, anthocyanin, and elongation). For all analyses, P values below 0.05 were considered to indicate statistically significant differences. Statistical comparisons were performed using IBM SPSS Statistics, version 25 (IBM, Armonk, NY, USA). Regression relationships and descriptive statistical analyses, as well as data visualization, were calculated and presented using Microsoft Excel 2016 (Microsoft, Redmond, WA, USA).

3 Results

Elongation rates of M. spicatum were significantly reduced in plants exposed to any salinity level compared with that of control plants maintained at 0 ppt (Fig. 1, ANOVA P < 0.01 for 5 and 10 ppt; ANOVA, P < 0.05 for 1.5 and 2.5 ppt; Fig. 2). There was no significant difference between M. spicatum groups exposed to 1.5 ppt, 2.5 ppt, and 5 ppt salinities (ANOVA, P > 0.05). The elongation of M. spicatum plants exposed to 10 ppt salinity was lower than that of all other groups (ANOVA, P < 0.01). Compared with the control group, the elongation of M. spicatum in the 10 ppt group decreased by 8.6-fold. The elongation rate of C. demersum was not significantly affected by the exposure to 1.5 or 2.5 ppt salinities (ANOVA, P > 0.05; Fig. 2). Elongation was significantly reduced in plants exposed to 5 ppt salinity compared with that of plants in low salinity (0, 1.5, and 2.5 ppt) groups, (ANOVA, P < 0.05), but it remained significantly higher than the elongation of plants in the 10 ppt group (ANOVA, P < 0.01). The elongation rate of C. demersum exposed to 10 ppt salinity was approximately 3.5-fold lower than that of plants in the 0 ppt group (Fig. 2). Further, the usual leaf-closing mechanism (closed in the dark and open under light) of C. demersum was altered at 5 ppt and 10 ppt salinities: leaves remained fully closed under 10 ppt salinity exposure and partially closed under 5 ppt salinity exposure, regardless of light or dark phase. Under 1.5 ppt and 2.5 ppt salinities exposure, the leaf-closing mechanism of C. demersum was unaffected (Fig. 1).

Chl a level in M. spicatum differed between 2.5 ppt and 5 ppt salinity levels (t-test, P < 0.05). The rest of the salinity levels were statistically not different (ANOVA, P > 0.05; Fig. 3a). The Chl b levels of 5 ppt and 10 ppt groups were increased significantly than other groups (ANOVA, P < 0.05; Fig. 3b). In C. demersum, Chl a and Chl b levels of 1.5 ppt and 2.5 ppt salinities were not different from 0 ppt (ANOVA, P > 0.05; Fig. 3a and b). However, Chl a and Chl b levels significantly increased in C. demersum exposed to 5 ppt and 10 ppt salinities compared with other experimental groups (ANOVA, P < 0.01). There was no significant difference in Chl a and Chl b levels of C. demersum exposed to 5 ppt and 10 ppt salinities (t-test, P > 0.05). The levels of Car in M. spicatum were not significantly affected by any of the salinity groups compared with the 0 ppt group (ANOVA, P > 0.05; Fig. 3c). In the case of C. demersum, Car content did not differ between 0 ppt and 1.5 ppt groups (t-test P > 0.05; Fig. 3c). Plants exposed to 2.5 ppt and 10 ppt salinity had higher Car levels than those in 0 ppt and 1.5 ppt groups (ANOVA, P < 0.01). The C. demersum exposed to 5 ppt salinity had the highest Car content, which is higher than the Car contents of other salinity groups (ANOVA, P < 0.01; Fig. 3c).

Anthocyanin content in M. spicatum showed an increasing trend as salinity increased (Fig. 4). In plants exposed to 2.5 ppt, 5 ppt, and 10 ppt salinities, anthocyanin levels increased significantly compared with the 0 ppt group (ANOVA, P < 0.01), but there was no significant difference between these three groups (ANOVA, P > 0.05). No significant difference was observed between M. spicatum exposed to 1.5 ppt salinity and other groups due to high variability (t-test, P > 0.05). The concentration of anthocyanins in M. spicatum exposed to 10 ppt salinity increased significantly compared with that in the 0 ppt and 2.5 ppt groups (ANOVA, P < 0.05), whereas anthocyanin content in the 1.5 ppt and 5 ppt groups did not differ significantly (ANOVA, P > 0.05). A different pattern was observed for C. demersum, for which the anthocyanin content reached the highest value in plants exposed to 1.5 ppt salinity, which was significantly higher than all other groups (ANOVA, P < 0.05). Anthocyanin levels tended to decrease in C. demersum exposed to 5 ppt and 10 ppt salinities compared with the 0 ppt group, but statistically significant changes were detected only for 10 ppt exposure (ANOVA, P < 0.05; Fig. 4).

Hydrogen peroxide levels tended to decrease in M. spicatum exposed to 1.5 ppt, 2.5 ppt, and 5 ppt salinities compared with those in the control group (Fig. 5). In 1.5 ppt and 2.5 ppt groups, H₂O₂ content was significantly lower than that in plants maintained at 0 ppt (ANOVA, P < 0.05). There was no significant difference in H₂O₂ content between the 5 ppt group and the control group (P > 0.05). Myriophyllum spicatum exposed to 10 ppt salinity had the highest H₂O₂ content, which was significantly higher than that in all other groups (ANOVA, P < 0.01). In the case of C. demersum, H₂O₂ concentration showed a decreasing trend as salinity increased (Fig. 5). Hydrogen peroxide levels significantly decreased in C. demersum exposed to 5 ppt and 10 ppt salinities compared with those in the 0 ppt group (ANOVA, P < 0.01). Plants in the 5 ppt and 10 ppt groups did not differ significantly from each other (t-test, P > 0.05). Ceratophyllum demersum plants exposed to either 1.5 ppt or 2.5 ppt salinity were not significantly different from the control group in terms of H₂O₂ content (ANOVA, P > 0.05).

In M. spicatum, GPX activity showed an increasing trend as salinity increased up to 2.5 ppt when it reached the highest level (Fig. 6a); GPX activity was significantly higher in plants exposed to 2.5 ppt salinity than it was in all other groups (ANOVA, P < 0.05). In 1.5 ppt and 2.5 ppt groups, GPX activity was significantly higher than that in the control group (ANOVA, P < 0.01). As salinity increased to 5 ppt and 10 ppt, GPX activity tended to decrease and stabilize at levels similar to those observed in plants exposed to 1.5 ppt salinity. Nevertheless, GPX activity was still higher in M. spicatum exposed to 5 ppt and 10 ppt salinities than in control plants. (P < 0.05). With the exception of the 2.5 ppt group (t-test, P > 0.05), GPX activity increased in C. demersum exposed to saltwater (1.5 ppt, 5 ppt, and 10 ppt) compared to that in the control group (ANOVA, P < 0.01). Guaiacol peroxidase activity increased by 6.9-fold and 2.4-fold in C. demersum exposed to 10 ppt and 5 ppt salinities, respectively, compared with the control group (ANOVA, P < 0.01; Fig. 6a).

The exposure to saltwater at all levels significantly increased CAT activity in M. spicatum compared with the 0 ppt group (ANOVA, P < 0.01; Fig. 6b). There were no significant differences between 1.5 ppt and 2.5 ppt groups (t-test, P > 0.05) and between 5 ppt and the 10 ppt groups (t-test, P > 0.05) in terms of CAT activity. The highest CAT activity levels were observed in M. spicatum exposed to 5 ppt salinity. The 5 ppt and 10 ppt groups were significantly higher than that in 1.5 ppt and 2.5 ppt groups (ANOVA, P < 0.05). For C. demersum, CAT activity significantly increased in plants exposed to 1.5 ppt, 2.5 ppt, and 10 ppt salinities compared with that in the 0 ppt group (ANOVA, P > 0.05; Fig. 6b). Catalase activity in the 5 ppt group was not significantly different from that in the 0 ppt, 1.5 ppt, and 2.5 ppt groups (ANOVA, P > 0.05). The highest CAT activity for C. demersum was recorded in the 10 ppt group but was not significantly different from that in 1.5 ppt and 2.5 ppt groups (ANOVA, P > 0.05; Fig. 6b).

In M. spicatum, APX activity was the highest in plants exposed to 1.5 ppt salinity, whose APX levels were significantly higher than all other groups (ANOVA, P < 0.01; Fig. 6c). From 1.5 ppt to 10 ppt, APX activity tended to decrease and return to control levels. Ascorbate peroxidase activity in the 2.5 ppt group was not different from that in the 0 ppt group (t-test, P > 0.05), but it was significantly higher than that in plants exposed to 5 ppt and 10 ppt salinities There was no significant difference between M. spicatum exposed to 5 ppt and 10 ppt salinities (t-test, P > 0.05), but these two groups had APX activity levels significantly lower than those in the other salinity groups (ANOVA, P < 0.05). The exposure to saltwater at all levels significantly increased APX activity in C. demersum compared with the control group (P < 0.01; Fig. 6c). From 1.5 ppt to 5 ppt, APX activity tended to reduce in C. demersum. There was no significant difference between APX activity in plants exposed to 1.5 ppt and 2.5 ppt salinities (t-test, P > 0.05), while APX activity in C. demersum exposed to 5 ppt salinity was significantly lower than that in the 1.5 ppt group (t-test, P < 0.05). There was no significant difference between C. demersum exposed to 2.5 ppt and 5 ppt salinities (t-test, P > 0.05). Ceratophyllum demersum plants exposed to 10 ppt salinity had higher APX activity than that in plants exposed to 2.5 ppt and 5 ppt; (ANOVA, P < 0.01); APX activity in the 10 ppt group was not significantly different from that of the 1.5 ppt group due to high variability (t-test, P > 0.05; Fig. 6c).

Fig. 2 Changes in plant length of Myriophyllum spicatum and Ceratophyllum demersum expressed as elongation (%) after five days of exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt). Error bars represent the standard deviation.

Fig. 3 Levels of photosynthetic pigments, (a) chlorophyll a (Chl a), (b) chlorophyll b (Chl b), and (c) carotenoids (Car), in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Results are expressed as micrograms per gram of fresh weight (μg/g FW). Error bars represent the standard deviation.

Fig. 4 Relative change of anthocyanin content in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Anthocyanin levels were normalized to those of the respective control group. Error bars represent the standard deviation.

Fig. 5 Relative change of hydrogen peroxide levels in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Hydrogen peroxide levels were normalized to those of the respective control group. Error bars represent the standard deviation.

Fig. 6 Relative change of antioxidant enzymes activities, (a) guaiacol peroxidase (GPX), (b) catalase (CAT), and (c) ascorbate peroxidase (APX), in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Enzymatic activities were normalized to those of the respective control group. Error bars represent the standard deviation.

3.1 Relationships between parameters

Hydrogen peroxide levels showed a strong regression linear relationship with elongation rates in M. spicatum (R² = 0.965), whereas this relationship followed a second order polynomial distribution (R² = 0.863) in C. demersum (Tab. 1). The relationships between H₂O₂ content and levels of photosynthetic pigments (Chl a, Chl b, and Car) were weak for M. spicatum (R² = 0.168, 0.227, and 0.209 respectively), whereas these relationships were linear and strong for C. demersum (R² = 0.942, 0.944, and 0.858 respectively). As in the H₂O₂ levels, the relationships between elongation rates and photosynthetic pigments (Chl a, Chl b, and Car) were weak for M. spicatum (R² = 0.190, 0.394, and 0.077 respectively), whereas these relationships followed second-order polynomial distributions and strong at for C. demersum (R² = 0.945, 0.930, and 0.968 respectively). Further, elongation rates had a strong linear relationship with anthocyanin content in M. spicatum (R² = 0.807) and a second-order polynomial relationship with anthocyanin content in C. demersum (R² = 0.701). In M. spicatum, GPX activity had a weak second-order polynomial relationship with elongation (R² = 0.4824), whereas this relationship was linear and strong in C. demersum (R² = 0.824). A weak and linear relationship was found between CAT activity and elongation in M. spicatum (R² = 0.423), whereas this relationship was strong and followed a second-order polynomial distribution in and C. demersum (R² = 0.719). In the case of APX activity and elongation rate, a scattered distribution was observed for both M. spicatum and C. demersum (R² = 0.399 and 0.485, respectively; Tab. 1).

4 Discussion

We assessed the effects of short-term exposure to different salinities on the growth rate and biochemical adjustments of two key macrophyte species, M. spicatum and C. demersum. The results indicated both shared and species-specific responses. The two species shared the same trend in elongation rates. On the other hand, the H₂O₂ and anthocyanin levels in response to salinity followed opposite trends. The antioxidant and photosynthetic pigments (Chl a, Chl b, and Car) responses were showed deviated responses between two species. Therefore, M. spicatum and C. demersum will exhibit different levels of physiological responses for salt stress. In terms of elongation, C. demersum had a higher tolerance to salinity compared with M. spicatum. Its elongation was not significantly affected at 1.5 ppt salinity, while that of M. spicatum was significantly reduced at this level. Nevertheless, both species maintained elongation above 10% after 5 days exposure to 2.5 ppt or lower salinities. Therefore, it is expected that M. spicatum and C. demersum can maintain adequate elongation rates when exposed to salt water up to 2.5 ppt for a short period of time. However, according to extended duration studies on macrophytes responses to salinity (Thouvenot and Thiébaut, 2018; Warwick and Bailey, 1997), deviations in responses of these two species expected from short duration exposure, which is to be tested in further experiments.

The levels of photosynthetic pigments (Chl a, Chl b, and Car) in both species increased at high salinity levels (thought the pigments levels of M. spicatum were statistically not significant). Photosynthetic pigments concentrations in macrophytes have been reported to either increase or decrease in response to salinity exposure depending on the species and light intensity (Twilley and Barko, 1990). For example, under low light conditions (light intensity not mentioned), Chl a content increases in M. spicatum as salinity increases (Twilley and Barko, 1990). In the present study, the 80–85 μmol m⁻² s⁻¹ photosynthetic active radiation density in which plants were maintained is appropriate for the normal functioning of plants but can be considered as low light compared with natural conditions. Alternatively, the increase in photosynthetic pigment levels can be related to the excess Mg²⁺ in the artificial seawater used in the present study, which contains 0.94% MgCl₂ and, thus, increases Mg²⁺ supply as salinity increases. Magnesium is the most important ion for the synthesis of the central structure of chlorophyll molecules, and its increased availability stimulates the formation of chlorophyll (Candan and Tarhan, 2003; Terpstra and Lambers, 1983). Therefore, we speculate that the rise in pigment content in response to high salinity might also be related to the increased Mg²⁺ supply. However, further experiments are needed to evaluate the effect of artificial seawater containing Mg²⁺ on plant Mg²⁺ content.

In macrophytes under stress exposure, H₂O₂ content usually increases with the severity of the stress until a tolerance threshold level is reached (Asaeda et al., 2018). However, H₂O₂ content tended to decrease as salinity increased in C. demersum. The anthocyanin content also followed similar trends to those of the H₂O₂ levels for both species. The anthocyanin content of plants usually increases in response to high salinity stress (Eryılmaz, 2006), as well as to nutrient deficiency, cold stress, and excessive light (Ahmed et al., 2015; Liang and He, 2018; Pietrini et al., 2002). Increased anthocyanin content is also considered to be a strategy to enhance antioxidant defenses in plants (Liu et al., 2018) in response to reactive oxygen species (ROS) accumulation in plants under stress (Xu and Rothstein, 2018).

Antioxidant enzyme activities were not correlated with higher salinity or H₂O₂ content. Moreover, changes in the activity of antioxidant enzymes in response to high salinity differed between species. For example, CAT activity increased under all salt exposure conditions in M. spicatum, whereas it was rather stable in C. demersum. The patterns of GPX and APX activities changes were distinct between the two species, indicating that the antioxidant systems of M. spicatum and C. demersum respond differently to the salinity stress. Changes in antioxidant enzyme activities reflect distinct physiological states under different salinity levels. The salinity exposure of some terrestrial plant species also exhibited increased antioxidant activities without relating to a salinity gradient (Hishida et al., 2014; Srivastava et al., 2015). Therefore, exposure to high salinity stress is expected to elicit different antioxidant responses depending on the salinity level and species, which may not follow a consistent trend.

4.1 Biochemical parameters and elongation relationships

Temperature, light intensity, light duration, growth stage, and nutrient supply are the main factors that determine plant responses to stresses. In the present study, plants were maintained under controlled conditions within the optimal ranges for both species. However, in the field, where environmental conditions frequently change, these macrophytes might respond differently to changes in salinity compared with the responses recorded here. Despite changing environmental conditions, the correlations between physiological and morphological parameters are expected to predict the condition of the species under stress exposure. In that regard, we found strong correlations between some parameters, although their levels fluctuated along the salinity gradient. The regression relationships between H₂O₂ levels and elongation rates, and anthocyanin contents and elongation rates are significant (Tab. 1), and this warrants further investigation into their suitability for evaluation of expected growth responses under field conditions.

The correlations between elongation rates and pigment levels in M. spicatum and C. demersum were described mostly by linear regression distribution and sometimes by second-order polynomial regression distributions. These relationships are candidate approaches to estimate the expected growth response of macrophytes to salinity influences based on biochemical parameters or vice versa. However, as our findings indicate, such correlations are species-specific. Therefore, any application of these correlations for macrophyte monitoring should be carefully selected in relation to the species. Moreover, the responses of other macrophytes to long-term salinity exposure are expected to differ from those reported here and, thus, warrant further research.

5 Conclusion

Findings of the present research indicate the occurrence of species-specific responses, even under lower salinity levels. These distinct responses are not always correlated with salinity, indicating that macrophytes reach specific physiological statuses at each salinity level. The species-specific responses of most parameters measured in the present study suggest the inapplicability of common biochemical responses across species. On the other hand, the correlations between H₂O₂ levels and elongation rates, and anthocyanin content and elongation rates for both species have potential applications in the monitoring and management of M. spicatum and C. demersum. These two macrophyte species should be further studied for long-duration salinity exposure to distinguish short duration and long duration exposure responses. There can be a desiccating effect of on plant physiology and morphology, especially at higher salinity exposure, which was not measured in the present experiment. Considering the desiccating effect in further studies is recommended for understanding the plant response mechanism under salinity stress.

Acknowledgments

This work was supported by the annual budget allocations of the Saitama University, Japan.

References

Supplementary Material

Supplementary Figure S1. Access here

All Tables

All Figures

	Fig. 1 Representative images of experimental tanks containing the two species, Myriophyllum spicatum and Ceratophyllum demersum, after five days of continuous exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt). C. demersum plants exposed to 10 ppt salinity kept their leaves closed even when exposed to light; at 5 ppt leaves remained partially closed.
In the text

	Fig. 2 Changes in plant length of Myriophyllum spicatum and Ceratophyllum demersum expressed as elongation (%) after five days of exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt). Error bars represent the standard deviation.
In the text

	Fig. 3 Levels of photosynthetic pigments, (a) chlorophyll a (Chl a), (b) chlorophyll b (Chl b), and (c) carotenoids (Car), in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Results are expressed as micrograms per gram of fresh weight (μg/g FW). Error bars represent the standard deviation.
In the text

	Fig. 4 Relative change of anthocyanin content in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Anthocyanin levels were normalized to those of the respective control group. Error bars represent the standard deviation.
In the text

	Fig. 5 Relative change of hydrogen peroxide levels in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Hydrogen peroxide levels were normalized to those of the respective control group. Error bars represent the standard deviation.
In the text

Fig. 6 Relative change of antioxidant enzymes activities, (a) guaiacol peroxidase (GPX), (b) catalase (CAT), and (c) ascorbate peroxidase (APX), in Myriophyllum spicatum and Ceratophyllum demersum after exposure to different salinity levels (0, 1.5, 2.5, 5, and 10 ppt) for five days. Enzymatic activities were normalized to those of the respective control group. Error bars represent the standard deviation.

In the text