Ecological scheme of cyanobacterial responses to hydrogen peroxide and antioxidant enzymes against diurnal light intensity

doi:10.21203/rs.3.rs-1529688/v1

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Ecological scheme of cyanobacterial responses to hydrogen peroxide and antioxidant enzymes against diurnal light intensity

https://doi.org/10.21203/rs.3.rs-1529688/v1

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Diurnal variations of the oxidative stress condition of cyanobacteria were measured by field observations and laboratory experiments. In the field experiment, on clear summer days, transparent bottles, filled with surface water, were set up near the water surface, and several depths to 1 m deep. One bottle from each depth was collected every three hours together with the measurement of the photosynthetically active radiation (PAR). In the laboratory experiment, two cyanobacterial species were exposed to gradually increasing then decreasing light intensities. The samples were analyzed with the hydrogen peroxide (H₂O₂) concentration, the catalase activities (CAT), protein and chlorophyll a (Chl-a) contents. Regardless of difference in cyanobacterial species, similar diurnal trends were obtained. Protein contents did not change for one day experiments. Biologically generated H₂O₂ increased following the PAR intensity variation, with the delay of a few hours. The H₂O₂ concentration per protein was given as an increasing function of the exposure period (T) to the PAR. The amount of Chl-a content declined with the H₂O₂ per protein value.

The model was developed to estimate the H₂O₂ per protein value, elucidating that the effect of the cyanobacterial biomass and the surface layer mixing on the oxidative stress and mortality.

cyanobacteria

hydrogen peroxide

photoinhibition

diurnal behaviors

antioxidant activity

oxidative stress

Cyanobacteria are photosynthetic prokaryotes found in all habitats, and these organisms play a significant role in maintaining the ecological balance of nature. Like other organisms, cyanobacteria face an array of abiotic and biotic stresses from the environment, including metal ion toxicity, salinization, temperature, light conditions, eutrophication, allelopathy, and pathogens¹. As an adaptive mechanism, cyanobacteria, and other species, exhibit various responses in morphology, biochemistry, physiology, and anatomy to counter these stresses. Among these responses, biochemical responses have widely been used as biomarkers to detect the stress on organisms^2,3. The concentration of reactive oxygen species (ROS), activity of antioxidant enzymes, amino acid profile, and gene expression are some of the biomarkers employed in stress detections in aquatic plants, algae, and cyanobacteria.

As with other organisms, ROS accumulation causes harmful impacts on cyanobacteria, such as protein denature, impaired photosynthesis, growth inhibition, and anatomical damage^4,5, although ROS are important for growth regulation and signaling mechanisms^6,7. When biological systems produce ROS in their metabolism⁶, biological systems can control excess ROS production with the inherent scavenging enzymes and nonenzymatic components^{2, 6, 8–10}. ROS production exceeds its scavenging capacity under a stress environment, consequently, ROS accumulate inside cells, and induces oxidative stress.

One of the most prominent ROS is H₂O₂. The observed H₂O₂ concentration of a water body is widely distributed, and the production rate is enhanced by the nutrient content of the water body¹¹. Hence, the light intensity also is an important factor for the growth of cyanobacteria^{12, 13}.

Cyanobacteria are sensitive to even a minor change in light intensity causing stress for them¹⁴. Cyanobacteria can conduct photosynthesis and respiration via an elaborate electron transport pathway¹⁵. Collecting solar energy at photosystem II (PSII) in the thylakoid membrane results in the oxidation of water molecules and the reduction of plastoquinone, a molecule involved in the electron transport chain. The produced electrons are transported to PSI, where they are consumed in the synthesis of carbohydrates. An overabundance of solar energy results in the generation of ROS, including superoxide radicals, as the energy transfer rate is limited due to the underutilization of energy absorbed by the PSII antennae complex in the PSII reaction center^{16, 17}, and damages cellular components, such as D1 protein, which otherwise recovers the damaged photosynthesis apparatus^{2, 18, 19}. Superoxide dismutase (SOD) catalyzes superoxide radicals into H₂O₂, which is then detoxified into water by antioxidant activities²⁰. This mechanism has been intensively studied in the last two decades for many species, including cyanobacteria^{4, 6, 9}.

The H₂O₂ content and the activity of H₂O₂ scavenging enzymes can be used as good measures to detect the stress levels of these organisms. The physiological condition of cyanobacteria is highly dependent on the H₂O₂, content per protein ratio (Asaeda et al) ²¹.

Cyanobacteria is more vulnerable to H₂O₂ toxicity than another phytoplankton^22–24. Advancement in artificial endorsement of H₂O₂ has been considered to suppress cyanobacteria bloom in natural water^22,25,26,27.

Associated with increasing solar radiation, H₂O₂ is intensively generated biologically^28–30, and high irradiance at the surface layer makes a significant impact on cyanobacterial physiology via H₂O₂ production.

At the same time, in natural water, H₂O₂ is produced by the photolysis of dissolved organic carbon (DOC) by UV^31–34. Therefore, naturally generated H₂O₂ may have a possibility to contribute the decline of cyanobacteria.

The intensity of solar radiation changes diurnally, and cyanobacteria, under these changing light conditions, may have differing capacities to scavenge the ROS^{34, 36}, consequently, may exhibit the different damages in the diurnal variations.

Although the focus of the literature has been on the role of ROS, especially H₂O₂ in plants, in responding to environmental stresses^37–39, the behavior of cyanobacteria under diurnal light regimes is not well documented. Thus, this research was designed to study (1) the effect of light regime changes on cyanobacteria, (2) the relationship between solar radiation, H₂O₂ concentrations, and the antioxidant enzyme activities of cyanobacteria, and (3) the possibility of applying an H₂O₂ concentration as a proxy to detect stress intensity in algal management.

Diurnal variation of protein contents is shown in Fig.1, compared with PAR intensity. The protein content was nearly twice as higher in 2019 than in 2020.

In both field and laboratory experiments, the total protein (TP) content was nearly constant throughout the experimental period regardless of protein content in the early morning (Fig. 1). A similar trend was obtained for the optical densities, OD₇₃₀. These results indicate that the cell density changed only slightly with time.

The trend ofH₂O₂ concentration presented in Fig.2 was similar between the laboratory and field experiments. Although the magnitude was different, light-induced H₂O₂ production clearly indicated that the H₂O₂production increased in parallel to the PAR increment in the morning, however, peaked at 14:00 to 15:00, which was after the peak in the highest solar radiation at 11:40.

In the laboratory experiment, theH₂O₂ concentration was highest at 15:00, later than the peak PAR intensity. Species specifically, it was slightly higher with P.ambiguum compared with M.aeruginosa. The maximum PAR intensity, whether 300 or 600mmol m^-2 s^-1, did not affect thefluctuating trends of H₂O₂ concentration.

Fig. 3 postulates the H₂O₂ concentration increment from the early morning value, normalized by the TP content with respect to the PAR intensity.

In parallel to the increasing PAR in the morning, theH₂O₂ per protein increased with the decline of the increasing rate. In the afternoon, although PAR declined, theH₂O₂ per protein further increased for a while, achieving the peak value of 4, at around 15:00, then declined. Thus, it showed a large anti-clockwise hysteresis curve. The H₂O₂ concentration depends on the light intensity, and generally low in the lower level of the water body with the low light intensity.

The diurnal trend of CAT activity is shown as a function of H₂O₂concentration in Fig. 5.

The CAT activity gradually increased in the morning until 15:00, with a sharp peak, followed by a declining trend of the variation of H₂O₂concentration thereafter. The H₂O₂ and CAT activity of both species were positively correlated (M. aeruginosa, R= 0.865, and P. ambiguous, R= 0.910) and were statistically significant (p< .001). There was a similar trend with the field observation

(Fig. 4).

Although similar trends were observed for the activities of other antioxidant enzymes such as guaiacol peroxidase (POD), and ascorbate peroxidase (APX). However, CAT was by far the largest (CAT/POD~100-700, CAT/APX~30-300, data are not shown).

Species specifically, even though the APX activity hardly changed between P. ambiguum and M. aeruginosa, the CAT activity was comparatively several times higher in M. aeruginosa than in P. ambiguum.

Chl-a and protein contents slightly decreased in a day with increasingH₂O₂ per protein (R= -0.536, t= 3.03, p < 0.01 for Chl-a and R = -0.400, t = 2.09, p < 0.05) (Fig. 5).

H₂O₂ source under high solar radiation in the field

The same trend of H₂O₂ concentration of both the field and laboratory conditions indicates that the oxidative stress of cyanobacteria varies diurnally.

The H₂O₂ concentration rose up to the magnitude of 100–200nmol L^-1 in both laboratory and field before exposure to PAR. In the field samples, it was about 100 nmol L^-1 in 2019 and less than 10 nmol L^-1 in 2020 before exposure to solar radiation in the morning. In the laboratory, it was approximately 80 mol L^-1. The life span of H₂O₂ is 4 to 20 h implies that some level of H₂O₂ concentration is maintained though daily production⁴⁰. The H₂O₂ per protein in the early morning samples is likely attributed to the H₂O₂produced in the previous days, either biologically or by the photolysis of DOC.

Except for the H₂O₂ concentration in the early morning, the H₂O₂ concentration increased by 80 to 160 nmol L^-1 at the highest radiation period (Fig. 2). This is the amount generated biologically exposed to the high solar radiation.

Although biomass was not measured in this study, one third to one half of the cyanobacteria biomass is composed of protein⁴¹. Thus, the H₂O₂ per protein value indicates two to three times the amount biologically generated by a single cell exposed to the PAR.

The maximum H₂O₂ amount corresponds to about 3 to 4 nmol H₂O₂ per protein (mg), indicating that about 1 nmol of H₂O₂ per biomass (mg) was generated and contained in the cells. Considering the nearly neutral buoyancy of the cyanobacteria biomass, the H₂O₂ concentration is equivalent to approximately 1mmol H₂O₂ L^-1.

Many studies have been conducted on the lethal H₂O₂ concentration of cyanobacteria, mainly via incubations in laboratories, with the endorsement of different concentrations of H₂O₂.

Specifically, the growth of cyanobacteria was suppressed with H₂O₂ concentrations of 30mmol L^{-1, 25,26}, 100mmol L^{-1 27}, 118mmol L^{-1 12}, 325mmol L^{-1 42}, and 275mmol L^{-1 43}.

The maximum H₂O₂ concentration in the present study, 1nmol H₂O₂biomass^-1 (mg) (~1mmol H₂O₂ L^-1), is several times higher than the lethal concentration of cyanobacteria.

H₂O₂ concentration was measured in other types of plant species, where approximately 1mmol H₂O₂ g^-1FW was reported, and the growth was inhibited over 10mmol H₂O₂g^-1FW^{39, 44, 45}.

Considering that cyanobacteria are more vulnerable to H₂O₂ toxicity than other plants, the lethal H₂O₂ concentration in the present study agrees well with other plants results.

In a day, the high H₂O₂ concentration period continued for several hours only and did not last long. Cyanobacteria in surface water is in the lethal condition during the period and Chl-a as well as protein contents substantially declined associated with the increasing H₂O₂ per protein (Fig. 4). In the present field observation, many dead cells were observed after the period.

Diurnal patterns of generation and detoxification of H₂O₂

Associated with the solar radiation intensity, the H₂O₂ per protein was fluctuatedin a day. H₂O₂ is catalyzed majorly by the CAT activity, the activity of which is associated with the amount of H₂O₂ after transmitted by the signal^46-48. The high correlation between the H₂O₂ concentration and antioxidant activities indicates that the signal transmission of elevated H₂O₂ concentration is relatively quick⁷.

Although the light intensity sharply changed from the increasing phase to a decreasing phase at 11:40, the H₂O₂ concentration continued to rise slightly later, until around 15:00, and then gradually turned into the decreasing phase (Fig. 2).

Similar to this study, a one-hour delay from the solar radiation peak was observed in the photosynthetic quantum yield of the Microcystis bloom in Lake Taihu^49,50. As the mechanism is unknown, the effect is empirically estimated.

The H₂O₂ production rate was relatively low with the same intensity PAR in the afternoon. With morning and afternoon values together, the produced H₂O₂ per protein is assimilated as a function of PAR, such as

H₂O₂ protein^-1 = PAR^0.21(1)

(R = 0.732, t = 4.05, p < 10^-3).

Fig. 6 shows the observed H₂O₂ protein^-1 normalized by equation (1), E (= PAR^0.21), as a function of the period from 6:00, T (hrs.). There was a significant increasing trend for the normalized H₂O₂ protein^-1, and is assimilated by

E = 1.2*sin(p*(T-3)/24)²(2)

(R = 0.743, t = 6.38, p < 10^-6)

The evaluation of H₂O₂ per protein

The solar radiation is reflected at the water surface and attenuated in water.

The PAR intensity above the water surface is approximately given by

I₀(T)= I₀ cos (q) (3)

whereqis the incident angle of the solar radiation, approximately given by abs ((12 – T)p/12), and T is hrs. from 6:00.

Reflection coefficient at the water surface, Fr, is approximately given by

Fr = F₀ + (1- F₀) (1-cos (q))⁵(4)

Thus, the PAR intensity at time, T, and the depth, z, is provided by

I (T, z) = I₀(T) (1-Fr) exp (-kz)(5)

Where I₀(T) is the PAR intensity just below the water surface, z is the depth (m), k is the extinction coefficient of water (1 m^-1), given as a function of cyanobacteria biomass (protein content), or the protein content, given as

k = 0.25* Protein (1 m^-1) (6)

where Protein is the protein content in mg L^-1 (R= 0.807, t= 3.87, p < 0.01).

The PAR in water was obtained with sufficient agreement (R = 0.900, t = 6.72, p < 10^-4 for 2019; R = 0.620, t = 2.98, p < 0.01 for 2020).

Simulated H₂O₂ protein^-1 by equation (2) is postulated in Fig. 7, compared with the observed values.

The sufficient agreement was achieved (R = 0.705, t =3.30, p < 0.001 for 2019; R = 0.728, t = 5.10, p < 10^-5 for 2020).

In stagnant water, thermal stratification forms near the surface by the supplied solar energy from the surface. However, due to the disturbances, such as wind or cooling, the close to the surface is often mixed, where cyanobacteria distribution becomes relatively homogeneous.

H₂O₂ per protein in the mixed layer is postulated as a function of protein contents, 5, 10, and 30 mg L^-1, and thickness of the mixed layer, 0.1, 0.2 and 0.5m, in Fig. 8. At the surface, H₂O₂ per protein is independent of protein contents.

The H₂O₂ per protein value decreases with increasing mixed layer thickness, as the light intensity is attenuated in the deep layer. With low density of cyanobacteria, or low protein density, such as 5 to 10 mg L^-1, the H₂O₂ per protein values do not decrease much even deeply mixed because the light intensity is high in the deep layer due to the low attenuation rate. While with high density of cyanobacteria, H₂O₂ per protein substantially declines with increasing thickness of the mixed layer. This indicates that the mortality rate of cyanobacteria is higher with low density and is suppressed by rising cyanobacterial biomass.

The vertical migration of cyanobacteria

Lake cyanobacteria exhibit vertical migration behavior as an ecological strategy to minimize predation pressure, nutrient limitations, competition, and so forth^51-55. In most cases, cyanobacteria move to the deepest layers of the water body from 12:00 to 18:00, slightly later than the highest solar radiation period, and gradually rise thereafter. The present results indicate that the highest H₂O₂ protein^-1 occurs slightly later than the highest light time. The H₂O₂ concentration gradually lowers due to antioxidant activities afterwards. The cyanobacteria stay in the deeper zone in the early afternoon, likely to avoid the high solar radiation and the oxidative stress before the recovery of homeostasis by the increasing antioxidant activities⁵².

The effect of the species specific detoxification rate

In response to light intensity, M. aeruginosa has lower SOD activity per protein under the same light intensity compared with P. ambiguum. The generation rate of superoxide seems to be low in M. aeruginosa, likely because of the aggregation and forming scums of M. aeruginosa cells and being embedded by mucilage. These systems probably limit the light intensity, substantially weakening it by the time it arrives at the cell^{48, 55}.

The CAT activity per protein was twice to thrice times higher in M. aeruginosa than that of P. ambiguumfor the same H₂O₂ concentration. Although CAT is not a unique enzyme to catabolize H₂O₂, H₂O₂ was broken down more intensively into water and oxygen by M. aeruginosa than P. ambigum. Thus, the H₂O₂ concentration was lower with M.aeruginosa than with P.ambiguum in the period of high solar radiation. M. aeruginosa is reported to be weak under high H₂O₂concentration⁵⁶. However, together with higher antioxidant activities, it might exhibit a higher level of tolerance against light intensity than P. ambiguum, and it seems to enable M.aeruginosa to be dominant in the surface water under high solar radiation.

The efficiency of the destratification system on the blooming of these species

In the destratification system, to reduce algal biomass, the passive migration of algae into the deep layer by mechanical enforcement, such as artificial mixing, wind mixing, and night-time convection, is expected to prevent photosynthesis^57-59. The present study indicates a strong photoinhibition of cyanobacteria under solar radiation near the surface.As a result, slightly low light conditions are best for enabling cyanobacteria to grow. Compared with P.ambiguum, the destratification system will work more efficiently with M. aeruginosa, as it has a high tolerance to high solar radiation. However, the algal biomass can only be sufficiently reduced at a certain depth of the destratification layer. It is suggested that further studies estimate such aspects to potentially help to manage cyanobacterial blooms.

Regardless of difference in cyanobacterial species, similar diurnal trends were obtained. Protein contents did not change for one day experiments. Biologically generated H₂O₂ increased following the PAR intensity, with an approximately two to three-hour delay of the PAR exposure. Therefore, the oxidative stress indicator, the H₂O₂ concentration per protein showed the large hysteresis with respect to daytime PAR intensity variation, lower in the morning than in the afternoon with the same PAR intensity and is given as an increasing function of the T to the solar radiation. The amount of Chl-a concentration declined with increasing H₂O₂ per protein value. CAT activity, far largest compared with other antioxidant activities, solely follows the H₂O₂ concentration. Using these relationships, the prediction model to estimate H₂O₂ per protein value was developed with sufficient agreement with observation.

It was elucidated that the biologically generated H₂O₂ concentration was in the lethal level of cyanobacteria in the surface layer during high solar radiation period, and high biomass of cyanobacteria declines PAR intensity in water and avoids the oxidative stress.

These results indicate that H₂O₂ would be an effective biomarker to detect the stress level of cyanobacteria; the addition of H₂O₂ to freshwater may prove beneficial in designing the criteria for cyanobacteria management.

Ethical permission

This study has been approved by the Ministry of Environment of Japan (H30 Kokyo-Gaienbori ni okeru Kyokushoteki Ihicjiteki Aoko Taisaku-Gijutu Jisshou-Gyomu (Chidorighuchi Hibiyabori) ni Kakawaru Kikaku-Teian).

Field observation

Diurnal sampling in the field was performed. Field observations were made on a clear windless day on August 17, 2019, and August 29, 2020, at the boat jetty of the Chidorigafuchi moat of the Imperial Palace, Japan (35^o41’40” N; 139^o45’20” E) (Fig. 9). The moat was 2.5m deep in average. The highest solar radiation was observed at 11:40. Before 6:00, six semi cubic transparent bottles filled with surface water were set up at different depths, surface water (in 2020 experiment only), 0.1m, 0.3m (in 2020 experiment only), 0.5 m, and 1.0 m (in 2019 experiment only). At 9:00, 12:00, 15:00, 18:00 and 21:00, one bottle was collected from each depth and stored in a cool box (~0 ^oC) for transporting to the laboratory. At 13:30, another bottle was also collected, but only from a depth of 0.1m to see the effect of high solar radiation. Because all of the water is subject to convection during the night, the water was homogeneous at 6:00.

At each sampling time, light intensity, i.e., photosynthetically active radiation (PAR), was measured at each depth, along with temperature, pH, conductivity, and dissolved oxygen content (DO), by using a light intensity meter (Apogee, MQ-200, United States) and water quality meter (U-51, Horiba, Kyoto, Japan). On the observation day, the temperature was in the range of 28.3 ^oC to 32.6 ^oC in 2019, and 30.5 ^oC to 32.9^oC in 2020, while the pH fluctuated from 8.22 to 8.49 in 2019 and 8.85 to 9.45 in 2020. The turbidity changed from 91.8 to 99.6 NTU in 2019 and 27.5 to 30.2 NTU in 2020, and the conductivity varied from 21.1 to 25.5 ms/m in 2019 and 28.6 to 30.2 ms/m in 2020. The water samples were rich in Anabaena flos-aquae: the density at the surface and at 50 cm was 11,800 cells mL^-1(81% of all the cells) and 4500 cells mL^-1 (49% of all the cells), respectively, in 2019. Microcystis aeruginosa had a density of 55,300 cells mL^-1 (95% of all the cells) at the surface and 34,500 cells mL^-1 (90% of all the cells) at 50 cm, respectively, in 2020.

Laboratory Experiments

The laboratory experiments were conducted with more accurate condition, to confirm the rend observed in the field experiment.

Phormidium ambiguum (N2119) and Microcystis aeruginosa (N111) were obtained from the National Institute of Environmental Studies (NIES), Japan. These two strains were cultured and acclimatized for 14 days in an autoclaved BG 11 medium⁶⁰.

All experiments were conducted by using incubators (MIR-254, Sanyo, Tokyo, Japan) and in the nutrient level of BG-11 for 14 days at 20 ^oC with a 12h:12h light-and-dark (25 µmol s^-1m^-2) cycle, using a VBP-L24-C2 light source (Valore, Kyoto, Japan).

Each culture was divided into 36 flasks (10 ml) which were used as experimental units. During the experiment, the cultures were exposed to two levels of light: the intensity shifted from 0 µmol m^-2s^-1 at 6:00 to either 300 µmol m^-2s^-1 at 12:00 or 600 µmol m^-2s^-1 at 12:00 by changing the intensity to intervals of 25mmol m^-2s^-1 or 50 µmol m^-2s^-1, respectively. Then, the light intensity was decreased at the same rates until 18:00. The temperature was kept constant (20 ^oC) throughout the experiment. Every three hours, (i.e., at 6:00, 9:00, 12:00, 15:00, 18:00, and 21:00), three flasks were taken for a stress response analysis, while the remaining experimental units were manually shaken. The collected samples were subjected to bioassays, which are described later. The experimental units per each light condition were triplicated to confirm reproducibility.

Analyses

Chlorophyll concentration analysis

To begin, 10 mL of a homogenized treated cell suspension was centrifuged for 10 min at 3000 rpm, and the supernatant was removed. The cell pellet was washed using Milli-Q water, and then it was dried in a vacuum chamber. The dried pellet was vortexed using 3 mL of absolute acetone to dissolve it. Pigments of chlorophyll-a (Chl-a) were spectrophotometrically (UVmini-1240, Shimadzu, Japan) measured after being extracted into N,N-dimethylformamide by keeping them in darkness for 24 h⁶¹, and the specific extinction coefficient of 92.6 mL mg^-1cm^-1 at 663 nm for Chl-a was employed⁶².

Identification of cells and the cell density measurement

A small aliquot of 1 mL of each treated cell suspension was fixed at room temperature for 2 h using glutaraldehyde solution, and it was stored at 4 °C for counting. Cell identification and their densities were obtained by counting the fixed cyanobacterial cells using a light microscope supported with a digital camera (Nikon DXM 1200C) at its highest resolution (4116 x 3072 pixels), equipped with a bacteria-counting chamber with a depth of 0.02 mm (SLGC, Japan).

To estimate the growth of cyanobacteria, the OD₇₃₀ was measured by taking 1 mL of sample from each flask. The OD₇₃₀ was measured with a UV–Vis spectrophotometer (UVmini‐1240, Shimadzu, Kyoto, Japan) at an optical absorption wavelength of 730 nm, using a previously proposed methodology⁶³.

Total protein content analysis

In the same manner as the chlorophyll measurements, the cyanobacterial cells were harvested. Then TP was measured using the same method that was mentioned in⁶⁴, with minor modifications. After treating the cyanobacterial growth media, the cells were separated by centrifugation at 4 °C for 20 min at 10,000 rpm. After the pellet was washed once with distilled water, it was subjected to a freeze-thaw cycle and then grounded, using a motor. The total protein was extracted using a 0.5 M NaOH solution, and the extraction was centrifuged at 4 °C for 20 min at 10,000 rpm. The supernatant was used as crude protein extract, and the protein content was quantitatively analyzed with the aid of a Coomassie Bradford protein assay kit. The crude protein extract was stained with Coomassie (G-250) dye and incubated for 10 min at room temperature, and then the absorbance was measured at 595 nm using a spectrometer. The protein content was determined using a known concentration series of Albumin.

Stress assay

Enzyme extraction

The stress assays were performed by detecting the concentration of H₂O₂ and the activities of the antioxidant enzymes of catalase (CAT), superoxide dismutase (SOD), ascorbic peroxidase (APX), and peroxidase (POD). For the stress assays, the cells were extracted into an ice-cold phosphate buffer (pH 6.0, 50 mM) that contains polyvinylpyrrolidone (PVP) and masks the effect of phenolic compounds. Then, the extractions were centrifuged at 5,000 × g at 4 °C for 15 min. The supernatant was separated and incubated at -80 °C until further analysis. For each treatment, the extractions were performed in triplicate. All the results are expressed in protein basis.

H₂O₂Assay

The cellular H₂O₂ contents were estimated according to the e-FOX method⁶⁵, and the TiClO₄ method⁶⁶.

In e-FOX method, 1ml of the sample was taken from the flask and the supernatants were removed by centrifugation at 10,000 × g for 10 min at 4 °C. To extract the cellular H₂O₂, the cell pellets were homogenized in 1 mL of 0.1 M pH 7.0 phosphate buffer and centrifuged at 10,000 × g for 10 min at 4 °C. 25mM of ferrous ammonium sulfate is mixed with 20% H₂SO₄ to make reagent A alongside reagent B (sigma aldrich Co. Ltd) is prepared with help of 0.1M D-sorbitol, 0.125mM xylenol orange using Milli-Q water to make its final volume. A mixture of 1ml reagent B and an equal volume of 10μL reagent A and ethanol (99.5%) is used for a single analysis. The optical absorption mainly the xylenol orange complex that forms with oxidized metals such as ferric iron was measured at 560 nm using a spectrophotometer (UVmini-1240).

In TiClO₄ method, 1 mL was collected from each flask, and the supernatants were removed by centrifugation at 10,000 × g for 10 min at 4 °C. The cell pellets were washed once with ultrapure water (Milli-Q direct 5, Merck KGaA, Darmstadt, Germany). To extract the cellular H₂O₂, the cell pellets were homogenized in 1 mL of 0.1 M pH6.5 phosphate buffer and centrifuged at 10,000 × g for 10 min at 4 °C. A total of 750 μ L of 1% Titanium chloride in 20% H₂SO₄ (v/v) was then added to initiate the reaction. The optical absorption was measured at 410 nm using a spectrophotometer (UVmini-1240), following centrifugation (10,000 × g for 5 min) at room temperature (25 ± 2 °C).

The H₂O₂ concentration was determined using a standard curve of the optical absorption, prepared using a series of samples with known H₂O₂ concentration. However, the absorption at 560nm or 410 nm may have a possibility to include the effect of other soluble compounds^{65, 67, 68}. Thus, the H₂O₂ concentration was calculated from the slope of the standard curve obtained from the known H₂O₂ concentration, which was offset, derived by the intercept absorption rate with zero H₂O₂ concentration samples⁶⁷.

The results were compared with the results of both analyses, and suitable agreement was obtained (R = 0.85)⁶⁵.

CAT, APX, and POD activity assays

In the CAT activity assay, a total of 1 mL of each culture was centrifuged at 10,000 × g at 4 °C for 10 min. The supernatant was removed, and the cell pellets were homogenized in 1 mL potassium phosphate buffer (50 mM, pH 7.0), containing 0.1 mM EDTA. After centrifuging again (10,000 × g at 4 °C for 10 min), the supernatant was collected as the enzyme extract. The CAT activity was measured by reacting 15 μL of 750 mM H₂O₂, 920 μL of potassium phosphate buffer, and 65 μL of extract supernatant. Optical absorption was measured at 240 nm using UV mini-1240. The measurements were recorded every 10 s for 3 min, and the CAT activity was calculated using an extinction coefficient of 39.4 mM^-1cm⁶⁹.

For the APX assay, the reaction mixture contained 100 µL of enzyme extract, 200 µL of 0.5 mM ascorbic acid in 50 mM of potassium phosphate buffer (pH 7.0), and 2 mL of 50 mM potassium phosphate buffer (pH 7.0), mixed with 60 µL of 1 mM H₂O₂. The decrease in absorbance at 290 nm was recorded every 10 s for 3 min. The APX activity was calculated using an extinction coefficient of 2.8 mM^-1cm^{-1, 70}.

The POD activity was assayed following⁷¹, with slight modifications. The cyanobacteria cells were harvested by centrifuging 1-mL samples at 10,000 × g at 4 °C for 10 min and removing the supernatant and cell pellets, which were homogenized in 1 mL of potassium phosphate buffer (100 mM, pH 7.0). A total of 65 μl of enzyme extract was then mixed with 920 μL of potassium phosphate buffer (100 mM, pH 7) containing 20 mM of guaiacol. With the addition of 15 μL of 0.6% H₂O₂, the absorbance change was then recorded at 470 nm every 10 s for 3 min using UV mini-1240. POD activity was calculated with an extinction coefficient of 26.6 mM/cm.

Statistical analyses

The collected data were tested for normality with the Shapiro–Wilk’s test before the statistical analyses. All results were presented as the mean ± SD (n = 3). The data were subjected to a two-way analysis of variance (ANOVA), and the bivariate analysis was used and followed by Pearson's correlation method to evaluate the relationship among the parameters. The statistical analyses were performed with IBM SPSS V25.

Author contributions

T.A. contributed to the design of the experiment, observation, and writing the manuscript.

M.R. contributed to the revision of the manuscript.

A.M. carried out filed sampling and analyses.

D.H.L.A. carried out the experiment and analyses.

Acknowledgement

This work was financially supported by the Grant-in-Aid for Scientific Research (B) (19H02245) and Fund for the Promotion of Joint International Research (18KK0116) of Japan Society for the Promotion of Science (JSPS).

Conflicts of Interest

The authors declare no conflict of interest.

Data availability statement

The authors highly appreciate and stated that data is available for everyone in supplementary file named ‘Raw data.’

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Ecological scheme of cyanobacterial responses to hydrogen peroxide and antioxidant enzymes against diurnal light intensity

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