For the green synthesis of MgCl NPs, 300 mL of a solution containing 0.1 M MgCl₂ was prepared. 10 mL of pomegranate algae extract was added to the solution and refluxed for 2 h at 85°C under a reflux condenser. It was then placed in a reactor via Teflon tube. Hydrothermal reactions were performed at 180–195°C for 4 h to reduce nano-particle (NP) size. The precipitated MgO NPs were washed first via pure water and ethyl alcohol. They were preserved in an atmosphere free of moisture after being dried for 48 h at 60°C in an oven. MgS NPs were synthesized using the same procedure. 1 mol of Na₂S was added to the synthesis medium and the same process was repeated to synthesize MgS NPs. In the synthesis of CQD, the green algae (C. reinhardtii), known for its high non-toxic antioxidant content, was used as a carbon source. For this purpose, the algae extract was placed in a reactor containing sodium citrate as a reducing agent. CQD was synthesized by incubating at 180–195°C for 8 h.

Green-synthesised CQD, MgS and MgO NPs were characterized at the High Technology Application and Research Center of Eastern Anatolia (DAYTAM) at Atatürk University. X-ray microscopy (XRD) and FTIR analyses were performed for the characterization of CQD, MgS, and MgO NPs. The synthesized CQD, MgS, and MgO nanoparticles were characterized, including their size and morphology.

AOAC (71) guidelines were followed for chemical analyses. Kjeldahl was used to determine N content (AOAC, 71, Method 984.13). For the determination of Acid Detergent Fiber (ADF) and Neutral Detergent Fiber (NDF), Van Soest et al. (22) were used.

In vitro gas production is performed by taking rumen fluid from newly slaughtered cattle (as soon as they are slaughtered), as mentioned by Palangi et al. (10). Using a method validated by Menke and Steingass (23), it was found that 0.2 g of treated (CQD, MgS, and MgO nanoparticles at levels of 0.50, 100 ppm) and ground (1 mm) green algae (C. reinhardtii) samples were incubated in rumen fluid via 100 mL standardized glass syringes to measure in vitro gas production. Methane and gas volumes of feed samples were measured 24 h after incubation.

The mixed model of SAS version 9.4 (SAS Institute, Inc., Cary, NC, United States) was used in a completely randomized design to examine the data gathered from the in vitro study. The following model was used to statistically analyze the experiment:

where μ is the overall mean for each parameter, T_i is the effect of treatment, and E_ij is residual error. Differences among sample means with p < 0.05 were accepted as statistically significant.

The fundamental method for examining crystal size, phase purity, and crystal structure is X-ray diffraction (XRD) examination. As shown in Figure 1, the XRD pattern of the synthesized MgO exhibits various peaks corresponding to the (111), (200), (220), (311), and (222) reflection planes.

The particle size of the synthesized MgO NPs was determined from the Debye–Scherrer equation: D = K/cos (θ).

The MgO NPs’ median dimension and d-spacing values have been determined to be 20 nm and 0.25 nm, correspondingly.

Figure 2 shows the XRD pattern of CNPs. The XRD pattern exhibited an intense peak at 2θ = 22.90° and a weak peak at 2θ = 41.60°, corresponding to the (022) and (101) diffraction patterns of graphite carbon, respectively.

MgO NPs are characterized using the FTIR spectrum (Figure 3). In ambient settings, the spectra were captured at wavelengths ranging from 400 to 4,000 cm⁻¹. The peak at 651.94 cm⁻¹ indicates the stretching peak vibration of Mg-O bond, confirming that the obtained product is magnesium oxide. Moreover, H₂O adsorption on the metal surface is indicated by the peaks at 1552.0 cm⁻¹ and 3520.0 cm⁻¹.

The potential biomolecules in charge of the reduction of MgS NPs by green synthesis were found using Fourier transform infrared spectroscopy (FTIR) analysis. The FTIR spectra of MgS NPs made using Na₂S and pomegranate algae extract are displayed in Figure 4. In the spectrum, bands were observed at 3603.5, 1,725, 1,550, 1,232, 972 and 613 cm⁻¹. Particularly, the sharp band at 1,725 cm⁻¹ represents the C=O vibrations specific to the structure of flavonoids that can be found in pomegranate extract.

The characterisation of MgO NP production using pomegranate extract is depicted in Figure 5A, which is an image captured using a transmission electron microscope (TEM). Here, the scale bars are 500 and 200 nm. The images of TEM analysis of MgS NPs were taken and show the structures of MgS NPs (Figure 5B). The shape of these NPs was a small layer formation with a nearly spherical arrangement on a smooth surface. They had diameters ranging from 20–60 ± 1.6 nm and an average diameter of 55 ± 3.8 nm.

The nutrient composition and relative feed value of algal at various concentrations are indicated in Table 1. The extracts impacted significantly the chemical compositions (all p < 0.0001). Regarding fiber fractions such as ADF and NDF, the highest values were recorded for Algae +100 MgS. In contrast, in related to the CP and EE fractions, Algae +50 Mgs had the highest values (p < 0.0001).

The effects of algae extracts on in vitro rumen fermentation profiles are shown in Table 2. The parameters of gas production, pH, and OMD have influenced significantly by the different extracts (all p < 0.0001); the highest and lowest values regarding net gas, pH, OMD, ME, and NE_l were observed for Algae +50 MgO and Algae +50 MgS, respectively. The converse mentioned trend was observed for Algae +50 MgO and Algae +50 MgS in related to CH₄ production (p < 0.0001). Totally, not only measured total gas volume but also most of the measured parameters from the Algae-based rumen fluid were significantly influenced by the different nanoparticles (p < 0.0001).

Table 2 shows the volatile fatty acid (VFA) composition of rumen fluid. The effects of extracts on total VFA (TVFA) were substantial (p < 0.001), with the highest value found in the “Algae +100 MgS” group (163.12 mM) and the lowest in the CON group (139.59 mM). For the individual VFA, extracts have resulted in fluctuated amounts between treatments, in which the treatments influenced the individual VFA significantly (p < 0.001).

The observed peaks demonstrate the cubic structure of MgO and assign it to the pure phase of periclase MgO. In the spectra of other phases, no additional peaks could be seen. It confirmed that the prepared MgO was crystallized and was free of impurities. In addition, the presented peaks exhibit higher intensity and narrower spectral widths, indicating the product is in good condition. The XRD graph obtained for the crystallographic analysis of synthesized MgS nanomaterials is given in Figure 6. The 2θ values for MgS NPs peak at 37.94° (200), 45.42 (220) and 58.71° (221) at 200, 210 and 222. The characteristic peaks of the XRD spectrum at 2θ = 45.45° can be indexed at (220). Literature-based findings are consistent with the results obtained (24).

This is defined as OH stretching and bending, respectively. The metal-oxygen frequencies for the respective metal oxides published in the literature and observed frequencies coincide reasonably well. Using this method, MgO NPs can be analyzed for their chemical composition and surface properties (25). The -C-H bending vibrations in the aromatic amine groups of the flavonoid structure are linked to the absorption band at 550 cm⁻¹. Additionally, the peak at 972 cm⁻¹ shows the existence of MgS NPs as well as the distinctive C-S bond structure peaks. Under the aliphatic chain structure, the observed 613.4 cm-1 peak is part of the –CH2 group. The pomegranate algae extract’s bioactive components were verified using FTIR spectrum (26). Using this analysis, it is possible to determine the biomolecules involved in the synthesis of MgS NP. Figure 7 displays the carbon quantum dots of NP according to FTIR spectra. The band at around 3,242 cm⁻¹ is indicative of OH stretching vibration, which may arise from either the hydroxyl groups found in C black NP or water absorption. The peak recorded at 1,652 cm⁻¹ was exclusively found in pure CB and was ascribed to the material’s C=C stretching vibration. Peaks at 2,040, 2,166, and 2,015 cm⁻¹ are ascribed to the nanocarbon structure’s carbonyl group and C–O stretching (27).

TEM analysis is a very critical methodology to describe the particle size distribution, average particle size and shape of NPs. The produced MgO nanoparticles are less than 10 nm in size and spherical, as confirmed by TEM examination, despite being aggregated (28). The current green synthesis method approach has enabled the use of a simple and low-cost reducing agent for single-phase MgS NPs. This approach offers an effective method to synthesize MgS NPs in a non-toxic manner (24). Transmission electron microscopy (TEM) evaluated the morphology of pure carbon NP samples. These NPs are the samples showing the highest level of modification and are shown in Figure 5C. The TEM image shows semi-spherical primary particles with an average size ranging from 15 to 65 nm. These primary particles were formed and held together by agglomeration, resulting in agglomerates. The results corroborate other published studies in the literature, and the conclusions are consistent with current literature (29).

The present results on the chemical composition of macroalgae were in line with previous reports (30, 31). In disagreement with our findings, a recent meta-analysis of 47 published papers containing a broad variety of macroalgae was conducted and demonstrated that the average content of CP, NDF, ADF, and organic matter (OM) was 734.2, 189.2, 321.3, and 208.5 g/kg DM, respectively (32). Additionally, to confirm our findings Min et al. (33) published the different levels of CP (7.8 to 38.1% DM), NDF (16.6 to 43.1% DM), ADF (6.6 to 13.1% DM), and EE (0.3 to 3.9% DM) across eight macroalgae species. It’s important to note that the chemical composition and bioactive content of macroalgae are impacted by their taxonomic classification (brown, green, or red), and vary across genera and species. Seasonal fluctuations may also impact their composition during the growing and harvesting periods (31, 34). All algae and nanoparticles tested in our study had acceptable chemical compositions, particularly as a protein source; however, they should be included in a TMR ration to determine their potential advantages.

The current study’s findings about NDF and ADF were congruent with those of Mahmood Ameen (35). Feeds typically comprise 100–120 g/kg DM of ash. The crude ash levels of the feeds were comparable to those reported by Kamalak et al. (36) and Karabulut et al. (37). Differences in nutrient composition of feeds between studies may be qualified to numerous elements, such as climate, fertilization, species and type, harvesting time, feed storage conditions, and vegetative phase (36, 38). Also, it has been stated that the in vitro gas production level is affected by the nutrient composition of feedstuff, the presence of compounds inhibiting (such as tannins) gas production, the microflora and microfauna content of the rumen fluid (donor animal’s diet), and the quality of fermentation provided (36, 38).

Microalgae have mass balances ranging from 630 to 1,170 g kg⁻¹, although proximate analysis seldom provides 100% (39). Our investigation’s findings regarding the mass balance deficit suggest that other soluble components such as B vitamins, nonprotein nitrogen, chlorophyll, and soluble carbohydrates may be responsible. Microalgae fiber is low in hemicellulose and lacks lignin, even though it has a high fiber content (50–55% of total carbohydrate) (40). This enhances the probability that the protein will be readily available due to its lack of lignin complexation. In addition, the cell wall fraction in microalgae is highly digestible (41). Drewery et al. (42) found that supplementing post-extraction algal residue (CP = 179 g kg⁻¹ DM) increased OM digestibility in steers fed oat straw (CP; 45 g kg⁻¹ DM). Similarly, Tetracystis sp., N. bacillaris, and C. vulgaris have a higher lipid content, which improves the calorie density of the diet. It has been widely shown that lipids frequently diminish enteric CH₄ emissions from ruminants (43, 44).

The post-fermentation pH ranged from 6.75 to 7.79 among algal-extract treatments, demonstrating that algae supplementation promotes a more alkaline environment during microbial fermentation. Carbohydrates are the primary source of substrate for the creation of acetate and butyrate during ruminal fermentation, and as byproducts, CO₂ and hydrogen (H₂), are used by methanogenic archaea to produce CH4 (9). Furthermore, according to Kholif et al. (45), microalgae promote carbohydrate fermentation by rumen microbes, which is consistent with what was observed with the addition of microalgae and was attributed to the microalgae’s fulvic acids, which can provide carbon to ruminal microorganisms (46) and thus favor microbial growth and increase DMD. In turn, the increased degradability resulted in higher production of SCFA and ME, ascribed to enhanced carbohydrate degradation (45).

Although not investigated in the current study, the increase in SFCA and ME with microalgae might be due to increased activity of the fibrolytic bacteria (47) and increased propionate production. In contrast, decrease of SFCA and ME are attributed to a reduction in other SCFAs, such as acetate (48). In the meantime, the effects on DMD and SCFA associated with the content and degradability of feed carbohydrates may be reflected in the computed variations in CH₄ per unit of SCFA, ME, and OM (49).

Biogas production (BG) is intimately related to feed degradability and, as a result, the availability of highly-fermented nutrients for rumen microbial activity and growth (15). Although their production is predominantly reliant on the fermentation of carbohydrates to SCFA and proteins, and BG is mostly made up of CO₂ and CH₄, their contribution to BG is negligible in comparison to that of carbohydrates (50). Furthermore, the production of acetate and butyrate during rumen fermentation produces more gas than the formation of propionate, accounting for the majority of the BG (51).

Natural compounds of microalgae have been proposed as potential methods for controlling rumen fermentation, contributing to CH₄ generation (52, 53). A previous in vitro investigation (54) demonstrated that Schizochytrium spp. inhibited CH₄. Furthermore, several research (55, 56) found an increase in CH₄-producing bacteria and protozoa, demonstrating that not all microalgae have CH₄-reducing properties.

The anti-methanogenic effect observed in this study has been reported in studies involving other microalgae (Spirulina platensis, Chlorella vulgaris, and Schizochytrium spp.). The studies attribute this effect to the presence of docosahexaenoic acid (C22:6 n − 3) and eicosapentaenoic acid (C20:5 n − 3), polyunsaturated acids that decrease the concentration of acetate and increase propionate, which results in reduction the abundance of methanogenic archaea, the primary microorganisms producing CH₄ (15, 51, 52). Likewise, Sheng et al. (57) found that humic compounds, including fulvic and humic acids, can lower CH₄ production in ruminants. They ascribed this to a decrease of the molar proportion of protozoa and acetate populations (58), which minimizes the amount of H₂ available for CH₄ production (59).

The addition of the microalgae reduced BG production in the current study, which is in line with Elghandour et al. (15), who observed that the BG decreased with the addition of the microalgae Schizochytrium spp. and associated it with the antimicrobial and cytotoxic effects of the compounds of the microalgae (60), as well as the long-chain fatty acid profile (48). Also, it is likely that the microalgae have modified the structure of the microbial community during fermentation which is caused variations in the final fermentation products, including the SCFA profile (61).

Among the treatments, the highest amount of gas produced was observed for the MgS nanoparticles group. Additionally, the MgO treatments demonstrated a notable decrease in the production of methane, which indicate the ability of MgO nanoparticles to meet the needs of rumen bacteria during the incubation period (62). The two main sources of in vitro gas generation are carbon dioxide and methane, which are derived directly from microbial fermentation, and carbon dioxide released from a bicarbonate buffer, obtained indirectly by buffering short-chain fatty acids. Menke and Steingass (23) affirm that the only variables influencing gas generation are the feed’s physical and chemical composition. The fermentation rate, however, could be impacted by modifications in ruminal microbial activity.

Volatile fatty acids (VFAs) have been considered one of the most significant factors in achieving anaerobic fermentation. According to Makkar (63), fluctuations in gas production might alter the amounts or ratios of VFA produced. VFAs’ hydrophobic qualities enable them to penetrate the bilayer structure of the bacterial cell’s plasma membrane (64). Therefore, by changing the membrane structure and increasing its flowability and permeability, they can lower the rate of bacterial growth (65).

Previous research has shown that adding red algae (Asparagopsis taxiformis) and lipid-extracted microalgae to forage diets dramatically boosted propionate and butyrate levels in the rumen (66). This is deemed advantageous since previous research demonstrated that the energy from propionate was used more efficiently than energy from acetate (67, 68). Lodge-Ivey et al. (68) found that adding lipid-extracted algae (Chlorella or Nannochloropsis) to the diet increased total rumen VFA content, which is in consistent with our findings. In contrast to our findings, it has been proposed that the high lipid content of Chlorella may suppress cellulolytic bacteria in the rumen and finally reduction of total VFA (53). Furthermore, adding algae to a corn silage-based diet raised ruminal pH and reduced total VFA by up to 18% after 19 days (69). Other in vitro investigations (53–55, 70) found that supplementation with DHA-rich microalgae or marine algae increased ruminal propionate while decreasing overall VFA and CH₄ synthesis. The differences could be attributed to variances in supplementation levels and oil extraction.