S. latissima was collected from Trondheimsfjord (N63˚26'56'', E10˚10'48'') near Trondheim, Norway in August of 2017. The macroalgae were subsequently washed using tap water to remove particulates from the surface. S. latissima was then milled using a table-top blender with 10 mL of deionized water per 1 kg of macroalgae to produce a dense macroalgae pulp. The pulp was dried for 48 h at 30˚C and then stored in airtight plastic bags in a dry location for further use.

The dry macroalgae pulp (10% w/v) was suspended in 0.15 M sodium carbonate (Na₂CO₃) solution to a volume of 1 L at a starting pH of 9.0 in a stirred beaker for 2 h at 50˚C.

Commercial β-glucanase (G4423) from Trichoderma longibrachiatum (Sigma Aldrich, Germany) was used during enzymatic hydrolysis. This was an enzymatic mixture of β-1-3/1-4-glucanase, xylanase, cellulase, β-glucosidase, β-xylosidase, α-˪-arabinofuranosidase and amylase activities. The lysed algal pulp was adjusted to pH 6 using HCl acid solution. Then 5 mg of enzyme mix per g of macroalgae dry weight was added to the solution, and left in the stirred flask for 48 h at room temperature. Samples were taken every 12 h and analyzed for glucose concentration using a hexokinase glucose assay kit (GAHK20-1KT, Sigma, Germany).

Total carbohydrates, reducing sugar and glucose content were both determined by acidic treatment of pre-hydrolysis dry biomass. Biomass (0.5 g) was treated with 5 mL of 72% (v/v) H₂SO₄ at room temperature for 30 minutes with constant stirring via a magnetic stirrer. The sample was then diluted to a volume of 50 mL with deionized water, then autoclaved at 121˚C for 30 minutes. Once cooled, NaOH was added to the sample to reach a pH of 7.5. The total carbohydrates in the sample were then determined by using a phenol-sulfuric acid method [18] . Reducing sugars in the sample were determined using a dinitrosalicylic acid method [19] . Glucose content of the hydrolysate was determined using a hexokinase glucose assay kit (GAHK20-1KT, Sigma, Germany).

The microbial inoculum for the biomethane potential experiments was obtained from the Biokraft biogas plant (Skogn, Norway), from a large-scale continuous mesophilic multifuel anaerobic digester. Prior to the biomethane potential test, the inoculum was incubated at 39˚C to reduce endogenous biogas production. The inoculum was diluted to 9 g volatile solids (VS) per liter. The diluted inoculum had a pH of 7.5.

The processed samples were digested anaerobically in sealed batch bottles. These were performed in triplicate with inoculum and processed samples, with inoculum alone as a control batch. The following was added as trace minerals and nutrients: 10 mM NH₄Cl, 8 mM NaCl, 1 mM CaCl₂・2H₂O, 0.5 mM MgCl₂・6H₂O and 0.3 mM Na₂S・3H₂O, as previously described [20] . The final liquid volume of each experiment was 250 mL. The bottles were flushed with nitrogen gas and closed with rubber septums and sealed as described by Ekstrand et al. [21] . These were incubated at 39˚C indefinitely with regular agitation.

Analysis of the dry matter (DM) and volatile solids (VS) we performed on the steam exploded birch wood and inoculum before further experimentation using standard methods [22] . The dry matter was determined by incubation of the birch wood and inoculum at 105˚C for 24 h. The mass of the samples after the incubation was compared to the mass pre incubation to acquire a percentage of DM. New samples were then incubated at 550˚C for 4 hours in a muffle furnace. The weight of the samples after incubation in the muffle furnace were compared to the weight of the sample before incubation to acquire a percentage of VS. The DM was determined to be 14.63%; whereas, the VS was determined to be 9.13%.

Measurement of the produced gas volume was achieved using an optimised liquid displacement method. The volume of gas was then adjusted for temperature using Charles’s law.

Regular 10 mL samples of gas from each experiment were extracted for compositional analysis. The composition was analysed using gas chromatography (SRI 8610C, SRI Instruments, USA), equipped with a thermal conductivity detector using hydrogen as the carrier gas. A standard mixture of CO₂, CH₄, H₂ and N₂ was used as a calibrating gas. Using the volume of gas calculated with the liquid displacement method, and the percentage of methane present in the samples, methane production was calculated for the experiments. All methane production levels are the average of 3 separate experiments and have been corrected by subtracting the endogenous methane production levels for the inoculum only experiments.

All experiments within this study were conducted in triplicate with the results displayed as mean values ± the standard deviation.

Using the collected samples of S. latissima, the total carbohydrates, reducing sugars and the glucose yield were determined (Table 2). These observations are similar to those previously obtained using the same methodology [17] . Before the enzymatic hydrolysis, the total carbohydrates consisted of 55% ± 3.1% of the dry weight of S. latissima. Furthermore, the reducing sugar content was 36% ± 1.9% of the dry weight, whereas the glucose content was 10% ± 1.8% of the dry weight of S. latissima. These observations of carbohydrate composition are in agreement with previously observed compositions of S. latissima, including studies of S. latissima obtained from Trondheimsfjorden [17] [23] [24] [25] .

During a 2 h basic (pH 9) lysis method at 50˚C in a 0.15 M Na₂CO₃ solution, the polysaccharides from S. latissima were extracted. The lysate was then altered to a neutral pH (pH 6) and exposed to an enzyme mixture of β-1-3/1-4-glucanase, xylanase, cellulase, β-glucosidase, β-xylosidase, α-˪-arabinofuranosidase, and amylases for a period of 48 h at room temperature. The reducing sugar concentration of the sample was measured every 12 h during the enzymatic hydrolysis process.

From the initial basic lysis of S. latissima, 8.37 ± 0.53 g/L of reducing sugars were released. Further enzymatic hydrolysis yielded a further 21.74 ± 1.77 g/L of reducing sugars after the 48 h incubation period (Figure 1; Table 3). The total reducing sugars obtained from the basic lysis and enzymatic hydrolysis was observed to be 30.11 ± 2.30 g/L of reducing sugars. When using the dinitrosalicylic acid method for determining total reduced sugars [19] , the maximum amount of reducing sugars obtainable from the sample of S. latissima was observed to be 36% ± 1.9% of the dry weight of reducing sugars before the lysis and enzymatic hydrolysis. Therefore, the efficiency of this process was determined to be 83.6% of the maximum obtainable reducing sugars, which is in agreement with previously observed results [17] .

With regard to the rate of the enzymatic hydrolysation of the polysaccharides from S. latissima, the maximum rate was observed at 12 h of incubation with the enzyme mixture (Figure 2; Table 3). The rate gradually declined after this period, as typically observed in similar studies [12] [17] [26] [27] . Although it has been speculated that the decline in rate of the enzymatic hydrolysis may be due to the inhibition of the enzymes by the liberated glucose and cellobiose [12] [27] [28] , it may also be due to the reduction of functional enzyme units in relation to the reduced amount of complex polysaccharides left.

Anaerobic Digestion of the Hydrolysate from S. latissima

The anaerobic digestion of the hydrolysate from S. latissima was performed using inoculum from a biogas plant near Trondheim, Biokraft (Skogn, Norway). The sample was added in a 2:1 (VS/VS) ratio with the inoculum in 500 mL flasks sealed with rubber septums and flushed with nitrogen gas for 5 minutes to obtain an anaerobic environment. The bottles were then incubated at 39˚C indefinately, with regular mixing. The biogas production was measured regularly using a liquid displacement method, and gas samples were assessed for composition using a GC machine (SRI 8610C, SRI Instruments, USA).

Two experiments were prepared for anaerobic digestion to determine the effect of the enzymatic hydrolysis on the amount of biogas produced; one with the lysed algal pulp without the enzyme hydrolysis step, and one with the lysed algal pulp with the enzyme hydrolysis step. The total biogas obtained from the anaerobic digestion of S. latissima algal pulp without enzyme hydrolysis was 372 ± 22 mL per gVS. This is a similar biogas volume to previously observed values [20] [29] [30] , but variation is expected due to seasonal and locational differences. Comparatively, the total biogas obtained from the anaerobic digestion of S. latissima S. latissima algal pulp with enzyme hydrolysis was 459 ± 30 mL per gVS (Figure 3; Table 4). The gas contained 56% CH₄ as determined by GC, resulting in a biomethane production of 208 ± 12 and 257 ± 17 mL per gVS for the algal pulp without and with enzyme hydrolysis, respectively (Figure 3; Table 4). Furthermore, the enzyme hydrolysis step was observed to increase the rate at which biogas was produced, reaching a near full digestion earlier than the algal pulp without the enzyme hydrolysis step.

The results observed in this study of the anaerobic digestion of S. latissima show that the extraction of reducing sugars from 1 kg (w/w) of the macroalgae yields 30.11 ± 2.30 g of reducing sugars. The subsequent enzymatic hydrolysis allows the straight-forward anaerobic digestion of these carbohydrates into 459 ± 30 mL per gVS of biogas, resulting in 257 ± 17 mL per gVS of biomethane from S. latissima. With respect to area required, as shown in Table 1, the amount of wet macroalgal biomass obtainable from 1 ha of sea floor is as high as 75,000 kg [31] . The VS of S. latissima was determined to be 9.13%, resulting in 6847.5 kgVS per ha. Therefore, 1 ha of S. latissima could potentially yield 3142 m³ of biogas, composed of 1760 m³ of biomethane. Therefore, the carbohydrate-rich macroalgae S. latissima has a large potential for utilisation as a substrate for biogas production. Furthermore, the utilisation of fresh water sources, fertilisers and terrestrial land is not required due to their ability to grow wildly in the Ocean.

The advantages of CO₂ sequestration associated with the cultivation of macroalgae are also significant. Typically, the utilisation of 1 tonne of macroalgae sequesters 6 times the amount of CO₂ emitted during the processing, maintenance and transport of the feedstock [32] , and the close proximity of macroalgae farms to sources of pollution can improve their growth rate, while cleaning the water of excess nutrients [30] . Macroalgae also have a high level of marco nutrients, improving the digestate as a biofertilizer [30] [33] . Finally, the C:N ratio desired for optimal anaerobic digestion is between 20:1 - 30:1 [34] , where a ratio bellow 15:1 can result in excess levels of ammonia in the anaerobic digestion process, leading to unstable biogas production [35] . The C:N ratio of S. latissima has been observed to be 22:1, suggesting that it is an ideal substrate for anaerobic digestion [11] .

Despite this, it should be noted that the impact of the extensive harvesting of wild macroalgae in these regions is unknown and may cause local ecological issues. The turnover of macroalgae in Nordic regions is relatively high, but excessive removal of macroalgae from the seabed in a specific area could result in the collapse of the community, which may take several years to recover. Alternative methods of industrial-scale growth could avoid these drawbacks, by the growth of macroalgae on suspended systems in deeper waters. If the waters are deep enough, there would be a limited amount of wild macroalgae present in the area; however, microalgae would then be competing for light with these artificial systems, and their growth could be reduced.