# Development of a scalable design for biofuel production from green macroalgae hydrolysates

Hayet Djelal, Alaa Salma, Walaa Sayed, Audrey Cabrol, Maud Benoit, Amance Corat, Abdeltif Amrane, Ronan Pierre

► To cite this version:

Hayet Djelal, Alaa Salma, Walaa Sayed, Audrey Cabrol, Maud Benoit, et al. Development of a scalable design for biofuel production from green macroalgae hydrolysates. Algal Research - Biomass, Biofuels and Bioproducts, 2026, 94, pp.104487. 10.1016/j.algal.2025.104487. hal-05445881

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# Development of a scalable design for biofuel production from green macroalgae hydrolysates

Hayet Djelal<sup>a,*</sup>, Alaa Salma<sup>a</sup>, Walaa Sayed<sup>a,b</sup>, Audrey Cabrol<sup>b</sup>, Maud Benoit<sup>c</sup>, Amance Corat<sup>c</sup>, Abdeltif Amrane<sup>b</sup>, Ronan Pierre<sup>c</sup>

## ARTICLE INFO

### Keywords:

Green macroalgae

Fermentation

Third generation bioethanol

Lab-scale

Pilot-scale

## ABSTRACT

This study investigated the valorization of green macroalgae in ethanol by the yeast *Saccharomyces cerevisiae*. The proof of concept was conducted with two green macroalgae hydrolysates, namely *Ulva rigida* and *Chaetomorpha linum* and the fermentation process was scaled-up from 250 mL and 500 mL shake flasks to a 3 L stirred-tank bioreactor. Firstly, the experiments used a synthetic medium based on algal hydrolysate, examining factors like nitrogen source, inoculum size, and salt content. After 72 h, ethanol yields were similar across the systems: 0.43 g.g⁻¹ in the bioreactor, compared to 0.46 and 0.45 g.g⁻¹ in 250 mL and 500 mL flasks, respectively. Secondly, fermenting green algae hydrolysate was tested, under similar conditions (without aeration and pH adjustment, with nitrogen enrichment, and 1 % v/v inoculum) resulted in an ethanol concentration of 9.05 g.L⁻¹, a yield of 0.44 g.g⁻¹, and 85.86 % efficiency. Thirdly, operating the process at larger scale was carried out on pilot scale of 77 L with *Chaetomorpha linum* hydrolysate. Glucose was fully consumed in 9 h, and maximum ethanol production (5.61 g.L⁻¹) occurred after 11 h, with a yield of 0.52 g.g⁻¹ which corresponds to the theoretical yield.

## 1. Introduction

Fluctuations in oil prices, the ongoing depletion of non-renewable energy sources, and increasing greenhouse gas emissions have spurred global efforts to find sustainable and environmentally friendly alternatives to fossil fuels [1,2]. Among these alternatives, bioethanol is considered as a promising alternative energy resource [3,4]. Compared to fossil energies, ethanol is a clean and efficient energy and widely accepted as a potential substitute for fossil fuels [3–5]. In this context, the bioconversion of biomass to ethanol has received great importance. Bioethanol can be produced from various biomass feedstocks, especially from sugar cane [6,7], corn starch [8] and other lignocellulosic biomass [9]. Even though these biomasses are economically competitive with fossil fuels, they use agricultural resources and so influence the food chain by increasing the food price [10]. Thus, seaweeds have emerged as a highly promising biomass material for ethanol production [11,12], offering several advantages over first- and second-generation biomass [13,14]. While seaweeds are primarily used for biodiesel production due

to their high lipid content, their cellulosic structure and abundant carbohydrates also make them suitable for bioethanol production [15]. Additionally, seaweed has the advantage of easy depolymerization, as they do not contain lignin in their cell walls, unlike most terrestrial plants [16]. Their high carbohydrate content can be readily converted into simple sugars, which can then be directly fermented into bioethanol [17].

Among seaweeds, *Ulva* is a cosmopolitan genus of green seaweed that can be found from tropical to polar climates, from fresh to fully saline environments [18]. Although *Ulva* causes the green tides nuisance, it is also used in the food and feed industries [19]. Seasonal and environmental variations affect the biochemical composition of the macroalgae, including their cellulosic content, which is the principal source of carbohydrates [20].

Green algae also contain salts, such as sodium chloride, and sulfates from sulfated polymer like ulvan [21]. These components could affect the osmotic pressure of the culture medium and, consequently, the performance of ethanol production [22]. Many studies explored the

<footnote><sup>a</sup> UniLaSalle, ECLORE, ULR 7519, Campus de Ker Lann, 35 170, Bruz, France</footnote>

<footnote><sup>b</sup> Univ Rennes, Ecole Nationale Supérieure de Chimie de Rennes, CNRS, ISCR, UMR6226, F-3500, Rennes, France</footnote>

<footnote><sup>c</sup> CEVA, Presqu'île de Pen Lan, 22610, Pleubian, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>*E-mail addresses:* hayet.djelal@unilasalle.fr (H. Djelal), elaasalma93@gmail.com (A. Salma), walaa_91@hotmail.com (W. Sayed), Aud.cabrol@gmail.com (A. Cabrol), maud.benoit@ceva.fr (M. Benoit), amance.corat@ceva.fr (A. Corat), abdeltif.Amrane@uni-rennes.fr (A. Amrane), ronan.pierre@ceva.fr (R. Pierre).</footnote>
potential of green algae as a nutrients source for bioethanol production through fermentation process [23–25]. A worldwide interest in the utilization of bioethanol as energy source has driven research on the cost-effectiveness of ethanol production industrial processes [26]. Intensive research has been therefore conducted to develop efficient fermentative organisms, affordable fermentation substrates and optimum environmental conditions for fermentation [27].

Several microorganisms including bacteria, yeasts, and fungi have been used for ethanol production. However, some microorganisms have greater potential to accumulate high concentrations of ethanol [27]. *Saccharomyces cerevisiae* has historically been the most widely used microorganism for this purpose [28]. The biological production of ethanol is a complex process influenced by several fermentation variables that can affect microbial metabolism. Key factors that need to be controlled include substrate and product concentrations, the accumulation of inhibitory substance, aeration, agitation, temperature and pH [29].

Scaling-up a process allows unravel issues that were not significant at laboratory scale. To ensure that the fermentation yield is maintained, and to achieve total control over culture conditions; the critical points for the scale-up methodology are process stability and parametric sensitivity [30]. Increasing production capacity to commercial scale can be approached with confidence only when the process has demonstrated robustness at pilot scale [31]. The development of industrial bio- processes requires several key steps: screening microbial strains, opti- mizing conditions, understanding the regulatory network, and integrating environmental signals by cells [32]. Data from pilot-scale is used to estimate the life cycle profiles at full scale and, consequently, to estimate the environmental implications of deploying algae-to-energy systems [33].

This study explores the potential of using green macroalgae, partic- ularly *Ulva rigida* and *Chaetomorpha linum*, as promising biomass sources for bioethanol production. The main interest of is the scale up of the fermentation process from lab-scale experiments, conducted in shake flasks, to a pilot-scale bioreactor. Evaluation of the impact of various factors such as aeration, pH control, nitrogen source, inoculum size, and medium composition on the fermentation performance was conducted. Thus, this study trends to study the feasibility of the bioproduction of third generation bioethanol on pilot scale after the proof of concept was checked on lab-scale.

## 2. Materials and methods

### 2.1. Strain maintenance and inoculum preparation

*Saccharomyces cerevisiae* baker's yeast CLIB 95 (CIRM French) was used in this study. Before inoculation of the fermentation medium, the yeast strain was maintained in a pre-culture medium whose composition consisted in (g.L⁻¹): glucose (20), peptone (10), yeast extract (10), and agar (20). The culture was maintained on Petri plates and agar slants at 28 °C during 24 h and then stored at 4 °C.

The cells from a single well-isolated colony were then inoculated into 250 mL Erlenmeyer flasks containing 25 mL of sterilized culture me- dium, whose composition was the same as MY medium, without agar. The cultures were incubated on a rotating shaker (New Brunswick, Innova 40) at 180 rpm and 28 °C, during 18 h to obtain high cell density. Then, the cells were recovered aseptically by centrifugation (3000 rpm, 4 °C and 5 min), re-suspended in 25 mL KCl (150 mmol.L⁻¹) and centrifuged again in similar conditions. The suspension obtained after harvesting cells and re-suspending in 10 mL of KCl was used as inoculum for the experiments [34].

### 2.2. Algal hydrolysate

The hydrolysis of *Ulva rigida* biomass (20.8 % dry matter) whose origin had described in previous work [35], was conducted in a 120 L

pilot. The seaweed was harvested, frozen, thawed, and milled. A 10 % dry matter suspension was prepared by adding water, heated to 60 °C, and the pH adjusted to 5.5 using HCl (1 mol.L⁻¹). Gelatinization was achieved by heating the biomass to 100 °C for 30 min while stirring. After cooling, *Liquozyme®* rich in α-amylase was added at 2.9 % of starch dry weight (DW), and the mixture was heated to 85 °C for 2 h for liquefaction. *Spirizyme®* rich in glucoamylase, was then added at 11.6 % of starch dry weight, and the mixture was heated to 65 °C for 2 h for saccharification. Enzymes were deactivated by heating to 100 °C for 30 min, then cooling to 70 °C. The pH was adjusted to 0.88 with sulfuric acid, and the mixture was heated at 70 °C for 6 h to hydrolyze co- extracted ulvans. The mixture was cooled but maintained at 40 °C to avoid excessive viscosity and the pH adjusted to 5 with Na₂CO₃. The hydrolysate was filtered through an 11 µm sieve (Sweco Separator), then autoclaved at 120 °C and 1 bar for 20 min (SMI, Montpellier, France).

For the scale up study, hydrolysate of *Chaetomorpha linum* was used. This alga, rich in cellulose, was cultivated under stress in the French Centre for the Study and use of Algae (Centre d'études et de valorisation des algues, CEVA, Pleubian, Brittany-France), to also trigger starch accumulation and to increase glucose content. Hydrolysates were pre- pared by CEVA using a cascade of enzymatic hydrolyses (Ctec3 rich in cellulase/ *Liquozyme®* / *Spirizyme®*) after physico-chemical treatment as previously described.

### 2.3. Synthetic media

To optimize culture and ethanol production conditions, assays were conducted on synthetic media similar to macroalgae hydrolysate. Syn- thetic media prepared following the composition of green macroalgae *Ulva* sp., was composed of simple sugars (glucose, galactose, xylose, rhamnose and arabinose) at levels close to those of hydrolysates (12 g L⁻¹). Mineral supplementation was added in culture media to enhance the growth of the yeast *S. cerevisiae*. The composition was well described in our previous work [22]. To sterilize the medium without modification of the composition, medium was filtered through a 0.2 µm pore size filter (Sartorius) under aseptic conditions.

### 2.4. Fermentation reactors

Small-scale production, 250 mL and 500 mL flasks (working volume of 100 mL and 200 mL respectively) were used for ethanol production. After inoculation, fermentation medium was incubated on an orbital shaker (INNOVA 40).

For the stirred-tank bioreactor production, ethanol fermentation was carried out in a 3 L stirred-tank bioreactor (Bio Flo / CelliGen 115) of 1.5 L working volume., The aeration flow rate was controlled manually by two Rotameters. It is possible to cascade dissolved oxygen (DO) to Agitation (AGIT); consequently, the agitation speed can be varied be- tween the user-specified minimum and maximum set points to maintain the set percentage of dissolved oxygen.

For the pilot scale, the fermentation was conducted in an agitator bioreactor with total and working volume of 77 and 33 L respectively.

### 2.5. Fermentation tests

Algal hydrolysate of green macroalgae *Ulva rigida* contains salts which could influence osmotic pressure and consequently the meta- bolism of yeast. The synthetic medium was therefore supplemented with sodium chloride and sulfate at similar concentrations found in *Ulva rigida* hydrolysate (0.25 and 0.21 mol.L⁻¹ respectively). Two other pa- rameters, nitrogen source, (NH₄Cl, 1 g.L⁻¹ or peptone, 5 g.L⁻¹), and the size of inoculum (0.1 and 1 % v/v), were also investigated to maximize the production of ethanol. The tests were conducted in 500 mL agitated flasks at 28 °C and 180 rpm. The Effects of source of nitrogen, inoculum size and salts on fermentation after 48 h using *S. cerevisiae* growing on a synthetic medium in the following conditions: A: with NH₄Cl as nitrogen
source, 0.1 % v/v inoculum size and without salts, B: with peptone, 0.1 % v/v inoculum size and without salts, C: with peptone, 0.1 % v/v inoculum size and with salts, D: with peptone, 1 % v/v inoculum size and with salts.

Effect of pH and aeration on ethanol production, were conducted in 3 L agitated bioreactor with synthetic medium based on *Ulva rigida* hydrolysate' composition at 28 °C, 180 rpm with 1 % v/v of S. cerevisiae inoculum. When it is necessary, the pH was maintained at 6.0 by automatic addition of KOH (2 mol.L⁻¹) to prevent acidification.

The algal hydrolysate of green macroalgae *Ulva rigida* was subjected to fermentation without enrichment and for an inoculum size of 1 % (v/v). Batch cultures were carried out in 3 L bioreactor using S. cerevisiae. Temperature (28 °C) and agitation (180 rpm) were kept constant throughout the culture. Experiments were carried out without aeration and at a pH value adjusted at 6.0 but not controlled.

The scale up test was conducted in 77 L agitated bioreactor maintained at a constant temperature of 28 °C, with continuous agitation at 250 rpm, without external aeration. The bioreactor was filled with 35 kg dry matter of *Chaetomorpha linum* hydrolysate. This hydrolysate was used directly as the substrate, without enrichment or sterilization. *Saccharomyces cerevisiae* was introduced as the microbial strain at an inoculum size of 0.07 % (w/w) relative to the hydrolysate. The initial pH was adjusted to 6.0 but was not controlled during fermentation.

With the exception of the 77 L bioreactor test, all experiments were conducted in triplicate. Fig. 1 illustrates the scale-up process in bioreactor systems.

## 2.6. Analytical methods

Analysis of total sugars and uronic acids was performed using high-

performance liquid chromatography (HPLC) with a reverse-phase C18 column after depolymerization under methanol-acid hydrolysis reaction (methanolysis). Starch content was subsequently extrapolated from total glucose content [36].

The metabolites produced by the yeasts (ethanol, glycerol and acetic acid) and the sugar concentrations were measured using high performance liquid chromatography (HPLC), with an ions exclusion column HPX-87H (300 × 7.8 mm, Bio-Rad, Hercules, CA, USA), kept at 45 °C (Oven CrocoCil™; Cluzeau-Info-labo, Ste. Foy LaGrande, France). The eluent for separation was sulfuric acid (0.01 M), applied at a flow rate of 0.7 mL.min⁻¹. The column was coupled with the Shimadzu RIO-6 A Refractive Index Detector (Japan), used for the detection of the various compounds. The samples were withdrawn and centrifuged at 3000 rpm, 4 °C and 5 min before analysis.

Cell growth was monitored by analysis of absorbance at 600 nm, with a spectrophotometer SECOMAM Prim 500, after calibration using a non-inoculated medium. Biomass growth was also measured in terms of dry matter (g.L⁻¹). A volume of 30.10⁻³ L of medium was disposed in a previously weighed porcelain cup (P1) and placed in an oven at 105 degree C during 24 h. Dry medium was then weighed (P2) and the dry matter could be calculated as follows (Eq. 1):

$$Dry matter = \frac{(P2 - P1)}{30.10^{-3}} \qquad (1)$$

## 2.7. Fermentation parameters

During alcoholic fermentation, fermentable sugars are converted into ethanol and carbon dioxide by the action of microorganisms.

The yield of ethanol (YEtOH) to consumed sugar (g.g⁻¹) was defined as the ratio of the amount of ethanol produced to the amount of sugar

<figure><img src="image_4.png" /><figcaption>Fig. 1. Graphical of the scale-up process.</figcaption></figure>
consumed (S0-Sf, with S0 and Sf the initial and final sugar concentrations) (Eq. 2):

$$Y_{EtOH} = \frac{\text{Ethanol produced}}{\text{substrate consumed}} (\text{g.g}^{-1}) \qquad (2)$$

The ethanol productivity rate (g.L⁻¹.h⁻¹) was calculated as the ratio of the amount of ethanol produced (g.L⁻¹) during a given time (t, h) (Eq. 3):

$$\text{Ethanol productivity (g.L}^{-1}.h^{-1}) = \frac{[\text{ethanol}]}{\text{time}} \qquad (3)$$

Fermentation efficiency (%) corresponded to the ratio of the experimental yield of ethanol to the theoretical yield (Eq. 4):

$$\text{Efficiency (\%)} = \frac{\text{Practical yield of ethanol}}{\text{Theoretical yield}} \times 100 \qquad (4)$$

# 3. Results and discussion

## 3.1. Influence of salt and nitrogen

Synthetic media based on *U. rigida* hydrolysate composition was supplemented in nitrogen or salts. The results are summarized in Fig. 2 and Fig. 3. Assimilable nitrogen is an important component of culture media and it plays a significant role in the course of fermentation [37]. The structural complexity of the nitrogen source strongly affects both biomass and ethanol production by industrial strains of *S. cerevisiae* [38]. Fig. 2 (A, B) showed that nitrogen sources had significant effect on ethanol fermentation. The glucose consumption rate was 0.29 g.L⁻¹.h⁻¹ using NH₄Cl as nitrogen source, while this rate increased 1.7 times in the presence of peptone as nitrogen source. Ethanol volumetric productivity followed the same trend, 0.13 and 0.20 g.L⁻¹.h⁻¹ with NH₄Cl and peptone, respectively. In fact, with a better assimilation of nitrogen, cell growth is favored, leading to higher glucose consumption; the poor yeast performance observed using ammonium agreed with the related literature. The use of peptone led also to an increase of the cell growth (absorbance at 600 nm), 11.3 instead of 4.2 in the presence of ammonium (data not shown). In general, supplementation with a complex nitrogen source, such as peptone, is more positive for yeast metabolism, inducing higher biomass and ethanol production, and preserving yeast viability [38].

Fig. 2 (B, C) showed the impact of salt found in green macroalgal hydrolysate. Glucose consumption rate (20 g.L⁻¹ initially) was not significantly affected by the osmotic pressure, since irrespective of the presence of salts, all the glucose was consumed at the end of culture. But

<figure><img src="image_5.png" /><figcaption>Fig. 3. Biomass (C), ethanol production (Δ) and glycerol production (ΔΔ), biomass growth during fermentation by *S. cerevisiae* without aeration (Red color), and with aeration (Blue color) (Vt = 3 L). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

a rather weak decrease in the performance of *S. cerevisiae* was observed in the medium enriched with several salts if compared to the synthetic medium without salt. Ethanol production decreased from 9.71 to 8.34 g. L⁻¹, and ethanol volumetric productivity decreased from 0.20 to 0.17 g. L⁻¹.h⁻¹. Results (Fig. 2 (C, D)) show that increasing the inoculum size increased the ethanol production, 9.63 and 8.34 g.L⁻¹ for 1 % and 0.1 % inoculation. Ethanol production rose from 0.17 to 0.2 g.L⁻¹.h⁻¹ when the yeast load increased from 0.1 to 1 % v/v. In both cases, glucose was totally consumed leading to the same glucose consumption rate (0.5 g. L⁻¹.h⁻¹). In addition, ethanol yield and efficiency were also found to be optimal for 1 % v/v inoculum (0.40 g.g⁻¹ and 78.52 % - Fig. 3). High ethanol productivity is an economically relevant factor for industrial purposes and hence 1 % v/v inoculation level was selected and considered thereafter. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

The rate of ethanol production and glucose consumption were faster using peptone as nitrogen source, while the efficiency and the ethanol yield were not improved with peptone if compared to the use of NH₄Cl (Fig. 3). For *S. cerevisiae*, a mineral source allows to obtain a high ethanol yield with a low biomass yield; while an organic source, such as peptone, allows improving ethanol volumetric productivity despite a loss of carbon substrate for biomass formation. Using peptone as a source of nitrogen allowed rapid fermentation (high rate of production and consumption) by minimizing fermentation time, which makes the process more effective from an economic point of view.

Due to the presence of salts, yeasts need to adapt to a higher osmotic pressure and hence the maintenance energy is increased; it is multiplied by 4 when 1 M NaCl is added to the culture medium [39]. Accordingly, the efficiency of conversion of sugar into ethanol decreased from 79.2 to 68.0 % in the presence of salts (Fig. 3). In the presence of salts, glycerol and acetic acid yields rose (data not shown); salts impact ethanol production and promote glycerol production [40], in order to adapt to the osmotic pressure of the culture medium. The inoculum size is known to have an impact on the process; a very small inoculum concentration increases the fermentation time and a too high concentration can lead to a loss in ethanol yield due to sugar consumption by the multiplying yeast cells [41]. To maximize the ethanol production from the synthetic model medium of hydrolysate of macroalgae using *S. cerevisiae*, inoculum size was increased from 0.1 to 1 % v/v.

<figure><img src="image_6.png" /><figcaption>Fig. 2. pH (ΔΔ), ethanol production (ΔΔ) and biomass growth (C) during fermentation by *S. cerevisiae* with controlled pH at 6 (Red color), and without pH control (Green color) (Vt = 3 L). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

## 3.2. Up-scaling of the fermentation process on synthetic medium

Synthetic media based on *U. rigida* hydrolysate composition were used for this part of the study. The size of the inoculum was 1 % v/v
<caption>**Table 1**</caption>

Kinetic parameters of fermentation at different scales after 72 h of fermentation on synthetic medium based on *U. rigida* hydrolysate composition.

<table><thead><tr><th>Description</th><th>Flask<br>Fermentation<br>(250 mL)</th><th>Flask<br>Fermentation<br>(500 mL)</th><th>Bioreactor<br>Fermentation (3 L)</th></tr></thead><tbody><tr><td>Absorbance (600 nm)</td><td>2.60</td><td>3.10</td><td>3.50</td></tr><tr><td>Ethanol (g.L<sup>-1</sup>)</td><td>6.87</td><td>6.71</td><td>8.41</td></tr><tr><td>Ethanol volumetric<br>productivity (g.L<sup>-1</sup>.<br>h<sup>-1</sup>)</td><td>0.10</td><td>0.09</td><td>0.12</td></tr><tr><td>Ethanol yield (g.<br>g<sup>-1</sup>consumed sugar)</td><td>0.46</td><td>0.45</td><td>0.43</td></tr><tr><td>Efficiency (%)</td><td>88.43</td><td>87.11</td><td>84.84</td></tr></tbody></table>

(Table 1). After 72 h of fermentation the trends in the bioreactor were like those in the flasks. Absorbance was 3.5 in the bioreactor, compared to 2.6 and 3.1 in the 250 mL and 500 mL flasks, respectively. Ethanol concentration, yield, and efficiency in the bioreactor were 8.41 g. L⁻¹, 0.43 g.g⁻¹, and 84.84 %, closely matching those in the flasks: 6.87 and 6.71 g.L⁻¹ for ethanol concentration, 0.46 and 0.45 g.g⁻¹ for yield, and 88.43 % and 87.11 % for efficiency in the 250 mL and 500 mL flasks, respectively. Similar findings were reported in a scale-up of taro waste fermentation using *Kluyveromyces marxianus* K21, where the ethanol concentration was higher in a 250 mL flask (48.98 g.L⁻¹) than in a 6 L bioreactor (43.78 g.L⁻¹) [42]. A decrease in ethanol yield was observed when scaling up from lab to pilot scale in cassava flour fermentation, dropping from 17.2 % v/v (86.1 % of the theoretical yield) to 16.5 % v/v (83.6 % of the theoretical yield) [43].

### 3.3. Influence of pH in 3 L bioreactor

Its effect on ethanol production is shown on Fig. 4. Adjusting the pH of the culture medium to 6 had no significant effect on ethanol productivity (data not shown) and ethanol concentration. Indeed, ethanol concentrations were 8.86 and 8.41 g.L⁻¹ with and without pH control respectively; and the final biomass concentration remained roughly similar, namely close to 0.5–0.77 g.L⁻¹.

During fermentation, pH decreased from an initial value of 6 to 3.49 after 72 h, indicating acid production, which could alter metabolic activity if it reaches a limiting level. pH plays a crucial role by affecting nutrient solubility, membrane permeability, and cellular metabolism. Generally, yeast cells can adjust their metabolism efficiently in mild acidic environments. *S. cerevisiae* can grow within a pH range of 2.4 to 8.6, with an optimal pH between 4 and 4.5 [44]. Studies show that increasing pH from 4 to 5.75 enhances ethanol production by 4.47 units,

but further pH increases (pH = 7) reduce the ethanol production rate [45]. These findings align with those of [46], who reported that pH control (adjusting from 4.5 to 6.5) had no significant effect (P > 0.05) on ethanol productivity and yield during the fermentation of sugar beet juice using immobilized cells on loofa sponge. Therefore, efficient ethanol production is possible without pH adjustment.

### 3.4. Influence of aeration in 3 L bioreactor

In order to maximize the ethanol production using *S. cerevisiae*, the effect of aeration was tested; an optimization of this parameter could improve ethanol yield and production rate [47]. Experiments were conducted in bioreactor with 0.2 vvm aeration, a pH controlled at 6 and at 28 °C; the corresponding results are displayed in Fig. 5.

Results showed that the biomass concentration increased 4 times in the presence of aeration, from 0.5 to 2.0 g.L⁻¹ of biomass; the aeration increased the exposure to oxygen. While ethanol production increased in the absence of aeration; it was 34 % higher. Ethanol production reached 8.86 g.L⁻¹ within 72 h of fermentation under non-aerated conditions. *S. cerevisiae* produced ethanol with an overall yield of 0.46 g ethanol/g sugar, and productivity 0.12 g.L⁻¹.h⁻¹, comparatively to 0.27 g ethanol/ g sugar and 0.08 g.L⁻¹.h⁻¹ for ethanol yield and productivity in the presence of aeration, respectively. The kinetics of growth and

<figure><img src="image_7.png" /><figcaption>**Fig. 5.** Efficiency and ethanol yield obtained after 48 h of fermentation using *S. cerevisiae* growing on a synthetic medium in the following conditions: A: with NH₄Cl as nitrogen source, 0.1 % v/v inoculum size and without salts. B: with peptone, 0.1 % v/v inoculum size and without salt. C: with peptone, 0.1 % v/v inoculum size and with salts. D: with peptone, 1 % v/v inoculum size and with salts (Vt = 500 mL).</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Effects of source of nitrogen, inoculum size and salts on fermentation after 48 h using *S. cerevisiae* growing on a synthetic medium in the following conditions: A: with NH₄Cl as nitrogen source, 0.1 % v/v inoculum size and without salts. B: with peptone, 0.1 % v/v inoculum size and without salt. C: with peptone, 0.1 % v/v inoculum size and with salts. D: with peptone, 1 % v/v inoculum size and with salts (Vt = 500 mL).</figcaption></figure>
production could be affected by physical and mechanical factors such as temperature, aeration and stirring rate.

Studies found that the presence of an appropriate amount of dissolved oxygen made the yeast physiologically healthy and more productive [48]. In this study the higher concentration of biomass can be explained by the presence of oxygen; the fermentation was rapidly used up for the synthesis of membrane components which are essential for growth [49]. Also, aeration at 0.2 vvm led to a 23 % increase in the viable cell mass for fed-batch ethanol fermentation [50].

Under micro-aerated conditions the by-product formation was reduced, such as glycerol, which has important implications in downstream steps for industrial ethanol recovery [50]. Oxygen limited the production of glycerol, for that in the present study the concentration of glycerol under aerated conditions (0.64 g.L⁻¹) was lower than those obtained under non-aerated conditions (1.15 g.L⁻¹). The effect of aeration on glycerol production was evaluated in a research [50]; they demonstrated that a full aeration strategy (0.2 vvm) led to three-fold reduction in glycerol production (from 12 g.L⁻¹ to 4 g.L⁻¹) compared to micro-aerobic culture.

Micro-aeration was found to enhance ethanol yield and productivity by promoting sugar utilization, increasing the NAD supply, and boosting the glycolytic flux, particularly towards phosphoenolpyruvate, which improved glucose transport [51]. However, our results indicated that ethanol yield decreased in fermentation under aerated conditions (0.25 vvm). Previous research using ethanologenic recombinant *Escherichia coli*, has shown that higher oxygen levels (Oxygen Transfer Rate (OTR) = 8, 24 and 100 mmol.L⁻¹.h⁻¹) reduce ethanol productivity by diverting carbon to cell mass and CO₂ production [52]. Optimal ethanol production (19.66 g.L⁻¹) by *Escherichia coli* KO11, was achieved with moderate oxygen transfer rates (e.g., 0.035 vvm), while higher rates led to ethanol consumption by the cells and reduced production, even though the sugar uptake ratio was higher to that observed at OTR = 0 mmol.L⁻¹.h⁻¹ [47]. In agreement with this, other results show that the ethanol concentration increased for micro-aeration at OTR = 4 mmol.L⁻¹.h⁻¹ compared to an absence of aeration (OTR = 0) or a high aeration (OTR = 15 mmol. L⁻¹.h⁻¹) which resulted in the lowest ethanol concentration because of ethanol consumption by the cells [51].

The duration of aeration also affected ethanol yield, with prolonged aeration favoring biomass formation over ethanol production [52,53]. Overall, the results support the conclusion that non-aerated or minimally aerated conditions are more favorable for ethanol production than fully aerated conditions.

### 3.5. *Ulva rigida* hydrolysate fermentation in 3 L bioreactor

The main result for this part is that the yeast *S. cerevisiae* is able to growth on *U. rigida* hydrolysate, there is no inhibition by the substrate

<figure><img src="image_9.png" /><figcaption>**Fig. 6.** Ethanol production, biomass concentration, glucose consumption and glycerol production after 48 h of fermentation of *U. rigida* hydrolysate using *S. cerevisiae* (Vt = 3 L).</figcaption></figure>

for the growth of the yeast. As shown in Fig. 6, after 22 h of fermentation all the glucose was exhausted. Ethanol reached the maximum production at 28 h with a concentration of 9.05 g.L⁻¹ and then remained almost constant. The kinetic growth followed the same trend with a maximum at 24 h (3.2 g.L⁻¹).

Fig. 7 shows a comparison between the results obtained by fermentation of the algae and those obtained by means of the synthetic medium. The results showed a higher ethanol yield and efficiency (experimental ethanol yield/ theoretical ethanol yield) in the algal hydrolysate than those obtained in the synthetic medium, 0.44 g.g⁻¹ and 85.86 % in the algal hydrolysate whereas 0.40 g.g⁻¹ and 78.96 % in the synthetic medium, respectively.

It should also be noticed in this study that the culture on algal hydrolysate of the green algae *Ulva* was done without enrichment. Indeed, nitrogen, trace elements and vitamins are required to obtain rapid fermentation and high ethanol levels. On a laboratory scale, media are often supplemented with peptone and yeast extract. However, such an addition is not feasible in industrial fermentation processes due to the high costs associated; they maximize the capital costs and distillation energy [54,55].

Currently, the two most abundant and feasible biofuels for large-scale production are ethanol from corn or sugarcane and biodiesel from oil crops such as sov or oil palm [55]. The results in this study are also comparable with those obtained with other biomass used for ethanol production at bioreactor-scale. Studies on ethanol production from rice straw in 3 L laboratory batch fermenter under optimum conditions and found that the ethanol product yield and volumetric productivity reached up to 0.50 g.g⁻¹ and 0.53 g.L⁻¹.h⁻¹ respectively [45]. Ethanol productivity of 10.8 g.L⁻¹.h⁻¹ occurring after 3 h of fermentation of soybean molasses by *S. cerevisiae* in 8 L bioreactor, with a product yield of 0.45 g.g⁻¹ of consumed substrate, which represented 88.8 % of the theoretical maximum [56]. Lab-scale production of bioethanol from algal hydrolysate of macro green algae *Ulva* using the yeast *S. cerevisiae* was therefore successfully carried out, showing that algal hydrolysate is a performant source of carbon and nitrogen to produce bioethanol.

### 3.6. *Chaetomorpha linum* hydrolysate fermentation in 77 L bioreactor

For the scale up part, the choice of macroalgae was made on hydrolysate of *Chaetomorpha linum*. The dry matter and the concentrations of chloride, and glucose in the hydrolysate are in g.L⁻¹: 61.5, 9.0, 11 respectively. As shown in Fig. 8, glucose was completely consumed after 9 h of fermentation. Ethanol production began at 5 h (indicating a short acclimation time), with the maximum production of 5.61 g.L⁻¹ achieved after 11 h of cultivation. This production profile corresponds to rapid fermentation, enhancing the process's economic efficiency. Ethanol yield and productivity are crucial factors for the commercial viability of

<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Efficiency (%) and ethanol yields (g.g⁻¹) after 48 h of fermentation by *S. cerevisiae* of *U. rigida* hydrolysate (Vt = 3 L).</figcaption></figure>
<figure><img src="image_11.png" /><figcaption>**Fig. 8.** Glucose consumption (g.L⁻¹), Ethanol production (g.L⁻¹) by *S. cerevisiae* after 24 h of fermentation using the algal hydrolysat of *Chaetomorpha linum* (Vt = 77 L).</figcaption></figure>

any process [40]. In this study, the fermentation successfully converted all glucose into ethanol, reaching maximum ethanol yield and productivity after 7 h, with values of 0.52 g.g⁻¹ and 0.62 g.L⁻¹.h⁻¹, respectively. Table 2 presents review ethanol yield from various macroalgae species (g/g Sugar) with corresponding references.

The ethanol yield obtained in this study is notably higher than those reported in various other studies cited in the literature. Studies documented an ethanol production of 4.72 g.L⁻¹ from 11.85 g.L⁻¹ of glucose [57], while other studies reported an ethanol production of 14.89 g.L⁻¹ with an overall yield of 0.43 g.g⁻¹ and a productivity of 0.93 ± 0.02 g. L⁻¹.h⁻¹ [58]. Additionally, an ethanol yield of 0.45 g.g⁻¹ was achieved during the fermentation of the macroalga *Ulva fasciata Delile* [25]. Interestingly, the ethanol yield from *Chaetomorpha linum* (0.52 g.g⁻¹) is noteworthy, as it aligns closely with the yields reported for various lignocellulosic feedstocks. For instance, corn wheat delivers an ethanol yield of 0.48 g.g⁻¹ of sugars, corresponding to a theoretical conversion efficiency of 96 % [59]. Similarly, the shrub *Prosopis juliflora* achieves an ethanol yield of 0.49 g.g⁻¹ of sugars, with a theoretical yield of 98 % [60], while *Lantana camara* provides a comparable yield of 0.48 g.g⁻¹ of sugars, with a theoretical conversion rate of 96 % [61]. These results highlight the potential of *Chaetomorpha linum* as a promising feedstock for bioethanol production, comparable to well-known lignocellulosic materials.

Previous studies [58,62] estimated large-scale bioethanol production at 3.8 kg per 100 kg (dry weight) for the red alga *Gracilaria verrucosa* and 0.23 kg per 100 kg (fresh weight) for *Kappaphycus alvarezii*. Despite this, the yield and volumetric productivity of ethanol observed are promising, indicating potential for further large-scale and long-term studies to commercially exploit this process. However, the low ethanol percentage is a limiting factor for industrial production, necessitating improved starch enrichment in the algae to achieve adequate ethanol levels. Specifically, 43 kg of algae at 21 % dry weight (equivalent to

<caption>**Table 2**</caption>

Ethanol yields from various macroalgae species (g.g⁻¹ Sugar) with corresponding references.

<table><thead><tr><td>Macroalgae</td><td>Ethanol Yield (g.g<sup>-1</sup> sugar)</td><td>Reference</td></tr></thead><tbody><tr><td>Ulva rigida</td><td>0.44</td><td>This study</td></tr><tr><td>Chaetomorpha linum</td><td>0.52</td><td>This study</td></tr><tr><td>Ulva rigida</td><td>0.41</td><td>[35]</td></tr><tr><td>Saccharina japonica</td><td>0.41</td><td>[63]</td></tr><tr><td>Gracilaria verrucosa</td><td>0.48</td><td>[64]</td></tr><tr><td>Saccharina latissima</td><td>0.42</td><td>[65]</td></tr><tr><td>Gwangalli beach</td><td>0.45</td><td>[66]</td></tr><tr><td>Sargassum sagamianum</td><td>0.38</td><td>[67]</td></tr><tr><td>Saccharina japonica</td><td>0.17</td><td>[68]</td></tr><tr><td>Kappaphycus alverzii</td><td>0.39</td><td>[69]</td></tr></tbody></table>

9.03 kg of dry algae), yielded 175 g of ethanol from 35 kg of hydrolysate with 5.5 g.L⁻¹ ethanol, resulting in an overall yield of 2.04 kg per 100 kg of dry weight of *Chaetomorpha linum*. This corresponds to a 2 % ethanol production yield by the yeast *Saccharomyces cerevisiae* per 100 kg of dried *Chaetomorpha linum* macroalgae.

## 4. Conclusion

From the above results, it can be concluded that the green macroalgae *Ulva rigida* is an attractive raw material for bioethanol production. The laboratory process was scaled up with promising results; aeration was not needed and there was no need for adjusting the pH. Synthetic media adjusted on green macroalgae hydrolysate composition allowed controlling nitrogen, inoculum size and salt contents, and hence to understand the importance of these factors and their effects on ethanol production at pilot scale. The salt brought by the macroalgae decreased ethanol production; while an increase in the size of the inoculum, from 0.1 to 1 % v/v, improved performance. An ethanol concentration of 9.0 g.L⁻¹ was obtained during the culture on hydrolysate of green algae under optimized conditions, without nitrogen supplementation, pH control and sterilization of the algal hydrolysate.

Fermentation at pilot-scale of *Chaetomorpha linum* hydrolysate also exhibited very good glucose to ethanol conversion yield, at par with the theoretical yield of 0.51 g.g⁻¹glucose. However, glucose obtained during processing of the hydrolysates reduced overall biomass to ethanol yield, as well as final ethanol concentration (5.5 g.L⁻¹). It is therefore important to control the hydrolysis phase of green macrophages to optimize the production of easily fermentable sugars. Indeed, any starch not converted to dextrin during liquefaction is a lost opportunity for bioethanol production. It will be interesting to extend the experience in order scale up of fermentation. In this context, it should be noted that an area for improving the global process is the reduction of energy consumption.

### CRediT authorship contribution statement

**Hayet Djalel:** Writing – review & editing, Visualization, Validation, Resources, Methodology, Data curation, Conceptualization. **Alaa Salma:** Writing – review & editing. **Walaa Sayed:** Writing – original draft, Methodology, Conceptualization. **Audrey Cabrol:** Writing – original draft, Visualization, Validation, Conceptualization. **Maud Benoit:** Writing – original draft, Methodology, Conceptualization. **Amance Corat:** Writing – original draft, Methodology, Conceptualization. **Abdeltif Amrane:** Visualization, Validation, Conceptualization. **Ronan Pierre:** Writing – review & editing, Visualization, Validation, Project administration, Funding acquisition, Conceptualization.

### Informed consent statement

Not applicable.

### Institutional review board statement

Not applicable.

### Funding

This work was carried out within the French National Agency for Research for the funding support (Project ANR-14-CE05-0043– Green AlgOhol, Programme Grands Défis Sociétaux 2014).

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
# Data availability

Data will be made available on request.

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