The potential of bioethanol production in marine yeasts and investigation of the optimal conditions of production in the selected isolates

doi:10.21203/rs.3.rs-5265758/v1

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The potential of bioethanol production in marine yeasts and investigation of the optimal conditions of production in the selected isolates

https://doi.org/10.21203/rs.3.rs-5265758/v1

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Factors such as energy consumption, population growth, and anticipated increases in fuel prices are driving the world towards transitioning to cleaner sources of energy. One promising alternative is the production of bioethanol from marine microorganisms, which is gaining attention due to its economic viability and minimal environmental impact. The objective of this study was to isolate and identify yeast strains from the water and sediments of the Bushehr coast that have the ability to produce bioethanol. Furthermore, the study aimed to determine the optimal production conditions for the most effective strain. 18 yeast strains were isolated and identified using morphological and molecular methods. The results of the molecular analyses showed that the isolated yeasts belonged to the genera Pichia and Candida. After assessing the amount of CO₂ produced during the fermentation process, the following strains were selected as the top bioethanol producers: Pichia sp. isolate BK10, Pichia kudriavzevii isolate BK40, Pichia kudriavzevii isolate BK50, Pichia fermentans isolate MK20, and Candida parapsilosis isolate BK30. In our study, we produced bioethanol using the microfermentation method and experimented with different carbon sources like glucose, fructose, lactose, and sucrose. We found that the highest alcohol yield of 12.73% was achieved when glucose was used as the carbon source. Additionally, when we induced mutations with UV-30W light in the superior strain, bioethanol production increased to 15.2%. These marine yeasts have great potential for industrial use in the future, and they could potentially cover alcohol production at the level of small bioreactors.

Bioethanol

Candida parapsilosis

Marine yeast

Microfermentation

Pichia sp

The growing concerns regarding climate change, energy security risks, fossil fuel prices, environmental degradation, and resource depletion have led to increased research on environmentally friendly and sustainable energy alternatives (Osmad et al. 2023). On the other hand, the ever-growing population and shifting demographics have resulted in a continuous increase in global energy demand. It is predicted that total world energy consumption will rise from 549 quadrillion British thermal units (Btu). Consequently, world CO₂ emissions related to energy will also increase. Among greenhouse gases, CO₂ accounts for nearly 55% of global warming, making the reduction of CO₂ emissions from fossil fuels an urgent issue to address the global warming trend. Bioethanol is a renewable and environmentally friendly fuel that can be obtained through the fermentation or distillation of various raw materials. It is considered one of the best alternatives to petrol due to its liquid form and similar characteristics. As a result, many governments have implemented policies to increase the use of bioethanol (Zaky et al. 2018; Maroufpour et al. 2019).

In recent years, there has been significant progress in the field of microbiology and industrial biotechnology, leading to the widespread use of microorganisms, particularly yeasts, for the production of various compounds. This includes applications in the food industry, as well as the production of bioethanol and medicinal compounds. Historically, terrestrial yeasts have been commonly used in the fermentation process to produce bioethanol, while marine yeasts have not received as much attention. However, research on marine yeasts has revealed several promising features compared to their terrestrial counterparts, such as higher tolerance to extreme environments and higher productivity (Kutty and Philip. 2008; Zaky et al. 2014).

Fermentation is a metabolic process that converts sugars into acids, gases, or alcohols. This process occurs in yeasts, bacteria, and even muscle cells when oxygen is lacking. In general, fermentation refers to the growth of microorganisms on a culture medium, often with the goal of producing a specific chemical product. Fermentation occurs when the electron transport chain is nonfunctional, often due to the absence of a final electron acceptor. In such cases, fermentation becomes the primary source of energy production for the cell. During fermentation, NADH and pyruvate, which are produced in glycolysis, are converted into NAD⁺ and an organic molecule depending on the type of fermentation. When oxygen is present, NADH and pyruvate are utilized for ATP production through cellular respiration. This process, known as oxidative phosphorylation, generates more ATP than glycolysis. As a result, fermentation is typically not utilized when oxygen is available to the cells (Bamforth and Cook. 2019). Fermented products play a significant role in various industries, including food, beverage, and alcohol production. Africa is the leading producer of fermented starch products and legume-based food products, while South and Southeast Asia are known for their production of fermented fish products. On the other hand, North America is the largest producer of fermented beverages and meat products worldwide. Throughout the centuries, fermentation methods have evolved, improved, and expanded, leading to increased production of fermented products such as bread, cheese, butter, and yogurt on a global scale (Rajauria et al. 2016). Alcohol fermentation is a crucial process in the production of wine, beer, alcoholic beverages, bioethanol from carbohydrate sources, and the production of byproducts in the food industry. Today, ethanol derived from natural sources such as lignocellulosic compounds, corn starch, and wheat straw has gained popularity as an alternative in various industries. The use of fermenting microorganisms that utilize complex carbohydrates like cellulose and hemicellulose to produce bioethanol has also emerged as a promising approach Panahi, 2019; Vamvakas and Kapolos, 2020). One strategy to increase bioethanol production is the development of microbial strains for the fermentation process (Stanley et al. 2010). Mucor indicus fungi, Saccharomyces cerevisiae, and Kluyveromyces marxianus yeasts are among the microorganisms utilized in this field. For instance, M. indicus fungi achieved a production of 0.46 g/g of ethanol through anaerobic cultivation (Karimi et al. 2006). Similarly, the yeasts Kluyveromyces marxianus (Sansonetti et al. 2011) and Saccharomyces cerevisiae (Zaky et al. 2018) produced 22 g/l and 82 g/l of bioethanol, respectively. Recently, seaweeds have also been explored as potential raw materials for bioethanol production (Mushlihah et al. 2020). Moreover, various studies have demonstrated that a wide range of marine yeasts, including Candida albicans, Candida tropicalis, Pichia fermentans, and Rhodotrula minuta, are capable of producing bioethanol (Senthilraja et al. 2011). Marine yeasts exhibit great potential for industrial applications (Kutty and Philip, 2008). Ecological and technological research has shown that Saccharomyces yeasts are also involved in the fermentation of juices (Kurtzman et al. 2011) and play a significant role in the production of industrial alcohol (Benjaphokee et al. 2012). For example, the sequential inoculation of Saccharomyces cerevisiae and non-Saccharomyces yeasts is an interesting approach to obtain alcoholic beverages with enhanced aroma and taste. T. delbrueckii Biodiva strain, for instance, increases glycerol concentration, reduces volatile acidity, and improves foaming properties, thereby enhancing the shelf life of alcoholic and non-alcoholic beverages. Saccharomyces cerevisiae yeast, on the other hand, is considered one of the most compatible and valuable microorganisms in the fermentation and industrial production of bioethanol (Ceccato-Antonini and Covre. 2020; Zhang et al. 2015). This study aimed to isolate and identify marine yeast strains from the Persian Gulf that are capable of producing bioethanol, as well as investigate the optimal production conditions for the most effective strain in fermentation environments.

2.1. Samples and sampling

In this study, water and sediment samples were collected from the coast of Bushehr province in 2022. The sampling was conducted at three stations: the student park, Halile, and Delvar (Table 1). Seabed sediment samples were collected using a grab from a depth of approximately 2 meters below the water surface. These samples were then placed in sterile zipped plastic bags. Water samples were taken from a depth of 15 to 20 cm below the water surface and stored in sterile glass bottles. Once the samples arrived at the laboratory of Khorramshahr Marine Science and Technology University, they were immediately stored in a fridge at 4°C until isolation was performed (Zaky et al. 2016).

Table 1

Geographical location of the investigated stations.
Station	Longitude	Latitude
student park Halile Delvar	´´758 81´ ° 50 ´´092 87´ ° 50 ´´436 04´ ° 51	´´305 90´ ° 28 ´´413 84´ ° 28 ´´763 75´ ° 28

2.2. Isolation and purification of marine yeasts

2.2.1. Method of preparation and enrichment of sediment and water samples

YPD (yeast extract, peptone, and dextrose) liquid culture medium was used to enrich yeasts. Once the medium was prepared, the pH was adjusted to 7.2. The jars were then autoclaved for 15 minutes at 121°C. Replicate samples from each station were mixed together under sterile conditions using a flame, and 5 grams of sediment mixture from each station were transferred into separate 100 ml Erlenmeyer flasks containing 50 ml of YPD culture medium. As for the water samples, 5 mL of each sample were transferred to the 100 ml Erlenmeyer flasks containing 50 ml of culture medium, all done under sterile conditions. Finally, the jars were placed on a shaker for 5 days at 30°C and 200 rpm (Zaky et al. 2016).

2.2.2. Isolation method from water and sediment

After 5 days of incubation and completing the enrichment phase (which was determined by observing turbidity, indicating yeast growth), the fungal colonies were diluted. To achieve this, 1 mL of the enriched medium was added to a test tube containing 9 mL of sterile distilled water. After thorough mixing, 1 mL of this mixture was transferred to tube number 2, which contained 9 mL of distilled water. This process was repeated to prepare a series of dilutions ranging from 10 − 1 to 10 − 4. Next, 100 µL of each prepared dilution was transferred to solid YPD culture medium and surface cultured using the spread plate method. The plates were then incubated for 48 hours at 33°C until colonies appeared. In the next step, colonies with different appearances were selected and cultured linearly on YPD solid culture medium. This process was repeated until the colonies were finally purified(Zaky et al. 2016). After 5 days of incubation and completion of the enrichment phase (indicated by turbidity, which signifies yeast growth), the fungal colonies were diluted. To dilutethe sample, 1 mL of the enriched medium was added to a test tube containing 9 mL of sterile distilled water. After thorough mixing, 1 mL of this mixture was transferred to another tube, labeled as tube number 2, which contained 9 mL of distilled water. This process was repeated to create a series of dilutions ranging from 10^− 1 to 10^− 4. Next, 100 µL of each prepared dilution was transferred to the solid YPD culture medium and spread evenly using the spread plate method. The plates were then incubated at 33°C for 48 hours, allowing colonies to develop (Gao et al. 2007; Zaky et al. 2016).

2.3. Yeast identification methods

Yeasts were identified based on morphological and molecular methods by sequencing 18S rRNA genes (Kurtzman et al. 2011a)

2.3.1. Identification of morphology

The first identification feature is the appearance of yeasts. Therefore, we investigated the characteristics of yeast colonies in terms of size, shape, color, and border. To ensure more accurate identification, various microbial tests and biochemical tests were including employed, sporulation tests, growth tests, glucose fermentation, assimilation of carbon compounds, and urea hydrolysis (Gao et al. 2007).

Each media was prepared into bottles as slopes after being autoclaved at 121ºC and15 minutes.

2.3.2. Sporulation test

An amount of 30 g per liter of PDA (Potato Dextrose Agar) culture medium was prepared. It was then poured into a plate and inoculated with a fresh yeast culture. The plate was grown in an incubator for 3 days. For the sporulation test, a smear was prepared and stained with 5% malachite green.The sample was then investigated under a microscope (Sulieman et al. 2015; Kurtzman et al. 2011).

2.3.3. Growth test

For the growth test at 37ºC and 40ºC, YMA (3g yeast extract, 3g malt extract, 10g glucose, 5g peptone, and 20g agar in 1000 ml distilled water) media was inoculated with a fresh culture of yeast and incubated aerobically at 37ºC and 40ºC (Sulieman et al. 2015; Kurtzman et al. 2011).

2.3.4. Fermentation of glucose test

The fermentation of glucose was carried out by filling test tubes with 10–15 ml of yeast extract broth containing 2% D-glucose with inverted Durham tubes, then autoclaved at 121ºC for 20 minutes. Each test tube was inoculated with a fresh yeast suspension and then incubated at 25ºC for 2 weeks, being examined frequently for gas bubbles(Sulieman et al. 2015; Kurtzman et al. 2011).

2.3.5. Assimilation of carbohydrates

For the assimilation of carbohydrates, small test tubes, each containing 10ml of sterile medium composed of 0.5% pepton water with 4% test sugar, were used. The tubes were inoculated with the selected yeast cultures in duplicate. A Vaseline-paraffin (Vaspar) layer, 2cm deep, was added to the top surface of one tube of the medium. The culture was incubated at 25ºC for four to five days. Fermentation was detected by the lifting of the Vaspar layer (Sulieman et al. 2015; Kurtzman et al. 2011).

2.3.6. Urea hydrolysis test

Urea broth medium was used for urea hydrolysis, which was distributed into 5ml tubes and then autoclaved at 121 degrees for 20 minutes. A ring full of cells from a one or two-days-old fresh culture was suspended in the broth and incubated at 37°C. The tubes were examined every half hour for a change in color to red, indicating urea activity (Sulieman et al. 2015; Kurtzman et al. 2011).

2.3.2.1. DNA extraction

The phenol-chloroform method was used to extract yeast DNA. To check the quality of the extracted DNA, the horizontal electrophoresis method was used on a 1% agarose gel. Electrophoresis was performed with a voltage of 120V and 200mA for 45 minutes, and finally, the DNA band and its quality were checked using a gel documentation system (Ghasemi et al. 2014).

2.3.2.2. PCR of ITS regions

For the amplification of the 18S rRNA gene, ITS primers shown in Table 2 were used. The PCR reaction was performed with a final volume of 25 µL and the materials shown in Table 3. All the steps of mixing the materials to prevent the activity of the polymerase enzyme and the formation of primer dimers were done on ice. Then, the PCR technique was performed with a thermocycler according to the thermal cycles, amounts, and materials consumed, in the order mentioned in Tables 2 to 4 (Augustine and Joseph. 2018; Greetham et al. 2019).

Table 2

Primers used for PCR.
Primer	Primer sequence	junction temperature
ITS1F ITS4	5´- CTTGGTCATTTAGAGGAAGTAA-3 5´-TCCTCCGCTTATTGATATGC-3´	49.3 ˚C 49.7 ˚C

Table 3

Quantities and materials used in PCR.
Material	Amount for reaction 25 µl
DNA extraction(100 ng/ µL) Master mix (µL 10 pmol/) primer1 (µL 10 pmol/) Primer2 Sterile distilled water	2µL 12.5 µL 1 µL 1 µL 8.5 µL

Table 4

Thermal cycle of PCR machine.
Cycle	Time	(ºC) Temperature	Level
1	3 Min	95	1
10X	30S 30S 1Min	95 62 72	2
20 X	30S 30S 1Min	95 52 72	3
1	10 Min	72	4
1	∞	4	5

2.3.2.3. PCR Product Quality Check Method and PCR Product Sequencing Method

To check the presence and quality of PCR products, the products were transferred to a 1% agarose gel, and electrophoresis was performed at 120 V for 45 minutes. DNA fragments were identified by comparison with a 100 bp DNA base pair marker (Lilja. 2013). Once PCR was performed and the quality and quantity of the product were confirmed, it was sent to Feza Research Company for sequencing. The obtained chromatograms were then analyzed using Chromas Pro software to verify the sequences and identify the yeasts. The starting and ending nucleotides of the sequences were trimmed, and suitable sequences were selected for searching in the gene bank. The BLAST tool on the NCBI website was used to identify yeasts using these sequences (Newell et al. 2018).

2.4. Fermentation

This experiment was conducted on a small scale using earlens, plastic covers, and syringes instead of a bioreactor. Each component was sterilized separately at 121°C for 15 minutes. Hararlen, containing the fermentation medium and yeast inoculation, was completely sealed with a plastic cap to create anaerobic conditions. It was then placed on top of the syringe cap to release gas (Zaky et al. 2018; Greetham et al. 2019).

2.5. Method for Detecting the Best Bioethanol-Producing Yeast (Screening)

One liter liquid culture medium, with a pH of 6, containing 55 g glucose, 20 g peptone, and 10 g yeast extract, was prepared and autoclaved at 110°C for 10 minutes. Test tubes were filled with 10 ml of this medium and inoculated with a loopful of yeast. The tubes were incubated at 30°C and 200 rpm in a shaker until they reached the appropriate turbidity, based on the peak growth time of each yeast. Cloudy tubes were vortexed for 10 seconds at their maximum activity, and 5 ml was transferred to Erlenmeyer flasks containing 50 ml of culture medium (maintaining a 1:10 ratio). The Erlens were then incubated at 30°C and 200 rpm in a shaker according to the peak activity time of each yeast. A syringe was placed on the cap of each Erlen to release gas (Zaky et al. 2018; Greetham et al. 2019).

2.6. Distillation and Production of Bioethanol

After the micro-fermentation, the Erlens containing the fermentation culture medium were placed in a simple distillation system during peak fermentation activity to separate the alcohol from the culture medium. The culture medium was heated to a fixed temperature of 78°C, and the distillation process lasted for approximately 3 hours. The percentage of isolated bioethanol was measured using gas chromatography (GC) (Somboon and Sansuk. 2018; Somboon et al. 2022).

2.7. Bioethanol Assay

2.7.1. Proofread version:

To measure the alcohol percentage, we used the Young Lin gas chromatography device from South Korea, equipped with a flame ionization detector (FID detector). The column used was TRB-5, purchased from TEKNOKROMA, with dimensions of 30m × 0.53mm × 0.5µm. A temperature program was designed for liquid sample analysis in GC. The initial temperature of the column was set at 70°C and maintained for 2 minutes. After 2 minutes, the temperature was raised to 250°C in a gradient of 10°C and held for 5 minutes. The detector temperature was 290°C, and the injector temperature was 270°C. The carrier gas (hydrogen) flow rate was 5 ml/min, and the sample volume injected into the device was 1 microliter. Each experiment was repeated three times to ensure accuracy, and the percentage of production was reported (Somboon and Sansuk. 2018; Somboon et al. 2022).

2.8. Optimization

2.8.1. Bioethanol production using different carbohydrates

A culture medium containing 55 g of various carbohydrates (lactose, sucrose, fructose instead of glucose as a carbon source), 20 g of peptone, and 10 g of yeast extract was prepared in a 1000 ml volume. The medium was then autoclaved at 110°C for 10 minutes (Urano et al. 2021). Following the mentioned procedure, micro-fermentation, distillation, and measurement of alcohol production percentage were carried out using a gas chromatography device.

2.8.2. UV-mutagenesis of the selected yeast isolate

The selected yeast isolate's culture was inoculated in 10 ml of YPD medium and incubated at 30°C for 12 hours until the cell density reached 2 × 10⁸ cells/ml. Subsequently, the cells were washed multiple times with 0.1 M phosphate buffer (pH = 5.4) and sterile distilled water. Once the cell density reached 100 cells/ml, 100 µl of the cell suspension was spread onto plates containing YPDA (yeast extract, peptone, dextrose, agar). The plates were then exposed to UV irradiation (wavelength 254 nm) at an intensity of 30 W for 5, 10, 15, and 20 minutes. After the exposure, the plates were kept in the dark for 24 hours and then incubated for 3 days at 30°C. The surviving mutant colonies were selected and streaked on YPD medium (Thammasittirong et al. 2013; Koti et al. 2016).

2.9. Statistical analysis

For statistical data analysis and graphing, SPSS v25 software was used. The Kolmograph-Smirnov normality test was employed to ensure the normal distribution of data. ANOVA-Tukey tests were conducted to compare the significance level between the samples. Data means and standard deviations were calculated using Microsoft Excel. In the statistical analysis, a P-value < 0.05 was considered significant.

3.1. Morphological analysis

During sampling, a total of 18 yeast strains were isolated, purified, and examined under the microscope using crystal violet staining. Among these 18 strains, 5 were selected as the most proficient bioethanol producers. This selection was based on the amount of CO₂ gas produced during the anaerobic respiration process, as shown in Table 5.

Table 5

Colony characteristics of isolated strains.
Strains	Colony color	Vegetative growth	Colony shape	Sampling location
B1	White - Dull matte	Budding	Spherical	Halile - Sediment
B2	creamy	Budding	Oroid	Halile - Sediment
B3	creamy	Budding	Spherical	Halile - Sediment
B4	White - Dull matte	Budding	Oroid	Water-Delvar
B5	White - Dull matte	Budding	Oroid	Sediment - Student Park

3.2. Biochemical analysis

Identification was done based on the results of biochemical tests and microscopic features. For more certainty, molecular methods were also used (Tables 6 and 7).

Table 6

Results of biochemical identification of bioethanol producing yeasts.
Strains	Assimilation of carbon compounds					Urea Hydrolysis	Fermentation of glucose
Strains	Fructose	Maltose	Sucrose	Lactose	Glucose	Urea Hydrolysis	Fermentation of glucose
B1 B2 B3 B4 B5	+ + + + +	- - - - -	- - + - -	- - - - -	+ + + + +	- - - - -	+ + + + +

Table 7

Results of biochemical identification of bioethanol producing yeasts.
Strains	Growth at 42 ºC	Growth at 37 ºC	Sporulation	Pseudohypha
B1	+	+	+	+
B2	+	+	+	+
B3	+	+	+	+
B4	+	+	+	+
B5	+	+	+	+

3.3. Molecular analysis

After examining the sequences and removing the first and last nucleotides, we selected the relevant parts of the sequences to search in the gene bank. We performed a search in the NCBI database to find similar sequences and identify the isolates based on their highest degree of sequence similarity with the information available in this database. Table 8 presents the results, showing that four strains belonged to the genus Pichia, while one strain belonged to the genus Candida.

Table 8

BLAST results of 18S rRNA in base.
Strains	Description
B1	Pichia sp. isolate BK10
B2	Pichia fermentans isolate MK20
B3	Candida parapsilosis isolate BK30
B4	Pichia kudriavzevii isolate BK40
B5	Pichia kudriavzevii isolate BK50

3.4. Measuring the growth rate of yeasts

As shown in Fig. 1, the rate of yeast growth in 24 hours follows a similar trend. Initially, the growth rate was slow, but after approximately 14 hours, strains B1, B4, and B5 reached their peak growth time. Strains B2 and B3, on the other hand, reached their peak growth after 18 and 22 hours, respectively.

3.5. Bioethanol production

Depending on the time of peak growth for each yeast, micro-fermentation (creating anaerobic conditions on a small scale) was performed at the specified time. Distillation was then carried out when the yeasts reached their peak fermentation activity (as indicated by visible foam production) (Fig. 2). The production percentage was measured using a GC device. Figure 3 shows a significant difference in the percentage of bioethanol produced by different strains. Strains B1, B4, and B5 produced 12.23%, 10.35%, and 12.71% bioethanol, respectively. Strains B2 and B3 produced 9.67% and 9.85% bioethanol, respectively.

3.6. Carbon source change analysis

In this analysis, four carbon sources were used for bioethanol production, in addition to glucose. Table 9 shows that the B4 strain had the highest bioethanol production at 10.14% when fructose was consumed, while the B2 strain had the lowest production at 5.31%. Strain B3 produced 4.53% bioethanol when consuming sucrose and 2.48% when consuming lactose. The B1, B2, B4, and B5 strains produced less than 1% bioethanol when consuming sucrose and lactose, indicating that they were not very efficient in absorbing and hydrolyzing these carbohydrates. Overall, glucose was found to be the most efficient carbon source for these yeasts, while lactose was the least efficient source.

Table 9

The results of bioethanol production by changing the carbon source.
Carbon source	B1	B2	B3	B4	B5
Glucose(%)	12.23	9.67	9.85	10.35	12.71
Fructose (%)	7.0	5.31	10.2	10.14	7.47
Sucrose (%)	0. 31	0.25	4.53	0.24	0.4
Lactose(%)	0.7	0.8	2.48	0.04	0.02

3.7. UV‑mutagenesis

The B5 strain was initially identified as the most efficient producer of bioethanol. Subsequently, a mutation was induced in this yeast using UV-30W light. At the 0 time point (control), approximately 100 colonies were observed to grow. However, after 5 minutes of exposure to UV light, only 2 colonies survived, and after 10 minutes, only 1 colony remained viable (Figs. 4 and 5). Following growth on YPD medium, the single surviving colony was transferred to the fermentation medium, resulting in the production of 15.2% bioethanol. The bioethanol production by the mutant strain was then compared to that of the wild strain. Figure 7 clearly demonstrates a significant difference in the amount of bioethanol produced between the parent strain and the mutant strain.

The reduction of energy resources, the increase of greenhouse gases, and environmental pollution have led researchers to focus on the production and use of renewable fuels derived from biological sources. In the last decade, the biosynthesis of alcohols, as a green chemistry approach, has garnered significant interest from researchers. Bioethanol production using biological resources is advantageous due to its minimal environmental risks, lack of harmful pollution, and affordability from an economic standpoint. Various methods have been developed for the biological production of ethanol, including fermentation of agricultural products containing starch and sugar from land sources, as well as marine sources. One approach involves the production of bioethanol from marine organisms and microorganisms such as algae, fungi, yeasts, and bacteria (Megia et al. 2021; Hang t al. 1986).

The production of bioethanol among different strains was carried out by controlling various parameters in the fermentation process, such as temperature, amount of substance, pH, etc. The results revealed a significant variation in the percentage of bioethanol produced by different strains. Therefore, the specific activity of each strain in fermentation and anaerobic conditions was the sole factor that influenced the percentage of bioethanol production. Despite the distinct phylogeny of B2 and B3 strains, there was not a substantial difference in their bioethanol production. Consequently, Pichia kudriavzevii isolate BK50, with a production rate of 12.71%, and Pichia fermentans isolate MK20, with a production rate of 9.67%, were identified as the strongest and weakest bioethanol producers, respectively, in this research. The micro-fermentation method (fermentation on a small scale) was utilized to establish anaerobic conditions. Zaky et al. discovered that nine isolated marine yeast strains exhibited potential for bioethanol production. Furthermore, these yeast strains were capable of producing bioethanol through microfermentation under anaerobic conditions (Zaky et al. 2018).These findings, combined with a comparison between the bioethanol yield from the activity of an industrial yeast strain S. cerevisiae NCYC2592 and a marine yeast strain isolated from the United States' sea, referred to as S. cerevisiae AZ65 by Zaky et al. in another study, indicate that marine yeasts and seawater hold great potential for bioethanol production using a small-scale fermentation process (Zaky et al. 2020). Studies have demonstrated that microfermentation is faster than conventional methods like bioreactor usage and requires minimal energy. Additionally, since this reaction occurs on a small scale, the by-products, such as heat and CO₂ production, do not disrupt the reaction process.

In the present study, changes in carbon sources were investigated while keeping other fermentation conditions constant. Bioethanol production was significantly higher with fructose as the carbon source, although the production rate was lower compared to glucose. Yeast strains using sucrose and lactose produced less bioethanol than the other two sources. Therefore, glucose and fructose, which are simple sugars, were identified as the best substrates for fermentation and bioethanol production. These strains were also found to be less efficient in hydrolyzing disaccharides such as sucrose and lactose to produce bioethanol. It appears that yeasts require less energy for the hydrolysis of simple sugars (monosaccharides) compared to the hydrolysis of disaccharides like lactose and sucrose. Despite the tests and result analysis, it seems that optimizing the carbon source parameter was not very successful. Mosier et al. also concluded that yeast cells generally prefer simple sugars and monosaccharides for growth. They also found that D-glucose, among monosaccharide and disaccharide sugars, is the most conducive substrate for yeast cell growth and bioethanol production (Mosier et al. 2005). Blomberg and Adler similarly concluded that carbon source directly affects bioethanol production, while the nitrogen source indirectly improves its efficiency (Blomberg and Adler, 1992).

In this research, after subjecting the best bioethanol producing strain to UV-30W lamp light for 10 minutes, only one colony of strain B5 was able to survive on YPDA medium. The mutant monocolony did not differ significantly in appearance and size from the wild strain, but after cultivation on YPD medium and the bioethanol production process, the production rate increased to 15.2% compared to the parental strain. It seems that the mutation created minor and repair changes in the gene of the parent strain, resulting in an increased amount of bioethanol. In general, exposure to ultraviolet rays strongly affects yeast activity and can enhance yeast's ability to absorb sugar through random mutagenesis, thus increasing the efficiency of bioethanol production. Hawary et al. (2019) observed that exposure of the Wickerhamomyces anomalus HH16 strain to ultraviolet (UV) rays (UV-30W light with a wavelength of 254 nm) for 15 minutes resulted in an 80.16 g/L increase in glycerol production. The mutation caused a decrease in the number of cells, but the remaining cells underwent mutation and enhanced production efficiency. This finding suggests that random mutagenesis in yeast using mutagenic agents like UV light or chemicals such as ethyl methane sulfonate can be a promising strategy for improving fat and sugar alcohol production (Hawary et al. 2019). Similarly, Aruna et al. (2015) exposed the Candida albicans strain to ultraviolet light for 5 to 100 seconds, resulting in mutation. The mutation destroyed over 90% of the colony. However, the mutant strain was able to produce 437 g of bioethanol from one kg of potato (used as a carbon source) through small-scale fermentation. This amount of bioethanol production was approximately 1.5 times higher than that of the wild strain. Hence, the Candida albicans strain exhibits less resistance to ultraviolet light and mutates in less than 2 minutes, thereby increasing bioethanol production efficiency (Aruna et al. 2015). In the study conducted by Koti et al. (2016), the mutagenesis of yeasts using ultraviolet light, as well as changes in environmental pH, were investigated for bioethanol production. Although changes in pH had an effect on the anaerobic fermentation process, yeast strains subjected to mutagenesis produced more bioethanol (Koty et al. 2016). Hawary et al. (2019) also concluded that mutagenesis with UV light is a crucial tool for enhancing production efficiency. Other variables, such as changes in carbon source, substrate type, nitrogen source, initial concentration, and fermentation period, significantly influenced yeast growth, but their efficiency was lower compared to mutagenesis (Hawary et al. 2019). This finding aligns with the results of the present study.

Conclusion.5

The current work presents The marine environment has great potential as a source for new yeast isolates with promising properties. The yeasts identified in this research have shown potential in participating in the fermentation process due to their high bioethanol production. Production rates can also be increased through mutations and metabolic engineering, potentially allowing for alcohol production in small bioreactors. There are various methods for optimization, such as changing temperature, carbon source, nitrogen source, and pH, as well as inducing mutations using ultraviolet light or chemicals. However, mutating with ultraviolet light has proven to be the most effective method for increasing bioethanol production. This method is preferred due to its lack of environmental pollution, unlike chemical usage, and its ability to improve production efficiency within a shorter time period. Furthermore, the research results suggest that the use of marine yeasts for bioethanol production could be an efficient alternative to fossil fuels.

Acknowledgements

gratitude and appreciation to Khorramshahr University of Marine Science and Technology for their invaluable support throughout our research project. We would also like to acknowledge the contribution of the technical staff and laboratory assistants. Their unwavering support in providing us with the necessary resources and equipment has greatly facilitated our research endeavors. Our gratitude also goes to the support staff and personnel who have facilitated the administrative processes and logistics involved in this project. Lastly, we would like to express our gratitude to the entire university community, including fellow students and colleagues. Your constant support, discussions, and constructive criticism have contributed significantly to our understanding and development of various aspects related to our research.

In conclusion, the support and encouragement we received from Khorramshahr University of Marine Science and Technology have been invaluable. We are truly grateful for the opportunity to be part of this esteemed institution and for the countless opportunities it has provided us.

Contributions

Authors' Contributions Banafsheh Khajeh and Hosein Zolgharnein conducted the experiments and data curation, Isaac Zamani and Kamal Ghanemi analyzed the results, Banafsheh Khajeh, Kamal Ghanemi, Hosein Zolgharnein reviewed and edited the manuscript, and Banafsheh Khajeh designed the study. All authors read and approved the final manuscript.

Funding

Faculty of Marine Science and Oceanography, Khorramshahr University of Marine Science and Technology, Khorramshahr, Iran.

Data Availability

The datasets used and analyzed during the current study are available.

Ethical Approval Not applicable.

Consent to Participate Not applicable.

Consent to Publish Not applicable.

Conflicts of interest/Competing interests The authors declare that they have no competing interests.

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The potential of bioethanol production in marine yeasts and investigation of the optimal conditions of production in the selected isolates

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Abstract

Figures

1. Introduction

2. Materials and methods

2.1. Samples and sampling

2.2. Isolation and purification of marine yeasts

2.2.1. Method of preparation and enrichment of sediment and water samples

2.2.2. Isolation method from water and sediment

2.3. Yeast identification methods

2.3.1. Identification of morphology

2.3.2. Sporulation test

2.3.3. Growth test

2.3.4. Fermentation of glucose test

2.3.5. Assimilation of carbohydrates

2.3.6. Urea hydrolysis test

2.3.2.1. DNA extraction

2.3.2.2. PCR of ITS regions

2.3.2.3. PCR Product Quality Check Method and PCR Product Sequencing Method

2.4. Fermentation

2.5. Method for Detecting the Best Bioethanol-Producing Yeast (Screening)

2.6. Distillation and Production of Bioethanol

2.7. Bioethanol Assay

2.7.1. Proofread version:

2.8. Optimization

2.8.1. Bioethanol production using different carbohydrates

2.8.2. UV-mutagenesis of the selected yeast isolate

2.9. Statistical analysis

3. Results

3.1. Morphological analysis

3.2. Biochemical analysis

3.3. Molecular analysis

3.4. Measuring the growth rate of yeasts

3.5. Bioethanol production

3.6. Carbon source change analysis

3.7. UV‑mutagenesis

4. Discussion

Declarations

References

Status:

Version 1