Skip to main content
NCBI home page
3 Biotech logoLink to 3 Biotech
. 2020 Feb 24;10(3):135. doi: 10.1007/s13205-020-2128-3

Bioprocessing of shrimp wastes to obtain chitosan and its antimicrobial potential in the context of ethanolic fermentation against bacterial contamination

Isabella C Tanganini 1, Ligianne D Shirahigue 1, Mariana Altenhofen da Silva 1, Kelly R Francisco 2, Sandra R Ceccato-Antonini 1,✉
PMCID: PMC7040113  PMID: 32158631

Abstract

This study investigated the bioprocessing of shrimp wastes to obtain chitin and its deacetylated product chitosan by a fermentation process mediated by Lactobacillus plantarum. The concentrations of glucose, bacterial inoculum, and shrimp wastes in the Man, Rogosa and Sharpe medium were optimized for the fermentation process performed in shake flasks to achieve the maximum titratable acidity to obtain chitin. The experiments were scaled up in a 700-mL working volume bioreactor, and the resulting chitin was deacetylated by the autoclave method. The bioextracted chitosan was characterized (Fourier transform infrared spectroscopy [FTIR], deacetylation degree, and molecular weight) and evaluated for its antimicrobial effects by comparing it with a commercial chitosan sample in the context of the ethanolic fermentation process for fuel alcohol production. The effect of chitosan on such a fermentation process has not been determined yet. The bacterial contaminant Lactobacillus fermentum and the main agent of ethanolic fermentation Saccharomyces cerevisiae were cultured in semi-synthetic medium and co-cultured in sugarcane juice to verify the effect of chitosan on their growth. The bioextracted chitosan (molecular weight 4.0 × 105 g mol−1 and deacetylation degree 80%) was comparable to commercial chitosan, although higher concentrations of the former were required to achieve similar antimicrobial activities. Both commercial and bioextracted chitosan samples exhibited antimicrobial activity against S. cerevisiae and L. fermentum, but the concentration that caused the inhibition of yeast growth was almost tenfold higher than for the bacterium. Moreover, bioextracted chitosan showed no yeast inhibition or lethality in the range of 0.0075–0.96% while for the bacterium, growth inhibition occurred in concentrations varying from 0.24 to 0.48% and lethality of more than 99% at 0.96%. These results indicate the potential use of chitosan and especially of bioextracted chitosan in the bioethanol industry as a safer and more natural approach to combat unwanted bacterial contamination.

Keywords: Bioethanol, Chitin, Lactic acid bacteria, Saccharomyces cerevisiae, Sugarcane juice

Introduction

Bioextraction of chitin and chitosan from the exoskeletons of crustaceans can be performed by one or multiple fermentation processes using microorganisms. These methods have been proposed as suitable and economic alternatives to the traditional chemical methods that involve the use of strong acids and alkali. The biological methods of chitin extraction from shrimp waste, which involve demineralization (DM) and deproteinization (DP) of the material, produce higher yields compared to the traditional chemical methods (Hamed et al. 2016).

Due to the high production rates of organic acids, lactic acid bacteria are preferred for shrimp shell fermentation as they prevent the deterioration of this highly perishable waste product (Prameela et al. 2010; Ploydee and Chaiyanan 2014; Neves et al. 2017; Sedaghat et al. 2017). The efficiency of the fermentation process, i.e., DP and DM, depends on the optimization of fermentation parameters such as substrate concentration (glucose), level and type of inoculum, load and particle size of shrimp shells, pH, and fermentation time (Ghorbel-Bellaaj et al. 2013; Sedaghat et al. 2017).

Bacteria such as Pseudomonas aeruginosa (Sedaghat et al. 2017), Bacillus subtilis (Sini et al. 2007), Streptococcus thermophilus (Mao et al. 2013), and Lactobacillus sp. (Jung et al. 2006; Wahyuntari et al. 2011; Zhang et al. 2012) have been used for shrimp waste fermentation. Lactobacillus plantarum, a homofermentative bacterium, exhibits rapid acidification and proteolytic activity (Liu et al. 2010), making it a promising microorganism for the bioextraction of chitin from shrimp waste (Rao and Stevens 2005; Khanafari et al. 2008; Prameela et al. 2010; Zhang et al. 2012; Ploydee and Chaiyanan 2014; Neves et al. 2017).

One of the most important chitin derivatives is chitosan, the only known natural cationic polysaccharide, which is obtained after chitin deacetylation (Du et al. 2014; Hamed et al. 2016). While chitin is insoluble in water and most organic solvents, chitosan is soluble in dilute aqueous acidic solutions. This polymer has special characteristics that make it useful for several applications in agriculture, medicine, cosmetics, food packaging, and additives, and also as an antimicrobial agent (Raafat and Sahl 2009).

Chitosan exhibits antimicrobial activity against a wide range of microorganisms, including filamentous fungi, yeasts, and bacteria, and is more effective against Gram-positive than Gram-negative bacteria (Jeon et al. 2001; Raafat and Sahl 2009; Verlee et al. 2017). It also exhibits antimycotic activity at concentrations ranging from 10 to 7750 mg L−1 but the effect is highly dependent on the microbial species (Verlee et al. 2017). Chitosan damages the cell membrane through electrostatic interactions with negatively charged phospholipids. Once the cell membrane is disrupted, it enters the cell and directly affects nucleic acid and protein synthesis (Marquez et al. 2013).

Lactic acid bacteria are considered the most important bacterial contaminants in the fermentation processes of fuel ethanol production, especially Lactobacillus fermentum (Beckner et al. 2011; Basso et al. 2014). Bacterial growth in the bioethanol industry is controlled by sulfuric acid treatment and/or the use of antibiotics, both posing safety and environmental problems (Ceccato-Antonini 2018). So far, no studies have reported the use of chitosan as an alternative method for controlling potential bacterial contaminants such as L. fermentum during fuel ethanol fermentation. Ristić et al. (2015) reported some controversial data about the effect of chitosan on Lactobacillus species; according to their study, chitosan exhibited both stimulant and inhibitory effects on bacterial growth. Lee et al. (2002) verified that chitosan oligosaccharides stimulated the growth of most Lactobacillus, while Ristić et al. (2015) established the minimum inhibitory concentration of 0.015625% (w/v) of chitosan for action against Lactobacillus sp. Moreover, some literature reports have focused on species of Lactobacillus other than L. fermentum. Jeon et al. (2001) verified that lactic acid bacteria including L. fermentum were inhibited by chitosan at concentrations lower than 0.03%.

Hence, for identifying natural and safer products to combat the issue of microbial contamination in the bioethanol industry, we aimed to (1) optimize a few conditions of the shrimp waste fermentation process by L. plantarum to obtain chitin and subsequently chitosan by chemical deacetylation, and (2) assess the potential of chitosan (bioextracted and commercial) application as an alternative antibacterial agent in the bioethanol industry using L. fermentum as the target microorganism.

Methods

Optimization of acid production by L. plantarum in a semi-synthetic medium

The strain of L. plantarum (CCT0580–ATCC8014) was cultivated in Man, Rogosa, and Sharpe (MRS, HiMedia®) broth to obtain a pre-inoculum with an optical density of 1.0 (approximately 1010 CFU mL−1), which was suspended in 250 mL Erlenmeyer flasks containing 150 mL of MRS broth, pH 6.8, and incubated under constant stirring (150 rpm) at 35 °C for 5 days. Two concentrations of inoculum (10% and 20% v/v) and three concentrations of glucose (20, 100, and 150 g L−1) were tested using a 2 × 3 factorial design in triplicate. At every 24 h, aliquots (10 mL) were collected and optical density was measured using a Biomate® spectrophotometer (540 nm). The aliquots were centrifuged at 8000 rpm at 4 °C for 10 min in a refrigerated centrifuge to determine titratable acidity expressed as the quantity of lactic acid (g) per 100 mL of the sample (AOAC 1995).

Fermentation of shrimp waste by L. plantarum

Shrimp waste

Fresh shrimp (Xiphopenaeus kroyeri) waste (head and shell) was collected from “Cooperativa de Indústrias Beneficiadoras” located in Santos, São Paulo (Brazil) and packaged under vacuum. The material was washed with distilled water, oven-dried at 60 °C for 24 h, freeze-dried (L101Liotop, Liobras®), crushed, sieved, and stored under frozen conditions. The waste samples were collected in triplicate for moisture, protein, and ash analysis (AOAC 1995).

Fermentation of shrimp waste in the flask and bioreactor scales

Flask scale tests were performed using previously optimized glucose (100 g L−1) and inoculum (10% v/v) concentrations, to which 5% and 10% (w/v) of the shrimp waste were added in the same incubation conditions as described in the previous section. In order to obtain a larger quantity of the product, fermentation was performed under the same conditions in a TECNAL® bench 1-L bioreactor using a 5% (w/v) waste load and a working volume of 700 mL at 190 rpm and 35 °C for 3 days. Aliquots of both flask and bioreactor fermentation batches were collected every 24 h to determine titratable acidity. After the fermentation processes were complete, the solid and liquid phases were separated by centrifugation at 5000 rpm for 10 min at 4 °C. The solid phase was washed and centrifuged three times with distilled water, oven-dried at 60 °C for 24 h, and used for determining ash content (AOAC 1995) and for deacetylation. The liquid phase was used for protein quantification by the Bradford method (Bradford 1976).

Demineralization (DM) and deproteinization (DP) degrees

The DM and DP degrees were calculated on a dry-weight basis according to Sedaghat et al. (2017).

Deacetylation of chitin

Chemical deacetylation of chitin (solid phase obtained after fermentation) was performed by the autoclave method (Hong et al. 2000) to obtain chitosan.

Characterization of chitin and chitosan

Chemical structure

The chemical structures of the bioextracted chitin and chitosan were determined by Fourier transform infrared spectroscopy (FTIR). Commercial chitosan (Polymar®, Fortaleza, Brazil) was also used for comparison. Approximately 0.2–2 mg of chitin and chitosan samples (bioextracted and commercial) was ground and mixed with 200 mg KBr. The powder mixture was compressed in a hydraulic press to form KBr discs. FTIR spectra were recorded using a Tensor II (Bruker®) spectrometer over the range of 4000–500 cm−1. Each spectrum was obtained by the accumulation of 128 scans ata resolution of 4 cm−1.

Determination of deacetylation degree (DD)

FTIR was also used to estimate the deacetylation degree of chitosan. The acetylation degree (DA) of chitosan was calculated according to the methods reported by Brugnerotto et al. (2001) and El Knidri et al. (2016).

Molecular weight determination of chitosan

First, the bioextracted chitosan was purified according to Tonhi and Plepis (2002) with modifications. The chitosan powder was added to a 3% acetic acid solution in a 1% (w/v) proportion and kept under magnetic stirring for 24 h and then centrifuged at 10,000 rpm for 40 min. The supernatant was recovered and filtered through a qualitative filter. Ammonium hydroxide was added until it reached pH 9–10. The precipitate was recovered by filtration in a quantitative filter, washed with distilled water, placed in Petri polystyrene dishes, and oven-dried (60 °C) overnight. After drying, the purified chitosan sample was dissolved in an aqueous solution of 0.3 mol L−1 acetic acid and 0.2 mol L−1 sodium acetate (pH 4.5) by mechanical stirring at 600 rpm for 24 h at 25 °C. Viscometric measurements of six chitosan solutions in the concentration range of 6.0 × 10−4–3.0 × 10−4 g mL−1 were performed at 25 °C using an Ostwald capillary viscometer (diameter of 0.75 mm) that was immersed in a cinematic bath (QUIMIS, Q–303SR26, Brazil). The molecular weight of chitosan was determined by calculating the intrinsic viscosity [η] of the polymeric solutions (Ydens et al. 2005) and using the Mark-Houwink Equation ([η] = K · Mva), where Mv is the viscometric average molecular weight and K and a are constants for a given solvent–solute system, with values of 0.074 and 0.76, respectively (Rinaudo et al. 1993).

Antimicrobial activity of chitosan

The antimicrobial activity of the bioextracted and commercial chitosan samples was examined against an industrial strain of Saccharomyces cerevisiae (PE–2) and a strain of L. fermentum (CCT0559-ATCC9338) maintained in Yeast–Peptone–Dextrose (YPD) medium (in w/v: 1% yeast extract, 2% glucose, 2% peptone, 2% agar) and in MRS medium, respectively. The slants were maintained at 4–8 °C, being periodically transferred to a new medium.

The minimum inhibitory concentration (MIC) and minimum lethal concentration (MLC) were obtained by twofold serial broth dilution in YPD or MRS broths (for yeast and bacterium, respectively) of a chitosan stock solution (prepared in a 2% acetic acid solution + culture medium in the proportion of 1:2.3, final pH 5.0) to obtain a final concentration ranging from 0.0075 to 0.96% (w/v). The final concentration of inoculum was around 2–6 × 105 CFU mL−1 prepared in the respective culture media. The tubes (final volume of 2 mL) were incubated at 150 rpm and 30 °C or 35 °C (for yeast and bacterium, respectively) for 24 h. The number of yeast and bacterial colonies was determined at the initial time (0 h) and after 24 h by plating aliquots in MRS or YPD medium using the drop-plate method (Herigstad et al. 2001). MIC was defined as the lowest concentration of chitosan at which there was no variation in the number of CFU mL−1 comparing the initial time and after 24 h. MLC99 was defined as the lowest concentration that caused a decrease of more than 99% in the number of CFU mL−1 when values at the initial time and after 24 h were compared.

A co-culture experiment using S. cerevisiae and L. fermentum (2–6 × 105 CFU mL−1 each) was performed in 10 mL sterile sugarcane juice (diluted to a concentration of approximately 4 g 100 mL–1 of total reducing sugars, pH 5.5). Three different conditions were tested: without chitosan, with 0.03% (w/v) of commercial chitosan, and with 0.96% (w/v) of bioextracted chitosan. The tubes were maintained at 30 °C and 150 rpm for 24 h. The number of bacterial and yeast colonies was determined as described above by adding chloramphenicol to a final concentration of 50 mg L−1 to YPD medium and nystatin at a final concentration of 50 mg L−1 to MRS medium, respectively.

Results and discussion

Optimization of the fermentation by L. plantarum in semi-synthetic medium to produce chitin and chitosan by chemical deacetylation

The composition of ash (54.07 ± 0.28% w/w) and protein (22.12 ± 0.45% w/w) determined using shrimp waste samples was similar to the values reported by Neves et al. (2017), which were 41% and 24%, respectively. Variations in ash and protein contents were reported by Panchakshari et al. (2016) but they may be attributed to the differences in the shrimp species and shell composition.

The highest acid production by L. plantarum in MRS medium (Fig. 1) occurred in 3 to 5 days under the conditions of 100 g L−1 glucose and 10% bacterial inoculum (Fig. 1b). These results were more favorable than those reported by Neves et al. (2017), wherein their study used higher substrate concentration (50% sucrose) and bacterial inoculum level (20–50%) but in a relatively shorter time (30 h). The increase in the inoculum concentration (from 10 to 20% v/v) did not result in an increase in acid production (Fig. 1d–f).

Fig. 1.

Fig. 1

Open in a new tab

Titratable acidity (gray bars) and optical density (solid line) in the fermentation assays by L. plantarum in MRS medium containing 20 g L–1 (a, d), 100 g L–1 (b, e) and 150 g L–1of glucose (c, f) and 10% (upper panel) and 20% (lower panel) of bacterial inoculum (v/v), at 35 °C, 160 rpm, during 5 days of fermentation

Lower waste load (5% w/v) yielded higher titratable acidity with increasing values of acidity until 4 days of fermentation (100 g L−1 glucose and 10% bacterial inoculum; Fig. 2). As expected, the DM and DP of the shrimp waste were also higher at 5% load compared to 10% (Table 1).

Fig. 2.

Fig. 2

Open in a new tab

Titratable acidity in fermentation assays by L. plantarum in MRS medium containing 100 g L–1 glucose, 10% bacterial inoculum (v/v), and 5% (black diamond) or 10% (black square) shrimp waste (w/v) at 35 °C, 160 rpm, during 4days of fermentation, in flask scale (solid line) and bioreactor scale (dashed line)

Table 1.

Demineralization (DM) and deproteinization (DP) of the shrimp residues fermented by L. plantarum to obtain chitin in flask and bioreactor scales in MRS medium containing 100 g L–1 glucose, 10% bacterial inoculum (v/v), and 5% and 10% shrimp waste (w/v) at 35 °C, under stirring, for 4 days (in flasks) or 3 days (in bioreactor) of fermentation

Fermentation scale Shrimp waste (% w/v) DM (%) DP (%)
Flask 5 48.0 33.6
10 47.1 29.0
Bioreactor 5 77.3 43.4
Open in a new tab

In the bioreactor run (100 g L−1 glucose, 10% inoculum, 5% waste, 3 days), faster acid production was observed soon after the first day but the rate became much slower on the third day. Neves et al. (2017) used a much higher shrimp waste load (33%) with higher acidity production in 72 h but they worked with cell reinoculation every 24 h with medium pH adjusted to 4.0–5.0. The DM and DP values were considerably higher in the bioreactor (Table 1) mainly due to better agitation and mixing conditions in the bioreactor vessel compared with those employed for fermentation using the Erlenmeyer flasks. Higher values for DP were obtained in bioprocesses to obtain chitin but the results varied with the bacterial species and the origin and type of fermentation (Neves et al. 2017; Sedaghat et al. 2017).

In our study, chitin was deacetylated by a chemical process. The bioextracted chitosan presented a DD of 80% while that for the commercial chitosan was 87.4% (as specified by the manufacturer). The DD is closely related to the antimicrobial effect of chitosan. The higher the DD, the more protonated amino groups will be available for electrostatic interactions with the cell surface, causing cell wall disruption and cell death (Verlee et al. 2017). Chitosan samples with a DD value of 81.2% (Zhang et al. 2017) and 87.7% (Sedaghat et al. 2017) were obtained by bacterial fermentation of shrimp waste and different deacetylation methods.

Characterization of chitin and chitosan

The FTIR spectra of chitin and chitosan (bioextracted and commercial) samples are presented in Fig. 3 and are comparable to the spectra reported in the literature (Campana-Filho et al. 2007; Liu et al. 2012; Chen et al. 2014; Sedaghat et al. 2017; Varun et al. 2017). For chitin (sample obtained after fermentation), bands representing the intermolecular bonds of hydrogen O–H (6)…O–C and O–H (3)…O–5 were observed in the range 3480–3441 cm−1. The axial deformation band associated with the hydrogen bond C = O…H–N can be visualized at 3262 and 3111 cm−1. The bands at 2931 and 2850 cm−1 resulted from the vibration of the groups CH3 and CH2. The band corresponding to amide I is divided in the wavelengths of 1655 and 1622 cm−1, characterizing the antiparallel alignment of α-chitin, which is the most stable and by far the most abundant polymorphic form of chitin (Muzzarelli 1977). Other absorption bands attributed to α-chitin are the angular deformation of N–H bond (amide II) at 1555 cm−1 and axial deformation C–N (amide III) at 1311 cm−1. The band at 1380 cm−1represents the acetamide groups and at 1160 cm−1 an axial symmetric deformation of C–O–C is observed (Fig. 3a).

Fig. 3.

Fig. 3

Open in a new tab

Fourier transform infrared spectra (FTIR) of chitin (a) obtained by fermentation of shrimp residues by L. plantarum; bioextracted chitosan (b) obtained by chemical deacetylation of chitin; and commercial chitosan (c)

In the spectrum for the bioextracted chitosan, a characteristic absorption band at 1641 cm−1 was observed due to the angular deformation of the primary amine groups. The absorption bands at 3262 and 3111 cm−1associated with the amide groups were not observed due to chitin deacetylation. The band at 3424 cm−1 showed an axial deformation of the NH2 group (Fig. 3b). Commercial chitosan exhibited a similar infrared spectrum with some variations that can be explained by the different chitin sources (Fig. 3c).

The average molecular weight of the bioextracted chitosan calculated by using the Mark–Houwink equation is approximately 4.0 × 105 g mol−1, which characterizes this biopolymer as a high-molecular-weight macromolecule (Verlee et al. 2017).With the information provided by the manufacturer regarding the commercial chitosan and according to O’Callaghan and Kerry (2016), its molecular weight is supposed to be in the range of 5.0 × 104–1.9 × 105 g mol−1, indicating a low-molecular-weight polymer.

Antimicrobial activity of chitosan

Both commercial and bioextracted chitosan samples exhibited antimicrobial activity against S. cerevisiae and L. fermentum, but a more remarkable effect was observed against the bacterium (Table 2). The yeast MIC after using the commercial chitosan was almost tenfold higher than the bacterial MIC. Moreover, bioextracted chitosan showed no yeast inhibition or lethality in the range of 0.0075–0.96% while for the bacterium, growth inhibition occurred in concentrations varying from 0.24 to 0.48% and lethality of more than 99% was seen at 0.96% while using bioextracted chitosan. These results indicate that the concentration of chitosan required to inhibit growth or kill the bacterial cells would not affect the growth and viability of the yeast. A high concentration of the bioextracted chitosan (i.e., 30 times) was necessary to cause a similar effect (MIC or MBC99) on L. fermentum compared to commercial chitosan.

Table 2.

Minimum Inhibitory Concentration (MIC) and Minimum Lethal Concentration (MLC99) for S. cerevisiae and L. fermentum in YPD and MRS media, respectively, with concentrations of commercial chitosan and bioextracted chitosan ranging from 0 to 0.96% (w/v), after 24 h

Chitosan (in % w/v) S. cerevisiae L. fermentum
MIC MLC99 MIC MLC99
Commercial 0.06 0.48 0.0075–0.015 0.03
Bioextracted * * 0.24–0.48 0.96
Open in a new tab

*No inhibition in the range of 0.0075–0.96%

The antimicrobial effect of chitosan is highly related to a combination of factors such as molecular weight, degree of deacetylation (DD), and type of microorganism. The interaction of chitosan with the bacterial cell wall is mostly electrostatic, and typically between protonated amino groups and negatively charged phospholipids, which promotes permeabilization. In Gram-positive bacteria, the presence of polyanionic teichoic acid improves the antibacterial activity of chitosan. Once it binds to the cell wall, it leads to destabilization and intracellular substance loss (Raafat et al. 2008; Kong et al. 2010; Verlee et al. 2017).

The antibacterial effect of chitosan is known to be dependent on its molecular weight (Jeon et al. 2001). However, high-molecular-weight chitosans, as in the case of the bioextracted chitosan, exhibit considerable antibacterial activity against lactic acid bacteria. A 1106 kDa-chitosan (1.1 × 106 g mol−1) inhibited the growth of L. plantarum, Lactobacillus brevis, and Lactobacillus bulgaricus at concentrations as low as 0.05–0.08% (No et al. 2002). Jeon et al. (2001) verified that L. fermentum was inhibited by a high-molecular-weight chitosan (685 kDa, or 6.85 × 105 g mol−1) at concentrations lower than 0.03%. Even ultra-high molecular weight chitosan (> 106 g mol−1) exhibits enhanced antibacterial activity (Li et al. 2016). The effects of chitosan according to different molecular weights were also elucidated in a recent study, confirming that its bactericidal effect is related to membrane disassembly (Gomes et al. 2020).

The relation between molecular weight and DD has a strong effect on the antimicrobial action of chitosan of different molecular weights against Gram-positive bacteria (Verlee et al. 2017). For a DD of 84%, the antimicrobial activity was observed with chitosan with a molecular weight of 2.3 kDa (2.3 × 103 g mol−1), whereas the antimicrobial effect of chitosan with a DD value of 52% was only verified with an 11.9 kDa (1.19 × 104 g mol−1) chitosan or higher (Mellegard et al. 2011). However, the DD values of commercial and bioextracted chitosan are comparable (87.4% and 80%, respectively), which should not have contributed to the difference between the bactericidal action of these chitosans.

The mode of action of chitosan on Gram-positive and Gram-negative bacteria is different and may be related to its molecular weight. High-molecular weight chitosan forms an impermeable layer around the cell wall by means of interactions with the cell wall compounds affecting nutrient transport. On the other hand, low-molecular-weight chitosan passes through the cell wall of Gram-negative bacteria (which is deprived of teichoic acid) and interacts with the cell membrane, thus disrupting cellular metabolism (Zheng and Zhu 2003).

Another possible explanation for the difference between the antimicrobial effects of chitosans is the fact that the bioextracted chitosan was not purified, and hence, more quantity of this chitosan would be required to have a similar effect when compared to commercial chitosan. A certain amount of chitin and carbonates may still be present in the extracted product after deacetylation, which would decrease the efficiency of chitosan (data not shown). Hence, for the use of chitosan in the bioethanol industry, the purification step would not be required. However, a comparison of the antibacterial activity between a purified and a non-purified chitosan sample would help elucidate the reason why a greater concentration of the bioextracted chitosan (30 times higher) is required to achieve a similar antibacterial effect when compared with commercial chitosan on L. fermentum. Moreover, improving the deacetylation step to increase the DD value, testing different purification methods, modifying the fermentation process such as substrate concentration, temperature, pH, etc., would contribute to achieving a more cost-effective process to obtain chitosan to be used as an antimicrobial agent in the bioethanol industry.

In co-culture fermentation using S. cerevisiae and L. fermentum in sugarcane juice, the growth of the yeast was not impaired by 0.03% commercial chitosan or 0.96% bioextracted chitosan and was quite comparable to the growth observed without chitosan use. This result is in agreement with the YPD experiment using pure culture (Table 2). The growth of L. fermentum was strongly decreased by the use of 0.03% commercial chitosan (99.9%, as in pure culture) while for the bioextracted chitosan, the decrease in growth was relatively lower (57%). It seems that the effect of the bioextracted chitosan was more inhibitory than bactericidal (as in pure culture) at the concentration of 0.96% (Fig. 4). It is worth noting that this co-culture experiment was performed using sugarcane juice and the medium composition affects the antimicrobial activity of chitosan (Hu and Gänzle 2018) but the effect was observed only on the bioextracted chitosan, which suggests again that the possible impurities in its preparation process may have interfered with the medium to decrease the antimicrobial activity of the bioextracted chitosan.

Fig. 4.

Fig. 4

Open in a new tab

Number of S. cerevisiae and L. fermentum (Lf) in co-culture experiments carried out in sugarcane juice, at 30 °C, 150 rpm, for 24 h, without chitosan, with 0.03% (w/v) of commercial chitosan and with 0.96% (w/v) of bioextracted chitosan

Three main considerations have arisen from the experiments: (1) chitosan exhibits a selective antimicrobial activity against the bacterial contaminant of bioethanol production (L. fermentum) without affecting the growth of the main agent, the yeast S. cerevisiae; (2) compared to commercial chitosan, higher concentrations of bioextracted chitosan are required to achieve similar effects on L. fermentum, again without affecting the growth of S. cerevisiae; (3) purified bioextracted chitosan may yield better results that are comparable to commercial chitosan, but it would add to the cost of the bioextraction process.

We believe that the utilization of chitosan in the bioethanol industry is prospective and suggests that the use of bioextracted chitosan may be a safer and more natural alternative to the use of sulfuric acid and/or antibiotics to control the growth of L. fermentum. Further experiments are required to optimize the use of chitosan for the ethanolic fermentation process.

Acknowledgements

This work was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo (research grant numbers 2014/17794-2, 2017/20006-4 and 2018/19139-2) and by Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (grant number PNPD/33001014051P7; finance code 001).

Author contributions

ICT was involved in the experimental design, performed the experiments and wrote the manuscript draft; LDS was involved in the experimental design, in the discussion of the results and in the edition of the manuscript; MAS and KRF were involved in the chitosan characterization and in the edition of the manuscript; SRCA was involved in the experimental design, in the discussion of the results and in the edition of the manuscript.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no conflict of interest in the publication.

Contributor Information

Isabella C. Tanganini, Email: isabella.tanganini@gmail.com

Ligianne D. Shirahigue, Email: ligianneds@yahoo.com.br

Mariana Altenhofen da Silva, Email: mariana.alt@ufscar.br.

Kelly R. Francisco, Email: kfrancisco@ufscar.br

Sandra R. Ceccato-Antonini, Email: antonini@ufscar.br

References

  1. AOAC . Official methods of analysis. Washington DC: Association of Analytical Chemists; 1995. [Google Scholar]
  2. Basso TO, Gomes FS, Lopes ML, Amorim HV, Eggleston G, Basso LC. Homo- and heterofermentative lactobacilli differently affect sugarcane-based fuel ethanol fermentation. Antonie Van Leeuwenhoek. 2014;105:169–177. doi: 10.1007/s10482-013-0063-6. [DOI] [PubMed] [Google Scholar]
  3. Beckner M, Ivey ML, Phister TG. Microbial contamination of fuel ethanol fermentations. Lett Appl Microbiol. 2011;5:387–394. doi: 10.1111/j.1472-765X.2011.03124.x. [DOI] [PubMed] [Google Scholar]
  4. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem. 1976;72:248–254. doi: 10.1016/0003-2697(76)90527-3. [DOI] [PubMed] [Google Scholar]
  5. Brugnerotto J, Lizardi J, Goycoolea FM, Argüelles-Monal J, Desbrières J, Rinaudo M. An infrared investigation in relation with chitin and chitosan characterization. Polymer. 2001;42:3569–3580. doi: 10.1016/S0032-3861(00)00713-8. [DOI] [Google Scholar]
  6. Campana-Filho SP, Britto D, Curti E, Abreu FR, Cardoso MB, Battisti MV, Sim PC, Goy RC, Signini R, Lavall RL. Extraction, structures and properties of α- and β-chitin. Quím Nova. 2007;30:644–650. doi: 10.1590/S0100-40422007000300026. [DOI] [Google Scholar]
  7. Ceccato-Antonini SR. Conventional and nonconventional strategies for controlling bacterial contamination in fuel ethanol fermentations. World J Microbiol Biotechnol. 2018;34:80. doi: 10.1007/s11274-018-2463-2. [DOI] [PubMed] [Google Scholar]
  8. Chen X, Chew SL, Kerton FM, Yan N. Direct conversion of chitin into a N-containing furan derivative. Green Chem. 2014;16:2204–2212. doi: 10.1039/c3gc42436g. [DOI] [Google Scholar]
  9. Du J, Tan E, Kim HJ, Zhang A, Bhattacharya R, Yarema KJ. Comparative evaluation of chitosan, cellulose acetate, and polyethersulfone nanofiber scaffolds for neural differentiation. Carbohydr Polym. 2014;99:483–490. doi: 10.1016/j.carbpol.2013.08.050. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. El Knidri H, El Khalfaouy R, Laajeb A, Addaou A, Lahsini A. Eco-friendly extraction and characterization of chitin and chitosan from the shrimp shell waste via microwave irradiation. Process Saf Environ Protect. 2016;104:395–405. doi: 10.1016/j.psep.2016.09.020. [DOI] [Google Scholar]
  11. Ghorbel-Bellaaj O, Hajji S, Younes I, Chaabouni M, Nasri M, Jellouli K. Optimization of chitin extraction from shrimp waste with Bacillus pumilus A1 using response surface methodology. Int J Biol Macromol. 2013;61:243–250. doi: 10.1016/j.ijbiomac.2013.07.001. [DOI] [PubMed] [Google Scholar]
  12. Gomes LP, Anjo SI, Manadas B, Coelho AV, Paschoalin VMF. Proteomic analyses reveal new insights on the antimicrobial mechanisms of chitosan biopolymers and their nanosized particles against Escherichia coli. Int J Mol Sci. 2020;21:225. doi: 10.3390/ijms21010225. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Hamed I, Ozogul F, Regenstein JM. Industrial applications of crustacean by-products (chitin, chitosan, and chitooligosaccharides): a review. Trends Food Sci Technol. 2016;48:40–50. doi: 10.1016/j.tifs.2015.11.007. [DOI] [Google Scholar]
  14. Herigstad B, Hamilton M, Heersink J. How to optimize the drop plate method for enumerating bacteria. J Microbiol Methods. 2001;44:121–129. doi: 10.1016/s0167-7012(00)00241-4. [DOI] [PubMed] [Google Scholar]
  15. Hong KN, Cho YI, Kim HR, Meyers SP. Effective deacetylation of chitin under conditions of 15 psi/121 °C. J Agric Food Chem. 2000;48:2625–2627. doi: 10.1021/jf990842l. [DOI] [PubMed] [Google Scholar]
  16. Hu Z, Gänzle MG. Challenges and opportunities related to the use of chitosan as a food preservative. J Appl Microbiol. 2018;126:1318–1331. doi: 10.1111/jam.14131. [DOI] [PubMed] [Google Scholar]
  17. Jeon YJ, Park PJ, Kim SK. Antimicrobial effect of chitooligosaccharides produced by bioreactor. Carbohydr Polym. 2001;44:71–76. doi: 10.1016/S0144-8617(00)00200-9. [DOI] [Google Scholar]
  18. Jung WJ, Jo GH, Kuk JH, Kim KY, Park RD. Extraction of chitin from red crab shell waste by cofermentation with Lactobacillus paracasei subsp. Tolerans KCTC-3074 and Serratia marcescens FS-3. Appl Microbiol Biotechnol. 2006;71:234–237. doi: 10.1007/s00253-005-0126-3. [DOI] [PubMed] [Google Scholar]
  19. Khanafari A, Marandi R, Sanatei S. Recovery of chitin and chitosan from shrimp waste by chemical and microbial methods. Iran J Environ Health Sci Eng. 2008;5:19–24. [Google Scholar]
  20. Kong M, Chen XG, Xing K, Park HJ. Antimicrobial properties of chitosan and mode of action: a state of the art review. Int J Food Microbiol. 2010;144:51–63. doi: 10.1016/j.ijfoodmicro.210.09.012. [DOI] [PubMed] [Google Scholar]
  21. Lee HW, Park YS, Jung JS, Shin WS. Chitosan oligosaccharides, dp 2–8, have prebiotic effect on the Bifidobacterium bifidium and Lactobacillus sp. Anaerobe. 2002;8:319–324. doi: 10.1016/S1075-9964(03)00030-1. [DOI] [PubMed] [Google Scholar]
  22. Li J, Wu Y, Zhao L. Antibacterial activity and mechanism of chitosan with ultra high molecular weight. Carbohydr Polym. 2016;148:200–205. doi: 10.1016/j.carbpol.2016.04.025. [DOI] [PubMed] [Google Scholar]
  23. Liu M, Bayjanov JR, Renckens B, Nauta A, Siezen RJ. The proteolytic system of lactic acid bacteria revisited: a genomic comparison. BMC Genom. 2010;11:36. doi: 10.1186/1471-2164-11-36. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Liu S, Sun J, Yu L, Zhang C, Bi J, Zhu F, Qu M, Jiang C, Yang Q. Extraction and characterization of chitin from the beetle Holotrichia parallela Motschulsky. Molecules. 2012;17:4604–4611. doi: 10.3390/molecules17044604. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Mao X, Zhang J, Kan F, Gao Y, Lan J, Zhang X, Hu Z, Li Y, Lin H. Antioxidant production and chitin recovery from shrimp head fermentation with Streptococcus thermophilus. Food Sci Biotechnol. 2013;22:1023–1032. doi: 10.1007/s10068-013-0179-5. [DOI] [Google Scholar]
  26. Marquez IG, Akuaku J, Cruz I, Cheetham J, Golshani A, Smith ML. Disruption of protein synthesis as antifungal mode of action by chitosan. Int J Food Microbiol. 2013;164:108–112. doi: 10.1016/j.ijfoodmicro.2013.03.025. [DOI] [PubMed] [Google Scholar]
  27. Mellegard H, Strand S, Christensen B, Granum P, Hardy S. Antibacterial activity of chemically defined chitosans: influence of molecular weight, degree of acetylation and test organism. Int J Food Microbiol. 2011;148:48–54. doi: 10.1016/j.ijfoodmicro.2011.04.023. [DOI] [PubMed] [Google Scholar]
  28. Muzzarelli RAA. Chitin. Oxford: Pergamon Press; 1977. [Google Scholar]
  29. Neves AC, Zanette C, Grade ST, Schaffer JV, Alves HJ, Arantes MK. Optimization of lactic fermentation for extraction of chitin from freshwater shrimp waste. Acta Sci Technol. 2017;39:125–133. doi: 10.4025/actascitechnol.v39i2.29370. [DOI] [Google Scholar]
  30. No HK, Park NY, Lee SH, Meyers SP. Antibacterial activity of chitosans and chitosan oligomers with different molecular weights. Int J Food Microbiol. 2002;74:65–72. doi: 10.1016/S0168-1605(01)00717-6. [DOI] [PubMed] [Google Scholar]
  31. O’Callaghan KAM, Kerry JP. Preparation of low- and medium-molecular weight chitosan nanoparticles and their antimicrobial evaluation against a panel of microorganisms, including cheese-derived cultures. Food Control. 2016;69:256–261. doi: 10.1016/j.foodcont.2016.05.005. [DOI] [Google Scholar]
  32. Panchakshari V, Srikanth K, Krishna PV, Babu CS. Extraction of chitin and chitosan from biowaste of scampi Macrobrichum rosenbergii and tiger shrimp Penaeus monodon. Int J Curr Microbiol Appl Sci. 2016;5:751–758. doi: 10.20546/ijcmas.2016.507.086. [DOI] [Google Scholar]
  33. Ploydee E, Chaiyanan S. Production of high viscosity chitosan from biologically purified chitin isolated by microbial fermentation and deproteinization. Int J Polym Sci. 2014;2014:1–8. doi: 10.1155/2014/162173. [DOI] [Google Scholar]
  34. Prameela K, Mohan CM, Smitha PV, Hemalatha KPJ. Bioremediation of shrimp biowaste by using natural probiotic for chitin and carotenoid production—an alternative method to hazardous chemical method. Int J Appl Biol Pharmac Technol. 2010;1:903–910. [Google Scholar]
  35. Raafat D, Sahl HG. Chitosan and its antimicrobial potential—a critical literature survey. Microb Biotechnol. 2009;2:186–201. doi: 10.1111/j.1751-7915.2008.00080.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Raafat D, von Bargen K, Haas A, Sahl HG. Insights into the mode of action of chitosan as an antibacterial compound. Appl Environ Microbiol. 2008;74:3764–3773. doi: 10.1128/AEM.00453-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Rao MS, Stevens W. Chitin production by Lactobacillus fermentation of shrimp biowaste in a drum reactor and its chemical conversion to chitosan. J Chem Technol Biotechnol. 2005;80:1080–1087. doi: 10.1002/jctb.1286. [DOI] [Google Scholar]
  38. Rinaudo M, Milas M, Dung P. Characterization of chitosan. Influence of ionic strength and degree of acetylation on chain expansion. Int J Biol Macromol. 1993;15:281–285. doi: 10.1016/0141-8130(93)90027-J. [DOI] [PubMed] [Google Scholar]
  39. Ristić T, Lasič S, Kosalec I, Bračič M, Fras-Zemljič L. The effect of chitosan nanoparticles onto Lactobacillus cells. React Funct Polym. 2015;97:56–62. doi: 10.1016/j.reactfunctpolym.2015.10.007. [DOI] [Google Scholar]
  40. Sedaghat F, Yousefzadi M, Toiserkani H, Najafipour S. Bioconversion of shrimp waste Penaeus merguiensis using lactic acid fermentation: an alternative procedure for chemical extraction of chitin and chitosan. Int J Biol Macromol. 2017;104:883–888. doi: 10.1016/j.ijbiomac.2017.06.099. [DOI] [PubMed] [Google Scholar]
  41. Sini TK, Santhosh S, Mathew PT. Study on the production of chitin and chitosan from shrimp shell by using Bacillus subtilis fermentation. Carbohydr Res. 2007;342:2423–2429. doi: 10.1016/j.carres.2007.06.028. [DOI] [PubMed] [Google Scholar]
  42. Tonhi E, Plepis AMG. Obtenção e caracterização de blendas colágeno-quitosana. Quím Nova. 2002;25:943–948. doi: 10.1590/S0100-40422002000600011. [DOI] [Google Scholar]
  43. Varun TK, Senani S, Jayapai N, Chikkerur J, Roy S, Tekulapally VB, Gautam M, Kumar N. Extraction of chitosan and its oligomers from shrimp shell waste, their characterization and antimicrobial effect. Vet World. 2017;10:170–175. doi: 10.14202/vetworld.2017.170-175. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Verlee A, Mincke S, Stevens CV. Recent developments in antibacterial and antifungal chitosan and its derivatives. Carbohydr Polym. 2017;164:268–283. doi: 10.1016/j.carbpol.2017.02.001. [DOI] [PubMed] [Google Scholar]
  45. Wahyuntari IB, Junianto SS, Setyahadi S. Process design of microbiological chitin extraction. Microbiol Indones. 2011;5:39–45. doi: 10.5454/mi.5.1.7. [DOI] [Google Scholar]
  46. Ydens I, Moins S, Degée P, Dubois P. Solution properties of well-defined 2-(dimethylamino) ethyl methacrylate-based (co) polymers: a viscometric approach. Eur Polym J. 2005;41:1502–1509. doi: 10.1016/j.eurpolymj.2005.02.002. [DOI] [Google Scholar]
  47. Zhang H, Jin Y, Deng Y, Wang D, Zhao Y. Production of chitin from shrimp shell powders using Serratia marcescens B742 and Lactobacillus plantarum ATCC 8014 successive two-step fermentation. Carbohydr Res. 2012;362:13–20. doi: 10.1016/j.carres.2012.09.011. [DOI] [PubMed] [Google Scholar]
  48. Zhang H, Yun S, Song L, Zhang Y, Zhao Y. The preparation and characterization of chitin and chitosan under large-scale submerged fermentation level using shrimp by-products as substrate. Int J Biol Macromol. 2017;96:334–339. doi: 10.1016/j.ijbiomac.2016.12.017. [DOI] [PubMed] [Google Scholar]
  49. Zheng LY, Zhu JF. Study on antimicrobial activity of chitosan with different molecular weights. Carbohydr Polym. 2003;54:527–530. doi: 10.1016/j.carbpol.2003.07.009. [DOI] [Google Scholar]

Articles from 3 Biotech are provided here courtesy of Springer

RESOURCES