Influence of detergents and sodium hypochlorite on Yarrowia lipolytica biofilms in utensils used in industrial production of colonial cheese

Abstract

The formation of microbial biofilms in materials used in the industrial production of dairy may lead to deterioration of these foods. Yarrowia lipolytica biofilms are widely found in dairy products and can modify the final characteristics of these products. Thus, this study investigated the effectiveness of hygienization by detergents and sodium hypochlorite on the formation of Y. lipolytica biofilms in different utensils usually employed during industrial cheese production, like polypropylene, hoses, and nylon/polyethylene. The utensils were sanitized using solutions of mild and alkaline detergents, and sodium hypochlorite, according to the cheese industry Standard Operation Procedure. Results showed that in all coupons there was biofilm formation with Y. lipolytica isolates. The contact angle measurements were favored to promote the adhesion of the biofilm in the evaluated surfaces. Even after treatment with sanitizers, a significant survival rate of planktonic cells was observed in all coupons tested. These results indicate that Y. lipolytica biofilms show a significant ability to adhere to polypropylene, presenting an important impact on the quality of colonial cheese.

Key words
Yarrowia lipolytica; biofilm formation; colonial cheese; hygienization; sanitizing

INTRODUCTION

Yarrowia lipolytica is a nonpathogenic yeast easily found in nature and frequent object of study due to its capability of producing metabolites with intense secretory lipolytic and proteolytic activities (Fukuda 2013). Regarded as a safe yeast, Y. lipolytica is widely employed in industry and is closely linked to dairies (Coelho et al. 2010, Jean-Marc 2012, Zinjarde 2014). This yeast is constantly associated with high proportions of fat or protein in foods and has already been identified in different types of cheese, at the surface and inside, contributing towards the process of maturation during production. Additionally, fatty acids inside Y. lipolytica throughout volatile compounds can promote undesirable effects such as organoleptic modifications, adverse effects in texture and discoloring (Groenewald et al. 2014, Zinjarde 2014).

The colonial cheese is traditionally produced by cow milk and commercial rennet. This production takes place in small industrial scale, handmade, without standardization, taking a maximum of 30 days until maturation (Borelli et al. 2006, Koelln et al. 2009, Fava et al. 2012). These operational process steps are illustrated in Figure 1. Utensils constituted of porous material are usually employed in traditional milk processing and during cheese production (Birhanu et al. 2013). For instance, PVC, hoses, beaker, spatulas, polypropylene molds and nylon/polyethylene wrapping.

Figure 1
Illustration of the production process of colonial cheese and steps where utensils are employed: (a) hose used to transfer milk, (b) beaker used to transfer the curd, (c) mold used in cheese forming and (d) package employed in the vacuum packaging process.

In the food industry, material and utensils that make up the contact surface with food exert a significant effect on the level of connection with the creation of biofilms (Van Houdt & Michiels 2009, Sokunrotanak et al. 2013). Biofilms can be defined as sessile communities of surface-attached cells encased in an extracellular matrix. This phase is followed by an intermediate state where the irreversibly attached cells form microcolonies on the surface of interest (Kiedrowski & Horswill 2011). The adherence of microorganism cells on the surfaces shows similarity to a physical-chemical process, resulting from the interaction between electrostatic forces and hydrophilic and hydrophobic interactions (Giaouris et al. 2012). In dairy production, the formation of microbial biofilm in different types of materials is very common (Galinari et al. 2014). Highly structured biofilms with sessile cells show greater resistance to antibiofilm treatment compared to planktonic cells (Bergamo et al. 2014).

When the hygienization procedure is not performed effectively, disinfectants do not penetrate through the biofilm matrix, avoiding the destruction of their living cells (Simões et al. 2010). To prevent biofilm maturation, the correct frequency of disinfection and sanitation must follow strict definitions in food processing. Moreover, the cleaning time and the type of sanitizing product are essential to prevent the formation of microbial biofilm (Van Houdt & Michiels 2009, Fouladynezhad et al. 2013). The food industry employs different categories of sanitizing products and chemical disinfectants to prevent the formation of these biofilms.

Regardless of the precise operation of the hygiene process in the industry, the materials employed during the process hold distinct characteristics that may ease up the formation of biofilms. Y. lipolytica is widely used in food industry and can have a significant impact on the final quality of the cheese. Moreover, there are few pieces of research regarding the formation of biofilms by this yeast. Thus, the aim of this study was to investigate the effectiveness of hygienization in the formation of Y. lipolytica biofilms in different utensils usually employed during industrial cheese production, like polypropylene, PVC, and nylon/polyethylene. Also, the Time-Kill Assay, Sessile Drop Method (SDM) and Emulsification Index (E24) tests were performed to evaluate the effectiveness of sanitizers in inhibiting the growth of Y. lipolytica planktonic cells.

MATERIALS AND METHODS

Microbial strains

This study employed six Y. lipolytica strains isolated from cheese marketed as colonial cheese (QU22, QU77, QU13, QU69, QU16, and QU50). Mattanna et al. (2014) performed the molecular identification through the sequencing with the domain D1/D2 of the great sub-unity (26S) ribosomal DNA using initiators NL-1 NL-4. All these isolates have 99% of sequence identity with Y. lipolytica type strains.

Biofilm formation assay on utensils

This methodology was conducted in accordance with Flach et al. (2014) and Bergamo et al. (2014) using different coupons: molds used for cheese production (polypropylene), hose (spiral PVC), beaker (polypropylene) and vacuum packaging (nylon/polyethylene), cut into shapes of 1 x 1 cm2. Before the experiments, a coupon sterilization process took place using ultraviolet radiation exposure in Biological Security Cabin Class II type A (Veco, Campinas, Brazil) for the period of 1 h.

Y. lipolytica isolates were grown on Sabouraud Dextrose Agar (SDA) with chloramphenicol (Himedia, Mumbai, India) during 24 h at 32 °C. Young cultures were added to 5 mL of Tryptone Soya Broth (TSB - Himedia, Mumbai, India) originating a suspension with 10⁶ CFU/mL, incubated at 32 °C for 24h. Then, 1 mL of this suspension was transferred for 9 mL of Peptone water 1 % (Merck, Darmstadt, Germany). Subsequently, the coupons were added to this solution and incubated for 96 h at 35 °C. Finally, the coupons were washed three times with peptone water for removal of poorly adhered cells and were added to another flask containing 50 mL of this solution. The adhered cells were released from the coupon by sonication at a frequency of 40 KHz (Unique, Indaiatuba, Brazil) for 10 min. Decimal dilutions were spread on SCA plates for assessment of microbial growth.

Evaluation of the antibiofilm activity of sanitizers in utensils

Y. lipolytica isolates were inoculated in TSB medium and incubated at 35°C for 24 h. After that, 1 mL of these cultures was added to 9 mL of sterile peptone to obtain the solution test, resulting in 106 CFU/mL. The utensils were submitted to a hygienization process, using the following sanitizing solutions according to the cheese Industry Standard Operation Procedure (SOP): mild detergent (3 %) for 5 min, alkaline detergent (6 and 8 %) for 10 min, sodium hypochlorite (1 and 1.5 %) for 10 and 20 min, followed by three washes using sterile distilled water. Then, the utensils were immediately immersed in the described fungal suspension for 96 h. Coupons were washed three times with peptone water to remove weakly adhered cells. Finally, the samples were sonicated for 10 min in 50 mL of peptone water to collect the biofilm for quantification. Decimal dilutions of sonicated peptone water were spread on SCA plates for assessment of microbial growth. This methodology followed Bergamo et al. (2014).

Time-kill assay

The Time-kill assay evaluated the sanitizing efficacy (fungicidal activity) against Y. lipolytica plantain cells according to Abreu et al. (2011), with modifications. The suspensions were prepared from isolates of Y. lipolytica containing 10⁶ CFU/mL and the following sanitizers were used: sanitizing solutions, mild detergent (3 %), alkaline detergent (6 and 8 %), sodium hypochlorite (1 and 1.5 %). The experiments were conducted using the ratio of 1.5 mL of sanitizing product to 0.5 of fungal inoculum. The contact times were 5, 10, 15, 20 and 25 min. Then, 1 mL of each fungal suspension was added to 9 mL of Peptone Water (1%) to obtain dilution 10^-3. After each contact time, 0.1 mL of this suspension was seeded in SDA and incubated for 24 h at 32 °C for determination of the number of CFU/mL.

Sessile drop method (SDM)

According to Locatelli et al. (2004), a drop with 20 µL of TSB containing a inoculum with 106 CFU/mL was carefully deposited above the coupons surfaces for later assessment of the contact angle of the drop on the surface. A Canon® Powershot SD200 digital camera captured the images showing the drops on the coupons surfaces after 5 sec of touchdown to enhance the drop surface stability. The measurement of the contact angle values occurred observing the straight-line inclination formed between the contact base radius and the height of the drop, supported by Image J software, and after three consecutive measurements (Skolodowska et al. 1999).

Emulsification index (E24)

The measurement of emulsification activity was performed according to Cooper & Goldenberg (1987). The fungal suspension of Y. Y. lipolytica in 10⁶ CFU/mL was added to 4 mL of xylene in TSB. After stabilization of the mixture, the emulsification index evaluation was performed by dividing the height of the emulsion layer by the total height of the mixture, multiplying by 100.

Statistical analysis

Statistical analysis of the results included mean ± DP and variance through ANOVA. In groups where significant statistical differences were found, the Turkey test was used along with the ’t’ test with a significance level p < 0.05. Mean values of Kolmogorov-Smirnov (KS) together with the log (CFU/cm2) results were applied to all isolates of Y. lipolytica in the study, in which all variables presented normality in its distributions. All analysis was processed by using software IBM SPSS Statistics v. 22.

RESULTS

In this study, all strains were able to form biofilms on mold, hose, beaker and wrapping (Table I). The counting of adherent cells ranged from 3.95 to 6.20 log CFU/cm2, with higher biofilm formation in the mold. Strain QU16 was the strongest biofilm formers in the mold, hose and beaker: 6.23, 5.87, 5.83 (log CFU/cm2), respectively. The strain QU50 was the strongest biofilm formers on wrapping: 6.27 log CFU/cm2. Strain QU22 was the weakest biofilm former on the mold, hose and beaker: 5.62, 3.95, 5.27 (log CFU/cm2), respectively. Strain QU69 was the weakest biofilm former on wrapping: 5.11 log CFU/cm2. However, there was no significant difference (p <0.05) between the mold and the other coupons.

Table I
Counting of sessile cells of Yarrowia lipolytica isolates adhered to coupons during the biofilm formation process.

	Log CFU/cm2*
	QU16	QU50	QU69	QU77	QU22	QU13
Mold (polypropylene)	6,23+0,03	5,89+0,09	5,73+0,02	5,72+0,01	5,62+0,06	5,68+0,58
Hose (PVC spiral)	5,87+0,12	5,78+0,17	5,60+0,05	5,02+0,05	3,95+0,35	4,95+0,06
Becker (polypropylene)	5,83+0,06	5,48+0,14	5,60+0,06	5,71+0,07	5,27+0,09	5,73+0,09
Wrapping (nylon/polyethylene)	5,30+0,18	6,27+0,02	5,11+0,06	5,20+0,04	5,53+0,14	5,45+0,02

• Yarrowia lipolytica isolates: QU16, QU50, QU69, QU77, QU22 and QU13; *Average values ± SD.

The evaluated biofilm inhibition on the utensils were statistically significant (p <0.05) among the evaluated treatments. In the mold, significant results were observed with the use of mild detergent (3%) (p = 0.002, t = 6.022, r = 0.813, IC = 0.281, 0.699) and sodium hypochlorite (1 %) for 10 min (p = 0.005, t = 4.798, r = 0.677, IC = 0.186, 0.617). The alkaline detergent (6 %) was the only one that did not present statistical significance in the results for this coupon. The beaker coupon was analyzed using only the mild detergent (3 %) in the period of 5 min and presented less statistical significance in the values comparing to the others in this study (p = 0.036, t = 2.839, r = -0.808, IC = 0.047, 0.953) (Figure 2). The hose coupons presented significant results with the sanitizing product sodium hypochlorite (1 %) in the period of 10 min (p= 0.001, t= 6.869, r= 0.843, IC= 0.688, 1.512). Sodium hypochlorite (1.5 %) in the period of 20 min presented similar results (p = 0.004, t = 5.188, r = 0.418, IC = 0.974, 2.890) (Figure 3 and Figure 4).

Figure 2
Survival of (a) QU16, (b) QU50, (c) QU69, (d) QU13, (e) QU77, (f) QU22 Yarrowia lipolytica isolates (log CFU/cm2) on the mold coupon before and after application of sanitation product (DN 3 %) mild detergent 03 - 05 min, (DA6 %) and (DA8 %) alkaline detergent and 8 % - 6% - 10 min (HS 1 %) sodium hypochlorite 1 % - 10 to 20 min. and (HS1,5 %) sodium hypochlorite 1.5% - and 10-20 min.

Figure 3
Survival of (a) QU16, (b) QU50, (c) QU69, (d) QU13, (e) QU77, (f) QU22 Yarrowia lipolytica isolates (log CFU/cm2) on the hose coupon before and after application of sanitation product (DN 3%) % mild detergent 03-05 min, (DA6 %) and (DA8 %) alkaline detergent and 8 % - 6 % - 10 min (HS1 %) sodium hypochlorite 1 % - 10 to 20 min and (HS 1,5 %) sodium hypochlorite 1.5 % - and 10-20 min.

Figure 4
Survival of QU16, QU50, QU69, QU13, QU77, QU22 Yarrowia lipolytica isolates (log CFU/cm2) on the beaker coupon before and after application of sanitizing product (DN3 %) % mild detergent 03-05 minutes.

Figure 5
Yarrowia lipolytica isolates used in the assay with mild detergent (3 %): QU16, QU50, QU69, QU77, QU22, QU13 at 5, 10, 15, 20, 25 min.

Regarding the time-kill assay, it was observed that there was no inhibition of growth in none of the tested times for the mild detergent at concentration of 3 % (Figure 4), except for isolate QU16. However, little change occurred in the course of time for the other isolates. It was also observed that there was no statistical significance (p < 0.05) in the results when comparing times 5 and 25 min (p = 0.650, t = 0.483, r = 0.650, IC = - 0.622, 0.425). However, the sanitizing products alkaline detergent 6 and 8 % and sodium hypochlorite 1 and 1.5 % were effective in inhibiting the growth of the other isolates of Y. lipolytica, in all assays.

The comparison between the angle of the water drop, Broth TSB drop and the angle of the inoculum 10⁶ CFU/mL did not present statistical significance in the results (p > 0.05). However, there were significant differences (p < 0.05) among the different coupons. With the contact angle measurements, a higher angle value of the hose coupon (overall average of 65.2° + 6.5°) demonstrated a less wetting surface property when comparing to other coupons (Table II). When relating all contact angles of the studied cultures, the coupon beaker presented significance in results (p < 0.05) compared to mold, hose and packaging, therefore presenting an enhanced wetting property. All the material culture angles presented significance in results (p < 0.05).

Table II
Sessile drops angle measurements in degrees on the coupons mold, hose and packaging.

Angle Measurements (ᶿ°)*
	Water	Broth TBS	QU16	QU50	QU69	QU77	QU22	QU13
Mold (polypropylene)	72,7+0,26	68,4+0,33	59,2+0,32	60,7+0,17	62,1+0,58	47,6+0,32	62,3+0,82	59,6+0,54
Hose (PVC spiral)	77,5+0,26	69,3+0,12	72,6+0,25	63,6+0,26	57,5+0,32	65,6+0,26	66,3+0,40	62,3+0,20
Becker (polypropylene)	60,9+0,04	61,2+0,77	49,5+0,14	48,7+0,30	52,8+0,04	34,4+0,42	34,9+0,23	52,5+0,31
Wrapping (nylon/polyethylene)	62,8+0,18	64,8+0,82	58,6+0,18	61,8+0,18	57,7+0,06	67,4+0,04	59,1+0,14	59,8+0,02

• Yarrowia lipolytica isolates: QU16, QU50, QU69, QU77, QU22, QU13; Broth TSB: Tryptic Soy Broth; *Average values ± SD.

The emulsification indexes (E24) (Table III) ranged from 49.7 to 88.3 % in isolates of Y. lipolytica and bio-emulsification was demonstrated for all tested isolates. Strain QU50 presented the lower E24 result with 49.7 % index. Consequently, it obtained a higher significance (p < 0.05) in the results when compared to the values obtained for the other isolates (QU16 p < 0.000, QU69 p < 0.000, QU22 p < 0.000, QU13 p < 0.001 e QU77 p < 0.002).

Table III
Emulsification indexes values in percentage of Y. lipolytica isolates.

Isolates	E 24 + SD
QU16	88,3 + 3,05
QU50	49,7 + 1,53
QU69	83,0 + 4,58
QU77	72,3 + 2,89
QU22	86,3 + 1,79
QU13	74,7 + 3,20

• Yarrowia lipolytica isolates: QU16, QU50, QU69, QU77, QU22 and QU13.

DISCUSSION

The strains presenting the strongest biofilm formation capacity on the coupons were from Y. lipolytica. Corroborating with our findings of Montel et al. (2014), Y. lipolytica is a biotechnologically relevant fungus capable of colonizing utensils used during the manufacture of colonial cheese. In addition, the presence of yeasts biofilms in artisanal cheese can lead to spoilage of the product, resulting in discoloration, gas production, undesirable flavor and changes in texture, as reported by Galinari et al. (2014). An investigation was performed to study biofilm formation on these materials, like stainless steel, rubber, silicon, glass, plastic, wooden surfaces and milking equipment. Molds, yeasts, and bacteria are the dominant microorganisms in this segment with stable formation of biofilms for an extended period (Montel et al. 2014). Biofilm formation by yeast can act in farm cheese in two ways, inducing product deterioration, creating undesirable flavor and discoloring the final product or can generate beneficial effect through proteolytic and lipolytic enzymes flavor enhancing during maturation.

Brugnoni et al. (2012) and Rosa et al. (2015) obtained yeast cell counting with results higher than 6.0 log (CFU/cm²) and 7.0 log (CFU/cm²) respectively. They also observed equal values between the counts in which there was no significant difference (p > 0.05), results that match our findings of 6.27 and 6.23, for instance, when coupons’ surface adhesion was evaluated. It is also clear that the strain of this yeast presents itself in biofilm growing mode under certain conditions. The same strain is associated with food deterioration, including cheese varieties that develop the tyrosine-processing capability, promoting the change in its coloration (Zinjarde 2014). Galinari et al. (2014) tested biofilm formation with yeasts in wooden utensils employed in cheese production and observed lower resulting values when compared to the results of this study.

Many studies reported the diversity of biofilm-forming microorganisms isolated from diverse areas of the food industry. In contrast to what was presented in our study, we can relate bacteria adherence with significant results found by Beltrame et al. (2014) and Santos Junior et al. (2014), respectively, in which maximum counts of 6.92 log (CFU/cm²) were observed in polyethylene coupons and, 6.17 log (CFU/cm²) in solid polypropylene surfaces.

The production of cheese marketed as colonial also uses several sanitizing products employed in the food industry. In these areas, sanitation takes place by using the sanitizing products tested in our experiments, varying only the concentration and the amount of time the product is applied. Figures 2, 3 and 4 demonstrate survival rate of isolates of Y. lipolytica in all coupons. Based on these results, we can notice that the mold coupon was the most adherence favorable material, followed by beaker and hose coupons, respectively. The sanitizing product alkaline detergent had the better efficiency (Figure 3) in hose coupons with QU22 strain. Also in this coupon, the effect of sanitizer sodium hypochlorite showed higher reduction of biofilm. The coupon received treatment in the concentration of 1 and 1.5% (Figure 3) in the period of 10 and 20 minutes.

According to Van Houdt & Michiels (2009) and Mogotsi et al. (2014), among the many sanitizing products available, active chlorine is probably the most used compound, and sodium hypochlorite proved to be an active oxidizing agent that can destroy protein cell activity. However, penetration only happens completely when they are in a de-ionized state. Moreover, for better disinfection and efficacy, cleaning agents such as detergents appear in combination with a chlorine-based solution.

In contrast, if we compare the sanitizing products’ effect on strains of Y. lipolytica there are no data regarding resistance of its adherence cells. Even so, there are other studies describing the action of disinfectants on yeast. Sodium hypochlorite and 70 % alcohol obtained effective action against a mixture of planktonic yeasts (Théraud et al. 2004). This solution reduced the adhesion of all strains of Candida albicans. For the majority of other species of Candida (non-albicans), the increase in the rate of blastospores against hyphae on polystyrene did not show cause-effect over the production or in the proteinase enzyme activity (Webb et al. 2007). Biofilms formed by strains of C. albicans were eradicated when exposed to sodium hypochlorite for 30 minutes in concentrations of 1:32 or higher (Dahlan et al. 2011). Finally, IIknur et al. (2012) demonstrated biofilm reduction against the control group in species of Candida exposed to polystyrene. However, no tested disinfectant completely removed the biofilm.

The time-kill curve was used to determine the fungicidal activity of sanitizing products on isolates of Y. lipolytica. This study demonstrates results similar to those presented by Brugnoni et al. (2012), using the sanitizing products hypochlorite against strains of yeast C. krusei, Zygosaccharomyces sp., K. marxianus and R. mucilaginosa. It was also noticed that for planktonic cells the reduction happened by using a lower concentration of sodium hypochlorite (0.02 %) in all tested strains. However, in studies with Gram-negative bacteria, lower effectiveness is noticeable in comparison to our study with yeasts. When using chlorine based sanitizing, the time-kill curve showed that an average of 83 min of action of sodium hypochlorite (0.02 %) (Sukplang & Thongmme 2014) and 60 min of action of sodium hypochlorite (0.05 %) is needed for an effective sanitizing to take place. (Mazolla et al. 2006).

The contact angle of the sessile drop served as a method to characterize the hydrophobicity/hydrophilicity of the surfaces. The relation between hydrophobicity and biofilm formation, and the correlation among them are, in most cases, clear and with physical-chemical surface properties regulating the initial adhesion of microorganisms. Thus, the hydrophobic characteristics of the biofilm make its adhesive properties attach easily to the surfaces of material (Tarifa et al. 2013, Cappitelli et al. 2014). The free surface energy defined by the roughness of the material can also influence the formation of biofilm (Flausino et al. 2014). Lehocký et al. (2007) emphasize in their study that yeast cells play a significant role in adhesion as well as the substrate’s surface. According to this concept, Y. lipolytica is an yeast capable of connecting only to very hydrophobic surfaces. As in our study, other results were favorable regarding the angle measurements with yeasts, promoting microbial adhesion. Gole et al. (2002) observed hydrophobic regions with contact angles of drops measured up to 105° in Y. lipolytica at the tested material surface. Gallardo-Moreno et al. (2004) with strains of Candida parapsilosis with measurements from 15° to 92°, also highlighted the contact angle measurements. The hydrophobicity of yeasts positively related to the adhesion rate of the tested material and with different levels of biofilm formation (Tarifa et al. 2013).

Many studies have demonstrated the formation of emulsification. Fontes et al. (2012) reported that strains of Y. lipolytica presented emulsification indices up to 68.0 and 70.2 %. Souza et al. (2012) also achieved good emulsifier production in their results with Y. lipolytica in the presence of seawater. To improve the production of biosurfactants, Fontes et al. (2010) achieved a better result of emulsification index (67.7 %) in a blend media with ammonium sulfate and yeast extract. Emulsification activity was also detected with Y. lipolytica in culture media in the study performed by Amaral et al. (2006).

Most emulsified hydrocarbons degrading microorganisms produce biosurfactants. The cell surface hydrophobicity is also an important aspect of the microbial cell adhesion to surfaces. Therefore, there is direct correlation between hydrophobicity, biosurfactant production, and microbial adherence (Youssef et al. 2004, Coimbra et al. 2009). All the isolates, excluding QU50 isolate, displayed relatively significant emulsification capabilities, with rates above 70%.

Groenewald et al. (2014) describe that, due to its lipolytic and proteolytic activities, Y. lipolytica strains has been widely employed in maturation or contributed to organoleptic characteristics, although it does trigger deterioration in some types of cheese. These quality altering effects include non- standard flavor, undesirable texture, surface browning and biogenic amine formation that contributes to product decomposition.

There are few studies on biofilm formation by Y. lipolytica. The results showed that Y. lipolytica isolates from colonial cheese shows significant ability to adhere to polypropylene. However, tests with sanitizers were not able to inactivate all adherent cells. Hydrophilic capability and bioemulsifier production were observed within these isolates.

ACKNOWLEGMENTS

The authors are grateful to the Brazilian agencies Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq, Public Notice Universal MCTI/ CNPq Number 14/2013).

REFERENCES

• ABREU CS, LOURENÇO FR & PINTO TJA. 2011. The efficacy and stability of five sanitizing agents challenged with reference microorganisms and clean area isolates. J Basic Appl Sci 32: 201-209.
• AMARAL PFF, SILVA JM, LEHOCKY M, BARROS-TIMMONS AMV, COELHO MAZ, MARRUCHO IM & COUTINHO JAP. 2006. Production and characterization of a bioemulsifier from Yarrowia lipolytica. Process Biochemistry 41: 1894-1898.
• BELTRAME CA, MARTELO EB, MESQUITA JB, STEFFENS C, TONIAZZO G, VALDUGA E & CASIAN RL. 2014. Adhesion of Listeria monocytogenes to cutting board surfaces and removal by different sanitizers J Verbrauch Lebensm 10: 41-47.
• BERGAMO VZ, DONATO DK, DALLA LANA DF, ORTEGA GG, SCHREKKER HS & FUENTEFRIA AM. 2014. Imidazolium salt as antifungal agents: strong antibiofilm activity against multidrug resistant Candida tropicalis isolates. Lett Appl Microbiol 60: 66-71.
• BIRHANU S, TOLEMARIAM T & TOLOSA T. 2013. Microbiological quality of ayib, traditional Ethiopian cottage cheese, in Jimma area, south-west Ethiopian. Afr J Microbiol Res 32: 4169-4175.
• BORELLI BM, FERREIRA EG, LACERDA ICA, SANTOS DA, CARMO LS, DIAS SR, SILVA MC & ROSA CA. 2006. Enteroxigenic Staphylococcus spp. and other microbial contaminants during production of Canastra cheese, Brazil. Braz J Microbiol 37: 545-550.
• BRUGNONI LI, LOZANO JE & CUBITTO MA. 2012. Efficacy of sodium hypochlorite and quartenary ammonium compounds on Yeats isolated from apple juice. J Food Process. Eng 35: 104-119.
• CAPPITELLI F, POLO A & VILLA F. 2014. Biofilm formation in food processing environments is still poorly understood and controlled. Food Eng Rev 6: 29-42.
• COELHO MAZ, AMARAL PFF & BELO I. 2010. Yarrowia lipolytica: an industrial workhorse A. Mendez-Vilas, v. 2. Formatex, p. 930-944.
• COIMBRA CD, RUFINO RD, TUNA JM & SARUBBO LA. 2009. Studies of the cell surface proprieties of Candida species and relation to the production of biossurfactants for environmental applications. Curr. Microbiol 58: 245-251.
• COOPER DG & GOLDENBERG BG. 1987. Surface-active agents two Bacillus species. Appl Environ Microbiol 53: 224-229.
• DAHLAN AA, HAVEMAN CW, RAMAGE G, LOPEZ-RIBOT JL & REDDING SW. 2011. Sodium hypochlorite, chlorhexidine gluconate, and commercial denture cleansers as disinfecting agents against Candida albicans: An in vitro comparison study. Gen Dent 59: 224-229.
• FAVA LW, HERNANDES JF, PINTO AT & SCHMIDT V. 2012. Characteristics of colonial hand- made cheese sold in an agricultural show. Acta Sci Vet 40: 1080-1084.
• FLACH J, GRZYBOWSKI V, TONIAZZO G & CORÇÃO G. 2014. Adhesion and production of degrading enzyme by bacterial isolated from biofilm in raw milk cooling tanks. Food Sci Technol Int 34: 571-576.
• FLAUSINO JS, SOARES PBF, CARVALHO VF, MAGALHÃES D, SILVA, WM, COSTA HL & SOARES CJ. 2014. Biofilm formation on different materials for tooth restoration: analysis of surface characteristics. J Mater Sci 49: 6820-6829.
• FONTES GC, AMARAL PF, NELE M & COELHO MA. 2010. Factorial desing to optimize biosurfactant by Yarrowia lipolytica. J Biomed Biotechnol 10: 1-8.
• FONTES GC, RAMOS NM, AMARAL PFF, NELE M & COELHO MAZ. 2012. Renewable resources for biosurfactant production by Yarrowia lipolytica. Braz J Chem Eng 29: 483-493.
• FOULADYNEZHAD N, ASFSAH-HEJRI, L, RUKAYADI Y, ABDULKARIM SM, MARIAN MN & SON R. 2013. Assessing biofilm formation by Listeria monocytogenes. Int Food Res J 20: 987-990.
• FUKUDA R. 2013. Metabolism of hydrophobic carbon sources and regulation of it in n-alkane- assimilating yeast Yarrowia lipolytica. Biosci Biotechnol Biochem 77: 1149-1154.
• GALINARI E, DA NOBREGA JE, DE ANDRADE NJ & DE LUCES CL. 2014. Microbiological aspects of the biofilm on wooden utensils used to make a Brazilian artisanal cheese. Braz. J. Microbiol 45: 713-720.
• GALLARDO-MORENO AM, GONZALEZ-MARTIN ML, PEREZ-GIRALDO C, BRUQUE JM & GOMEZ-GARCIA AC. 2004. The measurement temperature: An important factor relating physicochemical and adhesive properties of yeast cells to biomaterials. J Colloid Interface Sci 271: 351-358.
• GIAOURIS E, CHORIANOPOULOS N, SKANDAMIS P & NYCHAS G. 2012. Attachment and biofilm formation by Salmonella in food processing environments. Salmonella A Dangerous Foodbome Pathogen. Intech 8: 157-179.
• GOLE A, DIXIT V, LALA N, SAINKAR SR, PANTI A & SASTRY M. 2002. Patterned assembly of Yarrowia lipolytica yeast cells onto thermally evaporated octadecylamine films. Colloids Surf B 25: 363-368.
• GROENEWALD M, BOEKHOUT T, NEUVÉGLISE C, GAILLARDIN C, VAN DIJCK PW &WYSS M. 2014. Yarrowia lipolytica: safety assessment of an oleaginous yeast with a great industrial potential. Crit Rev Microbiol 40: 187-206.
• IIKNUR D, YASEMIN O & NURI K. 2012. Effect of disinfectants on biofilm development by five species of Candida. Afr J Microbiol Res 6: 2380-2386.
• JEAN-MARC N. 2012. Yarrowia lipolytica. Yeast 29: 409-418.
• KIEDROWSKI MR & HORSWILL AR 2011. New approaches for treating staphylococcal biofilm infections. Ann N Y Acad Sci 1241: 104-121.
• KOELLN FTS, MATTANA A & HERMES E. 2009. Microbiological evaluation of mozzarella cheese kind and colonial cheese marketed in the west region of Paraná. Rev Bras Tec Agro 3: 66-74.
• LEHOCKÝ M, AMARAL PFF, ST’AHEL P, COELHO MAZ, BARROS-TIMMONS AM & COUTINHO JAP. 2007. Preparation and characterization of orhanosilicon thin films for selective adhesion of Yarrowia lipolytica yeast cells. J Chem Technol Biotechnol 82: 360-366.
• LOCATELLI CI, ENGLERT GE, KWITKO S & SIMONETTI AB. 2004. In vitro bacterial adherence to silicone and polymetylmethacrylate intraocular lenses. Arq Bras Oftalmol 62: 241-248.
• MATTANNA P ET AL. 2014. Lipid profile and antimicrobial activity of microbial oils from 16 oleaginous yeasts isolated from artisanal cheese. Rev Bras Biocienc 12: 121-126.
• MAZOLLA PG, MARTINS AMS & PENNA TCV. 2006. Chemical resistance of the gram-negative bacteria to different sanitizers in a water purification system. BMC Infect Dis 131: 1-11.
• MONTEL M, BUCHIN S, MALLET A, DELBES-PAUS C, VUITTON DA, DESMASURES N & BERTHIER F. 2014. Traditional cheeses: rich and diverse microbiota with associated benefits. Int J Food Microbiol 117: 136-154.
• MOGOTSI L, SMIDT O, VENTER P & GROENEWALD W. 2014. Influence of sanitizer on the lipopolysaccharide toxicity of Escherichia coli strains cultivated in the presence of Zygosaccharomyces bailii. Scientific World J 1: 1-6.
• ROSA HS, CAMARGO VB, CAMARGO G, GARCIA C, FUENTEFRIA AM & MENDEZ ASL. 2015. Ecdysteroids in Sida tuberculata R.E. Fries (Malvaceae): Chemical composition by LC–ESI-MS and selective anti-Candida krusei activity. Food Chem 182: 193-199.
• SANTOS JUNIOR AC, SALIMENA APS, CARDOSO MG, ALVES E & PICCOLI RH. 2014. Action of sanitizers on Staphylococcus aureus biofilm on stainless steel and polypropylene surfaces. Afr J Microbiol Res 36: 3347-3353.
• SIMÕES M, SIMÕES L & VIEIRA M. 2010. A review of current and emergent biofilm control strategies. LWT, Food Sci Technol Res 43: 573-583.
• SKOLODOWSKA A, WOZNIAK M & MATLAKOWSKA R. 1999. The method contact angle measurements and estimation of work of adhesion in bioleaching of metals. Biol Proced Online 1: 114-121.
• SOKUNROTANAK S, IQBAL KJ & SANG-DO H. 2013. Biofilm formation in food industries: a food safety concern. Food Control 31: 572-585.
• SOUZA FA, SALGUEIRO AA & ALBURQUERQUE CDC. 2012. Production of bioemulsifiers by Yarrowia lipolytica in sea water using diesel oil as the carbono source. Braz J Chem Eng Brazilian 29: 61-67.
• SUKPLANG P & THONGMEE A. 2014. In vitro kill-time test of disinfectants against Pseudomonas aeruginosa recovered from water associated with hemodialysis applications. RJAS 4: 39-45.
• TARIFA MC, BRUGNONI LI & LOZANO JE. 2013. Role of hydrophobicity in adhesion of wild yeast isolated from the ultrafiltration membranes of an apple juice processing plant. Biofouling 29: 841-853.
• THÉRAUD M, BÉDOUIN Y, GUIGUEN C & GANGNEUX JP. 2004. Efficacy of antiseptics and disinfectants on clinical and environmental yeast isolates in planktonic and biofilm conditions. J Med Microbiol 53: 1013-1018.
• VAN HOUDT & MICHIELS CW. 2009. Biofilm formation and the food industry, a focus on the bacterial outer surface. J Appl Microbiol 109: 1117-1131.
• WEBB BC, WILLCOX MD, THOMAS CJ, HARTY DW & KNOX KW. 2007. The effect of sodium hypochlorite on potential pathogenic traits of Candida albicans and other Candida species. Oral Microbiol Immunol 53: 1013-1018.
• YOUSSEF NH, DUNCAN KE, NAGLE DP, SAVAGE KN, KNAPP RM & MCINERNEY MJ. 2004. Comparison of methods to detect biosurfactant production by diverse microorganisms. J Microbiol Methods 56: 339-347.
• ZINJARDE SS. 2014. Food-related application of Yarrowia lipolytica. Food Chem 152: 1-1.

Publication Dates

• Publication in this collection
  31 July 2020
• Date of issue
  2020

History

• Received
  20 Dec 2018
• Accepted
  7 Mar 2019