**The E**ff**ect of Eugenol and Chitosan Concentration on the Encapsulation of Eugenol Using Whey Protein–Maltodextrin Conjugates**

### **Iceu Agustinisari 1,2, Kamarza Mulia <sup>1</sup> and Mohammad Nasikin 1,\***


Received: 8 April 2020; Accepted: 29 April 2020; Published: 4 May 2020

### **Featured Application: The spray-dried eugenol formulation can be used as a food preservative preparation or for pharmaceutical purposes. The encapsulation formulation and method can be applied to other essential oils.**

**Abstract:** Eugenol has many functional properties for food and pharmaceutical purposes, especially as an antimicrobial agent. However, its use is constrained by its volatility and shelf life because it is easily degraded due to temperature, oxidation, and light. Research on encapsulation technology using biopolymers is still required to obtain the appropriate formulation in a eugenol delivery system. The aims of this research were to develop a new formulation of protein and polysaccharides in eugenol encapsulation and to determine the effect of eugenol and chitosan concentration on the characteristics of the emulsions and spray-dried powder produced. In this study, eugenol was encapsulated in whey protein–maltodextrin conjugates and chitosan through the double layer encapsulation method. The emulsions which were prepared with 2.0% eugenol were relatively more stable than those of 1.0% eugenol based on the polydispersity index and zeta potential values. Spray-dried powder which was prepared using an emulsion of 2.0% *w*/*w* eugenol and 0.33% *w*/*w* chitosan had the highest eugenol loading. The presence of chitosan resulted in more stable emulsions based on their zeta potential values, improved thermal stability of eugenol, increased eugenol loading to become twice as much as the loading obtained without chitosan, and modified release profile of eugenol from the spray-dried powders.

**Keywords:** eugenol; encapsulation; whey protein–maltodextrin conjugates; chitosan

### **1. Introduction**

Eugenol (C10H12O2) which is a phenylpropanoid group consisting of an allyl chain-substituted by guaiacol is a major bioactive compound that has a concentration of 45–90% in clove. This essential oil has shown many pharmacological uses due to its antibacterial, antifungal, antiplasmodial, antivirus, anthelmintic, anti-inflammatory, analgesic, and antioxidant activities [1]. However, eugenol has limited applications, because of its volatility, slight solubility in water, and ease of damage when exposed to high temperatures, air, and light. Nevertheless, encapsulation technology has been used to overcome these problems and to improve the utilization of essential oils as pharmaceuticals and food ingredients. Some benefits of encapsulating essential oils include increasing stability, protecting active compounds from interaction with other ingredients, increasing the activity or functional properties, and decreasing

volatility and toxicity [2,3]. In addition, encapsulation protects active compounds from oxidation, masks flavor, allows controlled release, and increases bioavailability and efficacy [4–6].

The benefit of encapsulation is achieved by using an appropriate polymer matrix as a wall material. Some factors affecting the retention and release of active compounds from the film include the type of polymer, the preparation method, the interaction between polymeric and active compounds, and the environment [7,8]. Generally, protein and carbohydrates are good combinations for wall materials for encapsulation [9]. Whey protein (WP) as a protein and maltodextrin (MD) as a polysaccharide are frequently used as a matrix in essential oil encapsulation, emulsifying agents, and stabilizers [10,11]. Moreover, protein-conjugated polysaccharides are better emulsifiers than proteins or polysaccharides alone. The difference between the present work and previous studies lies in the type of whey protein used. The whey protein used in this study was derived from bovine milk and had a protein content of 7–11%, whereas the isolates had a protein content of more than 90%.

No research has been conducted on double-layer eugenol encapsulation using a whey protein–maltodextrin (WPMD) conjugate as the first layer and the effect of chitosan as the second layer. Generally, double-layer encapsulation is formed using layer-by-layer electrostatic deposition of polyelectrolytes on oppositely charged surfaces and is processed by emulsification and homogenization [12,13]. The advantages of a double-layer system are better stability against environmental stress than a single layer [14] and the controlled release of active compounds [15].

Chitosan was chosen as the second layer in consideration of its characteristics and the findings of some related research. Previous studies have confirmed that a multilayer oil-in-water emulsion containing lecithin, chitosan, and pectin made through the layer-by-layer method exhibited good stability to aggregation [16]. Several studies on the effectiveness of chitosan as an encapsulating agent and its profile release have been carried out by some researchers [17–19]. Furthermore, according to Estevinho et al. [20], chitosan has the ability to form ionic or covalent bonds with crosslinking agents and form a network in which the active compounds can be maintained. These properties are highly advantageous and necessary for controlling the release of active compounds.

The objectives of this study were (1) to develop a new formulation of biopolymers using the double-layer method for eugenol encapsulation and (2) to investigate the effect of different eugenol and chitosan concentrations on emulsification, encapsulation performance, and the physical properties of spray-dried powders. In addition, the kinetics and mechanism of eugenol release from microcapsules were also evaluated.

#### **2. Materials and Methods**

#### *2.1. Materials*

WP from bovine milk (W1500) was obtained from Sigma-Aldrich, and technical MD was obtained from PT Sarana Mitra Anugrah (Bogor, Indonesia). Food-grade chitosan (88.59% degree of diacetylation) was purchased from Biotech Surindo (Cirebon, West Java, Indonesia), sodium acetate was purchased from Merck (127-09-4|106268), and eugenol (W246700) was obtained from Sigma-Aldrich.

#### *2.2. Preparation of Whey Protein–Maltodextrin (WPMD) Conjugates*

The preparation of WPMD conjugates was adopted from Shah et al. and Akhtar and Dickinson [11,21] with modifications. Whey protein was dissolved at 40 g/1000 g in distilled water and stirred for 15 min to homogenize the solution. The maltodextrin (80 g) was dissolved in the whey protein solution, which was, then, hydrated up to 18 h at room temperature (28–30 ◦C). The samples were spray-dried at 150 ◦C inlet temperature, 0.4 kPa compressed air, and 52 m<sup>3</sup> /h air flow rate using a spray dryer (Lab Plant Spray Dryer SD05). The outlet temperature was recorded at 70–80 ◦C, then, the spray-dried powders were heated at 90 ◦C in a cabinet oven for 1.5 h, which was then continued at 110 ◦C for 1.5 h.

#### *2.3. Encapsulation of Eugenol by Emulsification and Spray Drying*

Emulsification to encapsulate eugenol in the WPMD conjugate and chitosan as the wall material was adopted from the method reported by Shah et al. and Preetz et al. [11,12] with modifications. The WPMD conjugate product (5% *w*/*w*) was dissolved in distilled water and stirred using a magnetic stirrer at 700 rpm for 15 min to homogenize the solution. The conjugate solution had two functions, i.e., an emulsifier to emulsify the essential oil and a first layer in the double-layer system. In the first step, the emulsification was prepared using a high-shear homogenizer (Ultraturrax homogenizer, IKA T25, Germany). The conjugate solution was homogenized at 15,000 rpm for 1 min to produce the primary emulsion. This was achieved by adding eugenol of 2.0% or 1.0% *w*/*w* of the WPMD conjugate solution mass into the solution of WPMD conjugates under stirring at 15,000 rpm. Chitosan solutions (0.0%, 0.067%, 0.2%, and 1.0% in 0.1 M acetate buffer, pH 4.5) were gradually added, separately, to the primary emulsion according to the treatments that were determined. The addition was carried out under stirring at 15,000 rpm for 3 min to obtain the secondary emulsion. The mass ratio between the eugenol primary emulsion and chitosan solution was 2:1. The secondary emulsion was treated with a high-pressure homogenizer (Panda 2000, Gea Niro Soavi) for five cycles at 400–450 bar. There were eight emulsion samples consisting of four chitosan concentrations in each of two eugenol concentrations. The final emulsions had eugenol concentrations of 2.0% and 1.0% and were coded as F1 and F2, respectively. The chitosan concentrations in the final emulsion were 0.0%, 0.067%, 0.20%, and 0.33%. All emulsion samples were spray-dried at 150 ◦C inlet temperature to obtain dried powder samples. The formulation and the name of the samples is described in Table 1.


**Table 1.** The formulation of emulsion and name of the samples obtained.

#### *2.4. Encapsulation Performance*

Total eugenol was determined using the method of Shah et al. [11] with modifications. The spray-dried powder (40 mg) was dissolved in 10 mL of 60% methanol by sonication for 30 min. The samples were filtered through a membrane with a 0.45 µm pore size and the injection volume was 20 µL. Analyses were performed using HPLC equipped with an RS diode array detector at 280 nm. The mobile phase consisted of 60 mL/100 mL aqueous methanol using an isocratic mode at a flow rate of 1 mL/min in a C-18 column (Agilent, Zorbax Eclipse Plus; 4.6 mm × 25 cm, 5 µm, 100 Å). The eugenol concentration was determined from the sample peak area and used in the calculation of the loading capacity (LC). Encapsulation efficiency (EE) is defined as the percentage of total eugenol mass in spray-dried product with reference to the corresponding mass of eugenol in feed. The calculation of mass of eugenol in feed involved non-solvent mass in emulsion and mass of collected product.

$$\text{Loading (g/100g)} = \frac{\text{Mass of euogenol}}{\text{Mass of collected product}} \times 100\tag{1}$$

$$\text{Mass of equogenol in feed } (\text{g}) = \frac{\text{mass of eugenol in emulsion}}{\text{non solvent mass in emulsion}} \times \text{mass of collected product} \tag{2}$$

$$\text{Encapsulation efficiency } (\text{g}/100 \text{g}) = \frac{\text{Mass of unequal}}{\text{mass of unequal in feed}} \times 100 \tag{3}$$

#### *2.5. Particle Size and Zeta Potential Measurement of Emulsion and Spray-Dried Powder Dispersion*

The particle size and zeta potential of the emulsion and the dispersion of dried powder were measured using a Zetasizer Nano ZS (Malvern Instrument, Malvern, UK). The measurement principle for particle size was based on dynamic light scattering, while zeta potential was determined using laser doppler micro electrophoresis. The light source is He-Ne laser and a detector angle of 173◦ . Three drops (0.07 g) of emulsion samples were diluted in 20 mL of distillation water. The particle dimension of the spray-dried powder was measured by dispersing the powder (1.33 mg) in distilled water (20 mL), followed by sonication (Branson 3510) for 5 min. The dispersion was poured into the disposable plastic micro cuvette for particle size and disposable folded capillary cell (DTS 1070), for zeta potential measurement. The particle size measurement results showed the mean of a droplet diameter (z-average) and polydispersity index (PdI) value. Each data value was an average of three measurements.

#### *2.6. Di*ff*erential Scanning Calorimetry (DSC)*

The thermal denaturation properties of eugenol-encapsulated microparticles using the WPMD conjugate and chitosan as wall materials were analyzed with a DSC 8000 Perkin Elmer. The samples were weighed to be approximately 7–10 mg and prepared in aluminum pans. The samples were heated from 40 to 300 ◦C at a rate of 10.00 ◦C/min. The thermal transitions were evaluated in terms of peak transition temperature (Tp) and enthalpy (∆H). On the basis of the International Confederation for Thermal Analysis and Calorimetry (ICTAC) standard, the peak value in DSC thermograms is known as the melting point in polymer samples https://www.perkinelmer.com/CMSResources/Images/44- 74542GDE\_DSCBeginnersGuide.pdf.

#### *2.7. Fourier Transform Infrared (FTIR)*

All infrared spectra of eugenol microcapsules were obtained using a Nicolet iS50 FTIR spectrometer (Thermo Scientific). The instrument was equipped with a KBr beam splitter and a DTGS KBr detector. The scanning process was carried out at a resolution of 2 cm−<sup>1</sup> with a frequency range of 400 and 4000 cm−<sup>1</sup> .

#### *2.8. Study of In Vitro Release of Eugenol and Release Kinetic*

The in vitro release test of eugenol followed the method used by Hosseini et al. [22] and Chen et al. [23]. The sample (40 mg) was placed in an Erlenmeyer flask containing 10 mL of 60% phosphate-buffered saline (pH 7.4) and 40% ethanol. Agitation was carried out using a shaker incubator at a temperature of 27–30 ◦C with a speed of 75 rpm. At the appointed time intervals, 5 mL of the sample was sucked out for analysis and replaced with 5 mL of the buffer phosphate solution. The sample was transferred into a tube for further filtration using 0.45 µm PVDF and put into a 2 mL vial for the measurement process by HPLC. The cumulative percentage of the amount of active compound/essential oil released from the spray-dried powder was obtained by dividing the cumulative amount of essential oil released at any given time interval (Mt) by the initial weight of the encapsulated essential oil (M0) (Equation (3)):

$$\text{Release Cumulative Percentage} = \sum\_{t=0}^{t} \frac{Mt}{M0} \times 100\% \tag{4}$$

$$\frac{Mt}{M0} = \text{kt}^{\text{n}}\tag{5}$$

The release mechanism and release kinetics of eugenol from the spray-dried powder were investigated using the Korsmeyer–Peppas kinetic model [24] (Equation (4)). Mt is the amount of eugenol released at a given time, M0 is the initial amount of eugenol contained in the spray-dried powder or the maximum amount of eugenol that can be released from the spray-dried powder, t is the release time, k is the kinetic constant for the system, and n is the release characteristic and determines the release mechanism.

#### *2.9. Statistical Analysis*

The data analyses were carried out with the MINITAB 19 software using analysis of variance (ANOVA), and differences were considered significant at α < 0.05.

#### **3. Results and Discussion**

#### *3.1. E*ff*ect of Chitosan and Eugenol Concentration on Eugenol Emulsions*

The data showed that the droplet size of F1 emulsions tended to increase along with the increasing concentration of chitosan solution as the second layer in the eugenol encapsulation (Table 2). This result is in line with that of Lertsurthiwong and Rojsitthisak [25], who investigated oil encapsulation using chitosan and alginate as a biopolymer. The increased droplet size indicated polymer attachment on the surface of the oil core. However, the eugenol emulsion F2 did not show the same phenomenon. The results of the ANOVA test indicated that the increase in droplet size, due to the chitosan concentration, was not significantly different. Furthermore, the Tukey's test results showed that there were significant differences in droplet size between F2-chi 0.0% and F2-chi 0.067%.

**Table 2.** Droplet size, polydispersity index, and zeta potential of emulsions.


(\*) F1 and F2 contained 2.0% and 1.0% eugenol, respectively. Data represent mean ± standard deviation. Different letters indicate significant differences among samples (*p* < 0.05).

The eugenol emulsion of F2-chi 0.067% had the largest particle size and was significantly different from the other samples. This was probably due to the instability of the emulsion, which eased the aggregation. The instability of the emulsion was also shown by its PdI and zeta potential values. The PdI of F2-chi 0.067% was the highest among all the samples, whereas the zeta potential value was the lowest (Table 2). A high PdI value indicates instability of an emulsion [26].

Overall, the data showed that F1 emulsions had PdI values closer to 0.2, which represented the homogeneity of the emulsions. The F1 emulsions also showed a higher zeta potential than F2, but they were not significantly different. The F1 emulsion-chi 0.0% and the F2 emulsion-chi 0.0% showed negative values of zeta potential due to the absence of chitosan. The values were close to 30 mV, which revealed the stability of the emulsions. Zeta potential is an indicator of the stability of an emulsion and is influenced by the electric charge of the interface [27,28]. This phenomenon indicated that the formula F1 emulsion was more stable than the formula F2 emulsion. The difference between the F1 and F2 emulsion formulas was in the eugenol concentration. The ratios of the mass emulsifier (WPMD conjugates) and eugenol in F1 and F2 emulsions were 5:3 and 5:1.5, respectively. The higher eugenol concentration in this study produced a better emulsion. The effect of the clove oil content on droplet size and stability in nanoemulsions was investigated by Shahavi et al. [29].

#### *3.2. Performance of Spray-Dried Powder*

The spray-dried samples redispersed in water had a larger mean droplet size of emulsion before spray-drying. This phenomenon denoted that some of the nanoparticles formed aggregations during the spray-drying process. Similar studies on the change of particle size during drying have shown that the particle size of essential oils encapsulated in zein nanoparticle samples also increased after lyophilizing [30]. Increased particle size was probably caused by the imperfections of the hydration or the occurrence of structural changes within atomized droplets during spray-drying. From the research by Chen et al. [23], structural changes can occur due to the differences in the distribution of essential oils that evaporated and condensed during spray-drying which later cooled. The difference in the concentration of eugenol used in the F1 and F2 treatment samples perhaps caused differences in the particles' structural changes during drying. Therefore, F1 samples (2.0% eugenol) had a larger dispersed particle size than that of the F2 samples (1.0% eugenol). Moreover, increasing the chitosan concentration also increased the size of the dried particles. The chitosan content of 0.33% in the emulsion significantly increased the size of the spray-dried powder particles. The F1 powder-chi 0.33% had the largest particle size (1243 nm), as well as PdI value (0.71) (Table 3).


**Table 3.** Particle size, polydispersity index, and zeta potential of powders.

(\*) F1 and F2 contained 2.0% and 1.0% eugenol, respectively, in emulsion. Data represent mean ± standard deviation. Different letters indicate significant differences among samples (*p* < 0.05).

In contrast to the emulsions, the F1 spray-dried samples had smaller zeta potential values than that of the F2 spray-dried samples. However, the trend remained the same, where F1-chi 1.0% had a higher zeta potential value than that of the F1-chi 0.2% and F1-chi 0.067%, likewise for the F2 samples.

Encapsulation performance of eugenol using WPMD conjugates and chitosan was determined by the percentage of encapsulation efficiency (EE) and loading capacity (LC). The percentage of EE and LC increased with increasing levels of chitosan concentration of 0.067%, 0.2%, and 0.33% (Table 4). Chitosan content of 0.33% in formula F1 significantly increased the LC and EE, however, formula F2 showed a different phenomenon. This finding is similar to that of Dima et al. [31], who concluded that coriander essential oil-loaded microspheres prepared in an alginate/chitosan system with a 1:2 ratio had a higher value of entrapment efficiency than microspheres prepared in an alginate/chitosan system with a 1:1 ratio.

**Table 4.** Eugenol feed and encapsulation performance.


(\*) F1 and F2 contained 2.0% and 1.0% eugenol, respectively, in emulsion. Data represent mean ± standard deviation. Different letters indicate significant differences among samples (*p* < 0.05).

The F1 spray-dried sample with 0.33% chitosan solution (F1 powder-chi 0.33%) had the highest loading capacity and was significantly different from the other samples (Table 4). The spray-dried samples with 0.067% chitosan solution for both formulas F1 and F2 showed the lowest percentage of EE and LC. These values were lower than the EE and LC of formulas F1 and F2 without chitosan. This could be attributed to the instability of the emulsion and the droplet size. Chen et al. [23] stated

that large particle sizes at low pH caused a decrease in total oil content and encapsulation efficiency in spray-dried powder. Another researcher stated that during the spray-drying process, smaller particles had a higher mass transfer rate and more quickly formed semipermeable membranes around the atomized droplets, which reduced the loss of volatile compounds [32]. This phenomenon could have happened to the F2-chi 0.067% spray-dried powder sample. This sample came from an emulsion that had the largest emulsion droplet size, the highest PdI value, and a low zeta potential value. However, the spray-dried powder of F1-chi 0.067% had a small particle size and PdI value, and therefore the opinion of Jafari et al. [32] is not appropriate for this case. There could be differences in interactions between the WPMD conjugate, eugenol, and chitosan solutions in the acetate buffer and the interactions between the WPMD conjugates, eugenol, and acetate buffer without chitosan. The difference in interaction caused a higher percentage of LC and EE of eugenol in the spray-dried powder F1-chi 0.0% than in F1-chi 0.067%, as well as between F2-chi 0.0% and F2-chi 0.067%.

The eugenol spray-dried powder produced from this research had higher loading than some similar research results. The eugenol loading of the F1 sample powder-chi 0.33% (16.8%) was higher than the result of Woranuch and Yoksan [33]. They stated that eugenol loading in their eugenol-loaded chitosan nanoparticle study ranged from 0.85% to 12.80%. Research on eugenol loaded in zein/casein nanocomplexes resulted in a loading of 5.5% [23]. The encapsulation with the WPMD conjugate produced a loading ranging from 5.0% to 7.9% [11]. Generally, the encapsulation efficiency of the F2 powders was higher than that of the F1 powders, except for F1-chi 0.33%. We found that a higher eugenol concentration lowered the encapsulation efficiency. This result is in line with that of Hosseini et al. [22], who reported that EE decreased with an increase in the initial essential oil content. This could be due to the saturation of essential oil loading into the wall material during the encapsulation process and also the effect of chitosan concentration.

The level of saturation of eugenol was known from experiments encapsulating eugenol with four levels of eugenol concentration. The result showed that the percentage of EE increased at eugenol concentrations up to 2%, and then decreased when loaded with 4.0% eugenol. The LC increased until 4.0% eugenol concentration, and then decreased at 8% eugenol concentration (Figure S1). Volatile compound retention can be influenced by an optimum percentage of infeed solid that varies for each carrier or encapsulating agent. However, in this research, all formulas used the same mass and type of infeed solid (WPMD conjugate and chitosan). Each formulation is distinguished by the percentage of eugenol. In general, the encapsulation process with spray drying usually used 20% flavor load based on carrier solids. The higher percentage of that flavor load usually causes unacceptably high losses of flavors during spray drying [34]. The formula with a concentration of eugenol 2% (F-2%) has a maximum percentage of encapsulation efficiency (EE). The EE is influenced by the mass of eugenol in feed, which also involves a solid carrier in its calculation. The percentage of eugenol in feed for formula F-2% is 35.71% (data not shown). This value was greater than the formula with 1% eugenol (21.74%) and smaller than the eugenol formula 4% (52.63%), and the eugenol formula 8% (68.97%). The percentage of eugenol in feed for F-2% was probably the optimum value in the formulation of this study because it produced the maximum percentage of EE, although the maximum loading was at F-4%. At F-4% the percentage of EE decreased. The ANOVA test results showed that the loading percentage between F-2% and F-4% was not significantly different, while the percentage of EE was significantly different. Therefore, it can be concluded that the 2.0% eugenol concentration was the best at obtaining the optimal percentages of loading and encapsulation efficiency in the encapsulation of double-layer eugenol using WPMD conjugates and chitosan.

#### *3.3. Di*ff*erential Scanning Calorimetry*

All spray-dried powders containing chitosan, except the microcapsule of F2 powder-chi 0.067%, showed endothermic processes of temperature peak at 74.8 ◦C until 78.6 ◦C (Figure 1A,B). Chitosan had two endothermic peaks at 85.88 and 275.26 ◦C (Figure S2). The endothermic peak for eugenol is 258.81 ◦C [35]. The disappearance of the peaks located at these temperatures indicated that there

was an interaction between the WPMD conjugates and eugenol, as well as with chitosan. Previous studies reported that the individual characteristics of the graph polymer wall material and the active compounds are not visible in the DSC graphs if the interaction and encapsulation efficiency are good [36–38]. The interaction between core and the wall material determined the retention of the active compound. According to Hill et al. [39], the varied ratio of essential oil to the wall material resulted in the differences in the interaction of essential with the wall material which, then, influenced the encapsulation efficiency. –

**Figure 1.** Differential scanning calorimetry (DSC) thermogram of F1 (**A**) and F2 (**B**) spray-dried powders.

According to the thermograms, the spray-dried powders with 0.067%, 0.2%, and 0.33% chitosan concentrations for both eugenol concentrations (F1 and F2) appeared to have adjacent endothermic peak temperatures (Figure 1A,B). Therefore, the chitosan concentration did not have a different effect on the thermal behavior of the eugenol spray-dried powder. However, the addition of chitosan as the second layer in eugenol encapsulation can protect the sample from denaturation and decomposition. This was indicated by the absence of the exothermic peak and only one endothermic peak with temperatures below 100 ◦C. The endothermic peak in the range of 100 ◦C can be related to the gelatinization process. There was a decomposition of carbohydrates and similar structures in the presence of water during heating. This process was complex because, at the same time, water vaporization also occurred. Meanwhile, the endothermic peak in the range of 200 ◦C is related to the melting materials and the possibility of partial and liquid evaporation [40].

The thermograms showed that the formulations without chitosan (F1 powder-chi 0.0% and F2 powder-chi 0.0%) had endothermic and exothermic peak temperatures. The F1 powder-chi 0.0% had 85.75 and 189.97 ◦C endothermic and exothermic peaks, respectively. The temperature peaks of F2 powder-chi 0.0% at 81.49 and 180.61 ◦C, respectively, were lower than those of F1 powder-chi 0.0%. These two types of peak temperatures were also observed in the thermograms of the WPMD conjugate powder and the F2 powder-chi 0.067% at 70.14 and 213.09 ◦C and at 80.32 and 178.62 ◦C, respectively. The exothermic peak was attributed to the occurrence of decomposition.

According to Ronkart et al. [41] and Beirão-da-Costa et al. [42], the material reorganization during the spray-drying process could affect the changes in microcapsule melting temperatures. In their investigation, reorganization occurred when the polymer was heated above its transition glass temperature (Tg). In this study, the spray-drying process was held at 150 ◦C. The DSC measurements showed that the temperature of Tg maltodextrin and chitosan was not detected, while the Tg temperature of whey protein was 119.47 ◦C (Figure not shown). Therefore, the occurrence of material reorganization in eugenol encapsulation cannot be ascertained.

#### *3.4. Fourier Transform Infrared Spectroscopy (FTIR)*

The eugenol spray-dried powder of formula F1 with different levels of chitosan concentrations had similar spectra profiles (Figure 2A). The presence of eugenol and the formation of encapsulation could be detected from the appearance of peaks at wavenumbers in the range of 1517–1561 cm−<sup>1</sup> in the F1 spray-dried powder samples. The encapsulation process caused a shift in wavenumbers of the spray-dried powder sample, however, the wavenumbers were in the range of functional groups that represent C = C aromatic ring and secondary amines, NH bend. This wavenumbers' shift probably indicated physical interactions among eugenol, WPMD, and chitosan. The results of Piletti [43] explained that the physical interaction between eugenol with the β-cyclodextrin molecules caused the modification of the O-H functional group. The IR bands of eugenol, WPMD conjugates, and chitosan are presented in Table S1. The existence of a functional group in the sample could also be shown by the percentage of transmittance. In general, the percentage of transmittance in the wavenumber range of 1550–1650 cm−<sup>1</sup> for F1-chi 0.0% was lower than for F1-chi 0.067% and F1-chi 0.33%. This proved that F1-chi 0.33% had more chitosan than other F1 samples. – −1 s' explained that the physical interaction between eugenol with the β – −1

**Figure 2.** Infrared spectra of F1 (**A**) and F2 (**B**) spray-dried powders.

−1 – −1 – – −1 – −1 The existence of a C = C aromatic group in all F1 powder samples was seen from peaks in the 1508 cm−<sup>1</sup> region. These results are in line with those of Sajomsang et al. [44] and Woranuch and Yoksan [33]. Strong absorption was detected in F1 powder-chi 0.33%, indicating the higher eugenol content (Table S2). The data in Table 2 show that the F1 powder-chi 0.33% had the highest percentage loading capacity of eugenol. This was also reinforced by the detection of the OH bend at 1408–1411 cm−<sup>1</sup> and a C–H aromatic ring at 559–998 cm−<sup>1</sup> . The wavenumber range of 1400–1419 cm−<sup>1</sup> represents phenol and OH bend groups [45]. The intensity of these peaks in F1 powder-chi 0.33% was slightly higher than in F1 powder-chi 0.0%, 0.067%, and 0.2% (Figure 2A). The peak shifts that occurred were also likely related to the physical interaction between eugenol and the wall materials in the encapsulation process as explained above.

The profiles of FTIR spectra of F2 spray-dried powder samples were generally different from F1 powders (Figure 2B). Compared with the spectra of F1 powders, the percentage of transmittance of spectra of F2 powders was higher, meaning that the absorption was lower, which indicates the weak intensity of certain functional groups (Table S3). Peaks in the range of 3271–3295 cm−<sup>1</sup> experienced an increase in transmittance. This indicated the intensity of the hydrogen bonds between eugenol and the WPMD conjugate or between eugenol and free maltodextrin in F2 powders was lower than in F1 powders. Peaks in the range of 1563–1577 cm−<sup>1</sup> in a spray-dried powder containing chitosan represented NH functional groups that showed the presence of chitosan. The existence of eugenol was detected at peaks of 1401–1402 cm−<sup>1</sup> , which could have been shifted from specific wavenumbers of eugenol (Table S1). However, the intensity of these wavenumbers was lower than that of F1 powders, as seen from the percentage of the transmittance (Figure 2A,B). This was because the eugenol loading of F2 powders was lower than that of F1 powders (Table 4).

−1

–

−1

–

–

−1

#### *3.5. In Vitro Release Characteristic of Eugenol*

A release study of eugenol from the spray-dried powder was carried out to determine the effect of chitosan concentration on the eugenol released from microcapsules at a predetermined time. The results showed that F1-chi 0.0% powder had the lowest cumulative percentage of eugenol release (41.35%) among other F1 spray-dried powders at the end of the observation time (Figure 3A). However, among the samples containing chitosan, F1-chi 0.33% powder showed the lowest eugenol cumulative release (57.22%) as compared with F1-chi 0.067% (69.78%) and F1-chi 0.2% (59.22%). Chitosan concentration seemed to have the effect of inhibiting the release of eugenol in F1 encapsulation formulations.

**Figure 3.** Release profile of eugenol from F1 (**A**) and F2 (**B**) spray-dried powders.

The difference in eugenol concentration affected its interaction with the WPMD conjugates and chitosan, which was thought to influence the release profile of eugenol from F2 microcapsules. The spray-dried powder of F2-chi 0.0% (40.16%) had a greater percentage of eugenol cumulative release than the F2 powder containing chitosan at the end of the observation time (Figure 3B). The F2-chi 0.067% (34.68%), 0.6% (35.55%), and 1.0% (32.89%) had adjacent release graphs; however, the F2-chi 0.33% appeared to have the lowest cumulative release percentage at 48 h. Thus, it can be concluded that the presence of chitosan retarded the release of eugenol from microcapsules.

The percentage of eugenol release increased for 6 h and began to slow down after that until 48 h (Figure 3A,B). The retardation of eugenol release was probably related to the diffusion of active compounds contained in the particles [24]. Furthermore, Agnihotri et al. [46] explained that drug release through a diffusion mechanism began with penetration of the medium into the particle system, which caused the particles to swell, and then the drug was released from the swollen matrix.

The mechanism and eugenol release kinetic model of the spray-dried powders were also studied. The calculation to determine the mechanism and release kinetics used the Korsmeyer–Peppas equation (Equation (5)). The results of the calculation of the release kinetics showed that the values of the diffusion exponential (n) of all samples were less than 0.43 (Table 5). This indicates that the release mechanism in the eugenol microcapsules followed Fickian release (case I transport).

Release of eugenol occurred through a diffusion process. Ritger and Peppas [47] explained the criteria of the n value as follows: n ≤ 0.43 indicates Fickian release, n = 0.85 indicates case II transport, and 0.43 < n < 0.85 indicates non-Fickian release (an anomalous behavior). The results also revealed the diffusion constant value (k) and Pearson coefficient (*R* 2 ). The k value indicates the kinetics of the release. The greater the value of k, the faster the release of an active compound from microcapsules [48].


**Table 5.** Kinetic data of eugenol released from microcapsules using the Korsmeyer–Peppas kinetic model.

F1 and F2 contained 2.0% and 1.0% eugenol, respectively, in emulsion, n = diffusion exponential, k = diffusion constant, R<sup>2</sup> = Pearson coefficient.

Regarding some of the phenomena encountered in this experiment, the F1 formula samples had a cumulative release percentage greater than the formula F2 samples. The result was supported by the k value of the samples. The k value of F1 samples was higher than that of the F2 samples, indicating that the release of eugenol from F1 samples was faster than that from the F2 samples. The F1-chi 0.0% had the lowest cumulative release percentage and k value among other F1 microcapsules, while F2-chi 0.0% had the highest cumulative release percentage among other F2 microcapsules. According to Keawchaoon and Yoksan [24], the amount and release rate of a component were influenced by the pH of the media. However, the media used in this release trial had the same pH (7.4) for all samples. The differences between each sample were the combination of eugenol concentration and chitosan concentration. The eugenol spray-dried powders were obtained from eugenol emulsions with different emulsion pH values. The addition of different concentrations of chitosan solution to the first eugenol emulsion (primary emulsion) produced a different pH value in the second eugenol emulsion (secondary emulsion). The F1-chi 0.0% and F2-chi 0.0% emulsions had a lower pH than F1 and F2 containing chitosan. The pH value of the emulsion formula F1 was lower than the pH value of the emulsion formula F2. The pH difference could have influenced the electrostatic interaction among eugenol, WPMD conjugates, and chitosan which, then, influenced the release of eugenol. The mechanism was explained by Combrinck et al. [48]. Another possible reason for the release phenomenon of F1 and F2 powders is the difference in eugenol loading (Table 4). An increase in the rate of release of essential oils from microcapsules containing more essential oils is due to a decrease in the thickness of the microcapsule wall because insufficient wall material encapsulates the oil completely [49].

#### **4. Conclusions**

In this study, WPMD conjugates acted as an emulsifier, as well as a coating agent, in eugenol encapsulation. The concentration of eugenol and chitosan had an effect on the properties of the emulsions and spray-dried powders, including particle size, zeta potential, encapsulation efficiency, and loading capacity. The spray-dried powder prepared using an emulsion of 2.0% *w*/*w* eugenol and 0.33% *w*/*w* chitosan had the highest eugenol loading of 16.8%. The eugenol loading of this formulation with chitosan was twice as much as the loading obtained without chitosan reported previously. The eugenol encapsulation using chitosan as the second layer turned out to be effective at stabilizing the emulsion, as revealed by the zeta potential value. The presence of chitosan also improved the thermal stability and prevented the decomposition of the encapsulation product. The differences in eugenol concentration affected the release profile, whereas chitosan concentration had a role in slowing eugenol release. Generally, the results of the study indicate that the eugenol-loaded formulation yielded a biocompatible product with potential applications as food additives and pharmaceuticals.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2076-3417/10/9/3205/s1, Figure S1: Encapsulation efficiency and loading capacity of eugenol microcapsules loading with various concentration of eugenol, Figure S2: DSC thermogram of chitosan powder, Table S1: IR band WPMD conjugate, eugenol, and chitosan obtained from the FTIR measurement of the research, Table S2: IR band and % transmittance of F1 samples, Table S3: IR band and % transmittance of F2 samples.

**Author Contributions:** Conceptualization, I.A., K.M. and M.N.; methodology, I.A. and K.M.; formal analysis, I.A.; investigation, I.A.; data curation, I.A. and K.M.; writing—original draft preparation, I.A.; writing—review and editing, I.A., K.M. and M.N.; visualization, I.A.; supervision, K.M. and M.N.; project administration, I.A. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received funding for the experimental work (2017–2020) and the APC (2020) from the Indonesian Agency for Agricultural Research and Development.

**Acknowledgments:** The authors thank the Indonesia Agency for Agricultural Research and Development, Ministry of Agricultural and Universitas Indonesia for the research funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Relationship between Color and Redox Potential (Eh) in Beef Meat Juice. Validation on Beef Meat**

**Paolo Cucci, Aimeric C. K. N'Gatta, Supakakul Sanguansuk, André Lebert and Fabrice Audonnet \***

Institut Pascal, Université Clermont Auvergne, CNRS, SIGMA Clermont, F-63000 Clermont-Ferrand, France; paolo.cucci@orange.fr (P.C.); charlesaimeric@gmail.com (A.C.K.N.); supakakul1@gmail.com (S.S.); andre.lebert@uca.fr (A.L.)

**\*** Correspondence: fabrice.audonnet@uca.fr; Tel.: +33-4-73-40-78-17

Received: 20 March 2020; Accepted: 29 April 2020; Published: 1 May 2020

**Abstract:** In France, around 3.5 million cattle are slaughtered each year, which represents 1.3 million tons of beef carcasses. However, waste due essentially to organoleptic defects is estimated at 3.4% of the production or 45,000 tons of beef carcasses. Microbiological contamination and color are the two major causes of defect. In order to prevent color defect, a study was performed to develop a new method for measuring rapidly and instantly the redox potential as an indicator of color changes in carcasses without slowing down the slaughter line. This measurement would allow to classify them upstream according to their time of colors changes in order to sort them and to avoid food waste in the future. Meat juice has been shown to be a good mimetic medium for the study of color changes. The effect of different parameters was studied in order to fix experimental conditions. Color change is faster in the juice than in the meat and faster at 20 ◦C than at 4 ◦C. Redox potential allows following color changes and a symmetry has been highlighted between this thermodynamic measure and color changes.

**Keywords:** redox potential; color transfer; beef juice; beef meat

#### **1. Introduction**

In France, each year, around 3.5 million cattle are slaughtered for meat production, which represents 1.3 million tons of beef carcasses [1]. Most of this meat is distributed in large and medium-sized stores and alone accounts for 75% of purchases of fresh red meat in France [2]. One in two consumers buy fresh red meat based on color [3]. However, if the red meat does not meet the consumer's requirements (a beautiful color and a good smell), the latter does not buy it, which leads to food waste. In 2016, waste due to organoleptic defects in beef was estimated at 3.4% of production or 45,000 tons of beef carcasses [4]. Microbiological contamination and color are the two major causes of observed defect in beef meat.

The color of the meat is due to the presence of myoglobin. Myoglobin is a water-soluble intramuscular protein that binds to oxygen and allows cellular respiration in the muscle [5]. It is made up of globin and a prosthetic group called heme. At the center of the heme is an iron atom with its six coordination sites. The sixth coordination site makes it possible to bind a ligand such as dioxygen or even water [6]. Depending on the oxidation state of iron (ferrous Fe2<sup>+</sup> or ferric Fe3+) and the bound ligand, myoglobin is commonly found in 3 forms: deoxymyoglobin, oxymyoglobin, or metmyglobin [7]. Each redox form of myoglobin, when it is mainly present in meat, generates different colors. Deoxymyoglobin is purplish in color and is combined with a vacuum product or immediately cut. Oxymyoglobin is bright red in color and is associated with good quality red meat in trays, while metmyoglobin, brown in color on meat, and results in consumer rejection of the

product. Indeed, there is a dynamic of interconversions (redox reactions) between these three forms of myoglobin [7]. Initially, myoglobin or deoxymyoglobin undergoes oxygenation or blooming by binding to oxygen and becomes red-colored oxymyoglobin. Under low partial oxygen pressure such as during cellular respiration by muscle tissue, oxymyoglobin can revert to deoxymyoglobin, which is sensitive to oxidation. This oxidation at the level of the iron atom in the center of the heme causes the passage of the deoxymyoglobin of purplish color in metmyoglobin of brown color.

The aim of this study is to carry out a rapid and instant measurement of the redox potential, Eh, directly on a carcass (without slowing down the slaughter line), at the separation between forequarter and hindquarter in order to evaluate the time necessary for meat color to change from red to brown. To the best of the authors' knowledge, there is no technology to date to measure color and/or redox potential directly on slaughter lines. This would allow carcasses to be classified upstream in order to sort them. Indeed, a carcass whose E<sup>h</sup> value would indicate an early transfer would then be sent to a line of processed products, while a carcass whose E<sup>h</sup> value would indicate a later transfer would be distributed in large and medium-sized stores and offered in cut and packaged meat. However, since meat is a complex medium and redox probes for semi-solid media do not exist, the use of a mimetic medium such as meat juice (i.e., meat exsudate) has been proposed as a first approach.

The strategy applied in this work consisted in simultaneously measuring the color of a beef meat juice and the E<sup>h</sup> in order to search for a relationship between these two variables. The main sources of variability have been identified in order to limit their effect. A validation of the relationship on beef meat was also performed.

#### **2. Materials and Methods**

#### *2.1. Biological Products*

Rib steaks from young cattle (Puigrenier slaughterhouse, Montluçon, France) were semi-dressed, vacuum-packed, and labeled for all the experiments. After reception, the meat is stored in the refrigerator at (4 ± 2) ◦C. Rib steak was selected because it is easily and quickly accessible during the cutting of the carcass in the forequarter and hindquarter.

#### 2.1.1. Meat Juice

The meat juice preparation is based on the study by Kim and Jeong [8]. After reception, muscles were cut into pieces 3 cm thick with a weight of about 80 g and packaged in Polyamide/Polyethylene (PA/PE) storage bags impermeable to gases (3 welds, 200 × 300 mm, Sovapack, Cuiseaux, France). The bags were hermetically sealed either under half vacuum (500 mbar) or under vacuum with a double-chamber machine (Multivac, Lagny Sur Marne, France) for packaging in bags. The samples were then frozen at (−20 ± 2) ◦C for 24 h and then thawed at (4 ± 2) ◦C for 72 to 96 h depending on the size of the samples. The obtained juices after freezing and thawing of the samples were collected in a sterile manner under a microbiological safety station. The juices recovered have undergone different preparatory stages or not depending on the objectives of the study:


#### 2.1.2. Meat Decontamination

Rib steaks were decontaminated with peracetic acid following the study of Lebert et al. [9]. Then, they are cut into slices about 2 to 3 cm thick for a weight of 350 to 400 g. The slices are then placed in sealable trays in PS/EVOH/PE (Form'plast, Chantrans, France). They were finally placed in a modified

atmosphere using a T200 semi-automatic sealer (Multivac, Lagny Sur Marne, France). OPP/T504 film (Soussanna, Thiais, France) was used to seal the trays.

#### *2.2. Color Monitoring*

#### 2.2.1. Meat Juice

A climatic chamber (Binder KMF 240, VWR, Fontenay-sous-Bois, France) was used to reproduce the conditions of an industrial storage fridge for meat. Indeed, it allows temperature between 2 and 20 ◦C and relative humidity between 95 and 99%. In order to light up the meat and follow the color changes inside the climatic chamber, two strips with fluorescent tubes (T5, cool white color, 27.7 cm, 6 W, Diall, Paris, France) spaced apart 30 cm have been installed. The lighting delivers about 360 lux on the samples (dual-display traceable luxmeter, VWR Collection, France). In addition, a balance (Kern SXS-6K-3M, Timber Production, Esmans, France) was used in situ to monitor the weight over time to ensure that the samples did not dry out during the experiment. A camera (Logitech C270 720p, Logitech, Paris, France) was installed vertically approximately 20 cm above the samples to follow meat juice color.

#### 2.2.2. Meat

An industrial system (ADIV, Clermont-Ferrand, France) was used to follow the color changes of red meat. In a cold room (Dagard, Boussac, France) supplied by a cold group (Arcos, Gorrevod, France), strips with fluorescent tubes (T8, cool white color, 60 cm, 18 W, Sylvania, Saint-Etienne, France) were fixed on both sides others of the device with a distance of 45 cm between each strip. In addition, they were placed 30 cm above the meat trays just like the cameras (Logitech C270 720p, Logitech, Paris, France). Lighting at this facility delivers approximately 1700 lux on meat samples.

#### *2.3. Development of a Redox Probe for Solid Media*

Since commercial redox probes for semi-solid media do not exist, a built in-house redox electrode has been developed. It is composed of two "working" electrodes of 1 mm in diameter, one of which consists of a platinum rod (99.95% purity, Surepure Chemetals, New-Jersey, United States) and the other of an oxidized iron rod. Each rod is connected by electric cables in order to recover the electrical signals (measured voltages) at the level of the computer (the whole is molded in a resin).

#### *2.4. Physico-Chemical Measurements*

For meat juices, pH probes (HI11310, Hanna Instruments, Lingolsheim, France) and E<sup>h</sup> probes (HI36180, Hanna Instruments, Lingolsheim, France) were used. The measurements were taken and recorded automatically every 15 min using tablets (pH-/mV-meter Edge HI2002-02, Hanna Instruments, Lingolsheim, France). For meat, the built in-house probes were used. The data was measured and recorded using our internally developed software. The probes for liquid and semi-solid media were sterilized with 70◦ alcohol before being used.

Since at 4 ◦C, the time of color transfer is quite long (up to 400 h), and based on the fact that all the studied reactions are physico-chemical reactions, it has been decided to speed up these reactions by performing the experiment at 20 ◦C.

#### *2.5. Color Monitoring and Images Analysis*

Images of the samples (juice and meat) were taken every 15 min by using a software developed with LabVIEW 2018 and the Vision Development module (National Instruments Corporation, USA). These images in bmp format were saved on a computer before being processed.

An image processing process was performed for each saved image. Using ImageJ software (National Institutes of Health, Bethesda, Maryland, USA), the values of Lightness (L\*), Red/Green index (a\*), and Blue/Yellow index (b\*) were calculated automatically on the selected area (use of macros developed in-house). The target values were calculated to verify that the camera calibration did not drift over time. In this case, the difference between the initial target value and the target value after derivation has been subtracted or added to the measured value of the sample. This procedure gave the kinetics of a\*.

#### *2.6. Data Normalization*

The normalization allows adjusting of a series of values according to a transformation function in order to make them comparable with a few specific reference points. It is necessary when the results and their interpretations can be affected by the incompatibility of the units or scales of measurement between variables. This data normalization method was used for the comparison of the red index (a\*) and the redox potential (Eh) variation curves. The transformation function (Equation (1)) used is:

$$\text{f(V\_i)} = (\text{V}\_{\text{i}} - \text{V}\_{\text{max}}) / (\text{V}\_{\text{max}} - \text{V}\_{\text{min}}) \tag{1}$$

where V<sup>i</sup> = value to normalize, Vmax = maximum value, Vmin = minimum value. This function allows to normalize (and then compare) the red index a\* and E<sup>h</sup> between 0 and −1.

#### **3. Results**

#### *3.1. Operating Conditions*

In the next subsections, different operating conditions applied on the juices were tested: volume (20 mL, 35 mL and 50 mL), dilution (1/5, 1/10 and 1/15), water used for the dilution (milliQ or physiological), packaging (air or vacuum), and centrifugation (or not). Even if it should be interesting *per se*, it is worth mentioning that juice composition and visible spectrophotometry were not used in this study. Firstly, concerning the juice composition, we have considered the composition of the juice as a pre-slaughter factor. It is part of the initial conditions and is dependent on the animal such as the type of animal diet, the stress of animal experienced before slaughter, or also the type of muscle as mentioned by Bekhit et al. [10]. So, it was not relevant to master it for all the different experiments since each animal is different and it is impossible to perform this analysis on the slaughter line. Secondly, monitoring the evolution of visible spectrophotometry peaks of the meat juice in order to quantify the relative percentages of deoxymyoglobin, oxymyoglobin, or metmyoglobin over time using empirical equations as proposed by Tang et al. [11], were not undertaken because theses analyzes are not there either feasible on the slaughter line. Lastly, one has to note that the pH evolution of the different juices was also studied, but in all cases, its evolution was stable (between 5.7 and 5.5). These slight variations are in the uncertainties of measurement of the probe (± 0.1 pH unit, corresponding to two times the standard deviation) and do not explain color changes. Consequently, it has been decided to not represent it in the following parts.

#### 3.1.1. Influence of the Samples Volumes and Dilutions

The purpose of this experiment is to show the importance of standardizing samples when measuring color, since the volume and the dilution have an impact on the initial and final color (after transfer) of a meat juice. Samples with different volumes (20 mL, 35 mL, and 50 mL) and different dilutions (1/5, 1/10, and 1/15) were made in order to obtain the best possible compromise (Figure 1).

The juice samples were obtained following the freezing/thawing of a rib steak cut into small pieces (3 cm thick with a weight of about 80 g) packed in air; the juices were centrifuged (at 4 ◦C, 10,000 g for 5 min), filtered (through a filter with a porosity of 0.22 µm) and diluted with milliQ water. Each sample was transferred to a crystallizer of the same format and monitoring of the color change was carried out for 24 h at 20 ◦C in the climatic chamber, the choice of 20 ◦C being motivated by the acceleration of the reaction kinetics. At t = 0 h, the juices of 20 mL, 35 mL, and 50 mL diluted 1/5, 1/10, and 1/15, respectively, showed an intermediate red color. After 24 h, all the juices were found to have turned

from a red to brown color where the smaller the volume and dilution, the darker the color of the juice, rending the color change difficult to analyze. Moreover, in order to measure E<sup>h</sup> reliably, the largest volume (50 mL) is better to properly immerse the redox probes in these crystallizers. Finally, in order to modify the properties of the juice initially recovered as little as possible, the smallest dilution (1/5) was used. Following this analysis, a volume of 50 mL and a 1/5 dilution were used in identical crystallizers for all the experiments.

− −

−

**Figure 1.** Visualization of the color of the same rib steak juice with different volumes (20 mL, 35 mL, and 50 mL) and different dilutions (1/5, 1/10, and 1/15) at t = 0 h and t = 24 h, at 20 ◦C. Pure meat juice (undiluted) is also presented for comparison to diluted juices.

#### 3.1.2. Influence of the Water Type Used for Dilution

In this experiment, two types of water were tested, milliQ water and physiological water. These two waters were chosen because they are the most used in the world of the food industry (except running water). Each meat juice from the same whole rib steak, vacuum-packed before freezing and thawing, was centrifuged, filtered, and then diluted 1/5 in 50 mL milliQ water or 50 mL physiological water previously sterilized. A monitoring of the transfer of duplicate juices was implemented at 20 ◦C in the climatic chamber. The results showed that the two diluted meat juices had the same initial a\* value (Figure 2a). After the normalization of the data, it was found that the meat juice diluted with water milliQ turned faster than meat juice diluted with physiological water. The juice diluted in milliQ water turned completely yellow after 24 h while the juice diluted in physiological water changed completely in color (also yellow) after 48 h (Figure 2b). In fact, in order to not add ionic species in the juices, it was decided to use milliQ water for all the other experiments.

#### 3.1.3. Influence of the Packaging and Centrifugation Conditions

It is known that the oxygen has a role on the oxidation of myoglobin. Thus, it is necessary to determine whether the packaging conditions of the meat, in air or under vacuum, before freezing/thawing, have an influence on the color change of the juice from which it comes. Two rib steak muscles were packed whole under two conditions, one in air (21% of O2) and the other in vacuum (0% of O2) before being frozen/thawed. The juices obtained were diluted 1/5 in 50 mL of milliQ water, centrifuged, filtered, and then placed at 20 ◦C in the climatic chamber. It was found that the meat packaged in air gives more volume of juice (Factor 2) than that packed in vacuum. Figure 3a shows that the normalized values of a\* are identical.

When the juice has been collected after the freeze/thaw step, it may contain residues of connective tissue and adipose tissue. In order not to affect the color change due to the presence of residual, easily oxidizable fats, and so that the juice passes more easily through the filter during the filtration stage, the juice has been centrifuged and therefore clarified. The aim of this experiment was to verify the possible impact of centrifugation on the color change of the meat juice. The juice samples from a vacuum-packed whole rib steak were centrifuged or not centrifuged then filtered (through a 0.22 µm

porosity filter) and diluted 1/5 in 50 mL with milliQ water. Monitoring of the color change of duplicate juices was implemented at 4 ◦C in the climatic chamber. This temperature of 4 ◦C was chosen compared to previous experiments at 20 ◦C in order to have a first approximation of the transfer time of a juice at 4 ◦C. After normalization, it was found that the curves of the a\* values of the centrifuged and non-centrifuged juices are identical (Figure 3b).

**Figure 2.** Monitoring of the color change (a\*) as a function of time of (**a**) a rib steak juice (50 mL of centrifuged juice, filtered and 1/5 diluted) at 20 ◦C according to the type of water (milliQ water, black solid line, or physiological water, grey dashed line) used to dilute the meat juice as a function of time and (**b**) the same data in normalized values, using Equation (1).

**Figure 3.** Monitoring of the color change (normalized values of a\*) as a function of time for (**a**) a rib steak juice (50 mL of centrifuged juice, filtered and 1/5 diluted) at 20 ◦C depending on the type of packaging (air, in black solid line), or under vacuum, in grey dashed line) and (**b**) a rib steak juice (50 mL of filtered juice and diluted 1/5) at 4 ◦C with, in grey dashed line, or without, black solid line, the step of centrifugation (centrifuged or not centrifuged).

### *3.2. Correlation between Color Transfers and E<sup>h</sup> in the Juice*

In order to be as close as possible to industrial conditions, it was decided to carry out this experiment with no filtering of the juice. Indeed, in the meat industry, carcasses, muscles, and meat do not undergo any decontamination step between the start and the end of the process. The goal is to check if E<sup>h</sup> is really a good indicator of the color change in this case. The parameters of the protocol for obtaining the juice were as follows: freezing/thawing of a whole rib steak muscle under vacuum, centrifugation, and 1/5 dilution of the juice in 50 mL of sterile milliQ water. The same juice was separated in four crystallizers (two duplicates without probes and two duplicates with probes), which were then placed at 4 ◦C in the climatic chamber. The results of the normalized values of a\* and of E<sup>h</sup> are represented in Figure 4. The first observation which can be made is that a certain "symmetry" is observed between a\* and E<sup>h</sup> (when a\* decreases, E<sup>h</sup> increases). First, the values of a\* start with a phase of slight decrease, up to 105 h where the juices are still red. Then, for the next 45 h, a sudden decrease phase is observed; at this end point, the juices have turned and are yellow/greenish in color. Finally, a new plateau phase is observed. Concerning the E<sup>h</sup> evolution, a slight increase is observed during the first 96 h, then, an exponential increase is measured before reaching a plateau after 140 h. One has to note that a crossover is observed at 125 h, when the color transfer begins to be well established.

**Figure 4.** Monitoring of the color transfer (normalized values of a\*, in black solid line) of a non-decontaminated rib steak juice (50 mL of juice diluted 1/5) at 4 ◦C and of the redox potential (normalized values of Eh, in grey dashed line) as a function of time and associated photos showing the color transfer of non-decontaminated juices (duplicated on the right) at t = 1 h, t = 125 h, and t = 150 h.

#### *3.3. From Beef Juice to Beef Meat Measurements*

#### 3.3.1. Detection Limit for Color Changes in Meat

As the color transfers in the juice samples are homogeneous, the image processing process made it possible to detect the color transfers using the whole surface of the juice. However, for meat, a detection limit had to be determined. Indeed, if the values of a\* are determined by taking into account the entire surface of the meat, the tacking time cannot be detected on the curve (only some pixels are concerned by a color transfer) as shown in Figure 5a. To solve this problem, different sizes of meat sample area were used as the area for measuring the values of a\*. These different areas were calculated by dividing the total number of pixels of the area of the meat sample by 1, 2, 4, 16, 32, 64, and 128 (Figure 5a). The results presented in Figure 5b show that from the 1/64 cut, the values of a\* obtained no longer vary for the same sample. Indeed, the precision is such that the transfer is then detectable on the curve of

values of a\*. Therefore for all meat samples, a measurement of the values of L\*, a\* and b\* was carried out on 1/64 of the total area at the place where the tack was observed.

**Figure 5.** (**a**) Example of a limit detection of the color transfer, using a surface of 1/2 (white separation); (**b**) normalized a\* values for different areas of meat measured for the same sample (area divided by 1, 2, 4, 16, 32, 64, and 128) over time.

#### 3.3.2. Comparison between Beef Juice and Beef Meat

The aim of this experiment is to compare the color changes of beef meat and its associated juice in order to know if the juice is really a mimetic medium of the meat. For this experiment, a rib steak and its associated juice were collected and placed at 4 ◦C in the climatic chamber. The juice underwent the same protocol as for the other experiments and results are shown in Figure 6, with top left, pure juice and top right, 1/5 diluted juice. The results represent the normalized values of a\* of rib steaks treated in 1/64 and its 1/5 diluted juice. With regard to the juice, the values of a\* have the same tendency as previously observed: a slight decrease up to about 96 h, followed by a more pronounced slope up to 240 h. For meat, the measured a\* values follow exactly the same trends as the juice, i.e., first of all a slight decrease (t < 80 h) then a more significant fall (t < 240 h). A color change from red to greenish was identified. The normalized values shown in Figure 6 are pretty close, even if the fall was slightly more rapid for the a\* values of the juice than for those of the meat. The juice is therefore a good mimetic medium with regard to color transfers although the kinetics of the transfer is faster in the juice than in the meat (which is consistent because the diffusion coefficients are different).

#### 3.3.3. Calibration of the Built in-House Redox Probes

Meat juice being a good mimetic medium for meat to study color changes, it was decided to use the built in-house redox probe in the juice to perform their calibration. Indeed, the probes were built with two "working" electrodes, as shown in Figure 7a (see Material and Methods section for details), so that there is no longer a reference electrode and the values cannot be compared with values of commercial redox electrode. To verify the validity of our built in-house electrode, it was introduced into an unfiltered juice, diluted to 1/5 with a commercial redox probe (Hanna Instrument). The juice recovery protocol is the same as for the previous experiment. The results of this experiment, in normalized values, are shown in Figure 7b. Even if there is no reference electrode on our built in-house electrode, the normalized E<sup>h</sup> curves follow the trends. Following these results in the juice, the built in-house electrode developed for semi-solid media seems to be very promising for tests on meat.

**Figure 6.** Monitoring of the color transfer (normalized values of a\*) of a rib steak, in grey dashed line, and its associated juice (50 mL pure on the left and 50 mL diluted 1/5 on the right), in black solid line, at 4 ◦C as a function of time, and the associated photos showing their color transfers at t = 1 h, t = 144 h, t = 240 h.

**Figure 7.** (**a**) Built in-house redox electrode composed of two "working" electrodes of 1 mm in diameter, one of which consists of a platinum rod (left) and the other of an oxidized iron rod (right) molded in a resin; (**b**) Monitoring of normalized E<sup>h</sup> measured with a commercial redox probe (Hanna Instrument, in grey dashed line) and our built in-house redox probe (in black solid line) in a steak juice diluted 1/5 unfiltered as a function of time.

#### *3.4. Validation on Beef Meat*

This experiment was done under semi-industrial conditions. The rib steaks were placed in a heat-sealed tray and the built in-house probes were placed on the meat before air sealing (21% of oxygen). The experiment was carried out with 6 samples (6 successive slices) from the same muscle. As for the location of the probe on the meat, it was positioned in the muscle making up the rib steak which turned most often according to other experiments carried out during this study. At this stage, the important point is to test the probe on the meat medium to see if it works. The values of a\* were measured as close as possible to the probe (a few mm) and represent 1/64 of the slice of meat. The results of this experiment are shown in Figure 8. At the start of the experiment, the value of a\* is 20 and the meat is red. A sharp drop was measured after about 5 h. A color transfer was detected at the redox probe after 20 h of experience. A greenish color can then be observed at the level of the redox probe. After 96 h, the values of a\* reach a plateau corresponding to a marked green color on the beef meat. For Eh, the trends are the same as what has just been presented, the values ranging from 460 mV to 520 mV during the increase of the redox potential, then a plateau of a few hours was measured and finally a start of fall occurs, ending at 485 mV after 144 h. Normalized values are shown in Figure 8 on the right. This standardization highlights that E<sup>h</sup> follows the color change (change from red to green) and that there is a crossover after 20 h which corresponds to the start of the color change. The built in-house redox probe therefore makes it possible to follow the color change in the semi-solid medium that is beef meat.

**Figure 8.** Monitoring the color transfer a\* of a rib steak in a heat-sealed tray in air and the redox potential, Eh, measured with a built in-house redox probe at 4 ◦C as a function of time (**left**), the values normalized as a function of time (at **right**) and the associated photos showing their color transfers at t = 1 h, t = 20 h, and t = 144 h.

#### **4. Discussion**

The aim of this study is to carry out a measurement of the E<sup>h</sup> on a carcass, at the separation between forequarter and hindquarter in order to evaluate, by the measurement of the color transfer a\*, the time necessary for meat color to change from red to brown. Since meat is a complex media, linked to the fact no commercial redox probes exist for semi-solid media, a preliminary study was the use of meat juice as a meat mimetic medium. For that, the volume which has been retained is 50 mL so that the redox probes used can be completely immersed. Regarding the dilution, it was decided to modify the juice as little as possible to stay as close as possible to the pure juice, the 1/5 dilution in milliQ water was therefore selected. The more the juice is diluted the more the amount of myoglobin (Mb) present decreases, which can influence the results. In fact, the muscles containing high concentrations of Mb appear darker than those containing less. These differences in myoglobin concentrations, oxygen consumption rate and autoxidation rate may explain the variations in color stability between different muscles [12]. The juice preparation steps are just as important and should be monitored to ensure that they have the least impact on color changes. Pure juice is not a good choice to follow color changes because it is too dark. A 1/5 dilution was applied with two different waters. The results showed that the color change was slowed down with the use of physiological water. Indeed, the use of NaCl is well known to act as a preservative and this can explain this slowing down although the study done by Maruitti and Bragagnolo [13] has highlighted the pro-oxidant effect of NaCl on lipid oxidation and so, on color transfers. In contrast, milliQ water has the same behavior as distilled water and according to the experience of Thiem et al. [14] on the preparation of meat juices, distilled water was used and the components in the meat juices were not changed after dilution. This thus validates the method used in this study.

Experience on the packaging conditions of meat before freezing has highlighted the fact that the juice extracted from meat frozen in air has a value of a\* higher than that of meat frozen in vacuum. This can be explained by the fact that the meat packaged in air was able to oxygenate during the freezing and thawing stages, resulting thus in a higher beginning value. However, no difference was observed on the color transfer itself. According to Baran et al. [15], when fresh meat is packaged with films having a high impermeability to oxygen (under vacuum), anaerobic growth of native microorganisms is favored and the growth of aerobic native microorganisms is limited unlike packaging under oxygen (under air). Moreover, Daniloski et al. [16] also showed similar results with these two types of packaging. The different types of packaging can influence the color change due to the presence of microorganisms. However the filtration step done in this study has removed all the microorganisms from the juices. This could explain why no difference was observed in the color transfer between the two studied packaging conditions.

The centrifugation step did not affect the color transfer but only the initial a\* value. Indeed, the non-centrifuged juice has a value of a\* higher than the centrifuged juice. This can be explained by the fact that the centrifuged juice has been clarified and is therefore clearer than the non-centrifuged juice. As a result, only the value of a\* was impacted. The purpose of the centrifugation step is thus to standardize the juice samples by removing any debris recovered from the juice after thawing and to facilitate filtration.

During the experiments on the unfiltered beef juices at 4 ◦C and on beef meat at 4 ◦C, it was observed that the redox potential and color changes are linked; it is thus possible to follow the color changes using the redox potential and vice versa. Indeed, a symmetry was highlighted between this thermodynamic quantity (Eh) and the color changes (a\*). During the color transfer, heme iron oxidizes and changes from ferrous to ferric [17]. The increase in the redox potential would therefore be linked to the loss of the red color of the meat which is closely linked to the oxidation of heme iron. In the work of Ke et al. [18], the redox potential of the *Psoas major* muscle (PM) increased significantly between 0 and 7 days and was associated with an increase of the percentage of metmyoglobin and a decrease in color stability (values of a\*). Additionally, the redox potential of the *Longissimus lumborum* (LL) muscle did not show any significant variation over time, and was associated with a stability of the color of the muscle. We thus confirmed the observations of Ke et al. [18] in meat juice at 4 ◦C by the E<sup>h</sup> increase, reflecting the oxidation of the medium during the loss of red color of the juice. Moreover, the experience comparing the color transfer between beef meat and its juice has shown that the juice and the meat have a simultaneous color transfer but that the kinetics of

the transfer are faster in the juice. This latter observation is supported by the fact that the juice being a liquid medium, the physico-chemical reactions has to be faster than in a semi-solid media, where the species diffusion coefficients are greater.

The temperature has also an influence on the kinetics of the color change. The color transfer curves for juice or meat at 4 ◦C are generally composed of three phases in our study. First, a plateau which corresponds to the latency time, that is to say the time necessary for the oxidation to set up, then a fairly brutal fall corresponding to the oxidation (change from red to yellow/brown) in itself and finally a plateau phase once the transfer is complete. On the other hand, in experiments at 20 ◦C, this first stage of the plateau is shorter, even sometimes nonexistent and the transfer time is twice as short. Indeed, at 4 ◦C, transfers started between 96 and 144 h for juice and meat and ended after 144 and 240 h for juice and meat, respectively, while at 20 ◦C transfers started quickly between 1 and 10 h depending on the juices and ended after 24 h. This is explained by the oxidation of myoglobin which is kinetically favored and faster at high temperatures [17,19]. However, for the experiment on the color transfer of meat with the monitoring of E<sup>h</sup> with the built in-house redox probe at 4 ◦C, the color transfer was earlier than for the experiment of comparison of the color transfer meat and its juice. Two hypotheses can be put forward. The first would be that this meat is more sensitive to the color change than another because of its internal variability. The second would be that the redox probe favored the color change. Indeed, although the built in-house redox probe correctly follows the color change, with an almost constant delta compared to the values measured with the commercial redox probe, the oxidized iron that composes it may have initiated the color change at the location where it was inserted. Moreover, one has to note that this observation was exactely the same for the five other slices of meat under study, indicating that the probes helped trigger the color transfer on the six different samples. For exemple, Warner et al. [20] have shown that the iron concentration has an impact on the color change in mutton, where the color stability is reduced when the muscle had a high iron content. This argument would explain the systematic faster color transfer observed using the built in-house redox probe since, an electrode is made using an oxidized iron (Fe(III) state) rod. This iron rod was used since the aim was to not introduce other reducing metal species which would have produced metal ions which could have caused complexation reactions, which could themselves influence the color transfer of the meat. It is clear that the use of these probes is ultimately problematic in an industrial point of view, paricularly in slaughterhouses. However, the clear demonstrated relationship between the redox potential E<sup>h</sup> and the color transfer a\* seems to open the way to color measurement directly in contact with the meat. This would seem all the more interesting as this measurement is faster than that of the redox potential, for which probe development would also have to be carried out. Future work will then to model and predict these color transfers according to different factors, such as oxygen partial pressure, temperature or maturation time; this would allow carcasses to be classified upstream in order to sort them.

**Author Contributions:** Conceptualization, A.L. and F.A.; methodology, P.C.; investigation, P.C., A.C.K.N. and S.S.; writing—original draft preparation, P.C.; writing—review and editing, A.L. and F.A.; project administration, A.L. and F.A.; funding acquisition, A.L. and F.A. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was conducted with the support of the European Regional Development Fund (FEDER) program of 2014–2020, the Regional Council of Auvergne and the ADIV (Association pour le Développement de l'Institut de la Viande, Clermont-Ferrand, France).

**Acknowledgments:** Authors thank Pascal LAFON for wise advice for the choice and the implementation of cameras, David DUCHEZ for his help to take control of the software for image acquisition, Emilie PARAFITA and Valérie SCISLOWSKI for the relations with the slaughterhouse and the handling of the industrial installation of ADIV.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

MDPI St. Alban-Anlage 66 4052 Basel Switzerland Tel. +41 61 683 77 34 Fax +41 61 302 89 18 www.mdpi.com

*Applied Sciences* Editorial Office E-mail: applsci@mdpi.com www.mdpi.com/journal/applsci

MDPI St. Alban-Anlage 66 4052 Basel Switzerland

Tel: +41 61 683 77 34 Fax: +41 61 302 89 18

www.mdpi.com ISBN 978-3-0365-1841-1