*Article* **Assessing the Potential of Algae Extracts for Extending the Shelf Life of Rainbow Trout (***Oncorhynchus mykiss***) Fillets**

**María I. Sáez, María D. Suárez, Francisco J. Alarcón and Tomás F. Martínez \***

Departamento de Biología y Geología, Campus de Excelencia Internacional del Mar CEIMAR, Universidad de Almería, 04120 Almería, Spain; msc880@ual.es (M.I.S.); dsuarez@ual.es (M.D.S.); falarcon@ual.es (F.J.A.)

**\*** Correspondence: tomas@ual.es; Tel.: +34-950015267

**Abstract:** This study evaluates the potential of different algae extracts (*Crassiphycus corneus*, *Cc*; *Ulva ohnoi*, *Uo*; *Arthrospira platensis*, *Ap*; *Haematococcus pluvialis*, *Hp*) as additives for the preservation of rainbow trout fillets. The extracts were prepared with different water to ethanol ratios from the four algae species. The highest ferric reducing antioxidant power (FRAP) was observed in *Uo* extracted in 80% ethanol. *Ap* aqueous extract also had considerable FRAP activity, in agreement with a high total phenolic content. Radical scavenging activity (DPPH) was higher in *Cc* 80% ethanol extract, in agreement with a high total carotenoid content. In fact, when the algae aqueous extracts were assayed on the fish fillets, their antioxidant activity exceeded that of ascorbic acid (ASC). All algae extracts delayed microbial growth and lipid oxidation processes in trout fillets throughout the cold storage period compared to controls, and also improved textural parameters, these effects being more evident for *Ap* and *Hp*. With respect to the color parameters, the *Hp* extract prevented the a\* values (redness) from decreasing throughout cold storage, a key point when it comes to colored species, not least salmonids. On the other hand, the *Ap* extract was not as effective as the rest of treatments in avoiding a\* and b\* decrease throughout the storage period, and thereby the color parameters were impaired. The results obtained, together with the natural origin and the viability for large-scale cultivation, make algae extracts interesting fish preservative agents for the food industry.

**Keywords:** algae extracts; antioxidants; fish preservatives; total carotenoids; total phenolics; trout fillets

#### **1. Introduction**

The high polyunsaturated fatty acids content in fish fillets contribute to an increased susceptibility to oxidative processes, which leads to decreased shelf life and sensorial quality. Therefore, the prevention of lipid oxidation with natural additives represents a major challenge for the seafood industry [1]. The interest in algae, both macro and microalgae, as a valuable source of potential additives for the food industry has increased considerably in the last few years. They are acknowledged for being rich in a wide variety of bioactive compounds, such as polyphenols and carotenoids, linked to remarkable antioxidant and antimicrobial activities [2]. Thus, some studies have broached the application of algae solutions as natural preservatives for fish products [3,4].

*Crassiphycus* sp. and *Ulva* sp. are two of the most extensively assessed genera of edible macroalgae, owing to their easiness for cultivation, together with their richness in bioactive compounds [5]. On the other hand, the interest in microalgae as natural sources of bioactive compounds is increasing [6], not least taking into consideration that they can be grown at large scale in bioreactors. Their cultivation under controlled conditions not only enables continuous supply, but also guarantees a production free from the eventual bioaccumulation of toxic substances [7]. Among the vast variety of microalgae species, *Haematococcus pluvialis* stands out as a source of astaxanthin [8], a pigment included routinely in commercial feeds for salmonids. With respect to *Arthrospira*, this is one of

**Citation:** Sáez, M.I.; Suárez, M.D.; Alarcón, F.J.; Martínez, T.F. Assessing the Potential of Algae Extracts for Extending the Shelf Life of Rainbow Trout (*Oncorhynchus mykiss*) Fillets. *Foods* **2021**, *10*, 910. https:// doi.org/10.3390/foods10050910

Academic Editors: Matteo Alessandro Del Nobile and Amalia Conte

Received: 23 February 2021 Accepted: 14 April 2021 Published: 21 April 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

the most widely assessed cyanobacteria genera in aquaculture, not only owing to its high nutritional value, but also to its acknowledged antioxidant activity [9,10].

In view of the above, this study was aimed at assessing the potential preservative effects of four algae species (*Crassiphycus corneus*, *Ulva ohnoi*, *Arthrospira platensis*, and *Haematococcus pluvialis*) on the shelf life and quality attributes of rainbow trout (*Oncorhynchus mykiss*) fillets kept under cold storage. The phenolic and carotenoid contents, as well as the antioxidant activity of algae extracts prepared with different water to ethanol ratios, were also studied.

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

#### *2.1. Materials*

Freeze-dried biomass of two marine macroalgae, *Ulva ohnoi* (*Uo*) and *Crassiphycus corneus* (*Cc*) were provided by LifeBioencapsulation S.L. (Almería, Spain). SABANA project (H2020 EU research program) supplied cyanobacteria *Arthrospira platensis* (*Ap*) and microalgae *Haematococcus pluvialis* (*Hp*) freeze-dried powder. Ascorbic acid was purchased from Sigma-Aldrich (Madrid, Spain).

#### *2.2. Preparation of Extracts and Antioxidant Activity*

Algal extracts used in antioxidant activity determinations were prepared by mixing 10 g of lyophilized powder with 250 mL ethanol in distilled water at different concentrations (0%, 30%, 50% or 80% *v/v*), according to the procedure described by Santoso et al. [11]. The mixtures were homogenized by vigorous shaking (2 min) and then agitated for 24 h at room temperature (22 ◦C) in darkness with a magnetic stirrer. Then, the mixtures were centrifuged (8000× *g*, 20 min), and supernatants filtered through Whatman #1 paper. Extractions were carried out in triplicate. Extracts were stored at 4 ◦C until further use within the next 24 h.

#### 2.2.1. Total Phenolic and Carotenoid Contents

Folin–Ciocalteu spectrophotometric procedure was carried out as described by Singh et al. [12]. A gallic acid standard was prepared (0 to 200 μg mL<sup>−</sup>1) and the results for total phenolic content were expressed as mg of gallic acid equivalents g−1. Total carotenoid content of extracts was estimated spectrophotometrically at 470 nm according to the equations proposed by Lichtenthaler and Buschmann [13]. For both parameters, results were expressed as mg g<sup>−</sup>1.

#### 2.2.2. Ferric Reducing Antioxidant Power (FRAP)

The antioxidant capacity of algae extracts was estimated according to the methodology described by Hajimahmoodi et al. [14]. Working solutions were prepared by mixing 100 μL of the samples or standards with 3 mL of FRAP reagent (50 mL of 0.3 M acetate buffer, pH 3.6, 5 mL of 10 mM tripyridyltriazine prepared in 40 mM HCl, and 5 mL of 20 mM FeCl3), and kept in the dark for 20 min at room temperature. Absorbance was then measured at 593 nm. Standards were prepared with ethanolic solutions of 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox; Sigma Aldrich), and the results were expressed as μmol Trolox equivalents g<sup>−</sup>1.

#### 2.2.3. Radical Scavenging Activity Determination (DPPH)

This activity was measured according to the method described by Brand-Williams [15]. The reaction mixtures were prepared by adding 75 μL of the algae extracts into 150 μL of 100 μg mL−<sup>1</sup> 2,2-diphenyl-1-picryl-hydrazyl-hydrate (DPPH) solution, and then incubating at room temperature in the dark for 24 h. The transformation of DPPH from oxidized to reduced form was determined spectrophotometrically at 515 nm. Standards were prepared with ethanolic solutions of Trolox. Results were expressed as μmol Trolox equivalents g<sup>−</sup>1.

#### *2.3. Treatment of Fish Fillets with Algae Extracts*

#### 2.3.1. Preparation of the Dipping Solutions

Only aqueous algae extracts were used for dipping the fillets. The solutions were prepared by mixing 1.5 g of the respective lyophilized algae with autoclaved distilled water up to a final volume of 500 mL (0.3% *w/v* final concentration). The mixtures were homogenized by vigorous shaking (2 min), then agitated with a magnetic stirrer for 24 h at room temperature (22 ◦C) in darkness, and filtered through Whatman #1 paper. Ascorbic acid solution (0.3% *w/v*) was also prepared for comparative purposes. The six solutions were designed as follows: (i) control (CONTROL, distilled water); (ii) 0.3% ascorbic acid solution (ASC); (iii) 0.3% *Crassiphycus corneus* extract solution (*Cc*); (iv) 0.3% *Ulva ohnoi* extract solution (*Uo*); (v), 0.3% *Arthrospira platensis* extract solution (*Ap*); and (vi) 0.3% *Haematococcus pluvialis* extract solution (*Hp*). Values of pH and buffering capacity (mEq g−<sup>1</sup> biomass) of the experimental extracts were determined by acid–base neutralization in order to estimate their possible influence on fillet pH determinations.

#### 2.3.2. Fillet Treatment and Sampling

A total of 60 rainbow trout (500 ± 25 g) were provided by a commercial fish farm (Piscifactorías Andaluzas, Granada, Spain). Immediately after slaughtering, fish were mechanically filleted at the farm, then washed, dried, and kept on ice. Once in the laboratory (less than 2 h after slaughtering), fillets (210 ± 22 g) were distributed in 6 experimental lots of 20 units each (5 sampling points × 4 fillets). The 20 fillets of each treatment group were dipped into the experimental solutions. Autoclaved water was used for dipping control batch. Fillets were carefully placed in sterile trays where the solutions were previously added (500 mL solution per each 4 fillets), kept for 30 s, then removed and placed in a grid for drying during 10 min. Next, fillets were stored in groups of 4 in sterile polyethylene bags, and stored in a cold-room (4 ± 1 ◦C). Samples were withdrawn from each lot at 1, 4, 6, 8, and 12 days postmortem (dpm) for the experimental determinations.

#### 2.3.3. Microbial Counts

Total viable psychrophilic bacteria counts were carried out according to Sáez et al. [16]. Muscle pieces (1 g) were introduced into aseptic tubes with 10 mL of 0.1% (*w/v*) peptone water (Cultimed SL, Murcia, Spain) and homogenized for 60 s. Four 1-g pieces were withdrawn from each fillet. Appropriate dilutions were serially prepared (12 serial dilutions by mixing 100 μL with 900 μL of peptone water) and then 0.1 mL of each were spread onto plate count agar media. The prepared plates were incubated at 4 ◦C for 120 h. Microbiological loads were expressed as logarithm of colony-forming units (cfu) g−<sup>1</sup> tissue.

#### 2.3.4. Determination of Thiobarbituric Acid-Reactive Substances (TBARS)

TBARS were determined according to the method of Buege and Aust [17], as detailed in Molina et al. [18]. Extracts were prepared in triplicate. TBARS value was expressed as mg of malonyldialdehyde (MDA) kg−<sup>1</sup> fresh tissue.

#### 2.3.5. pH and Water Holding Capacity (WHC)

Fillet pH was determined by means of a penetration electrode as described in [19]. WHC was estimated by weighing 1 cm<sup>3</sup> samples before and after centrifugation, as described in Suárez et al. [19].

#### 2.3.6. Texture Profile Analysis (TPA)

Texture was measured by compression using a Texture Analyser (TXT2 plus, Stable Micro Systems, Surrey, England, UK), with a load cell of 5 kN. A cylindrical probe (20 mm diameter) was used for pressing downwards into the fillet at a constant speed of 1 mm s<sup>−</sup>1. The determinations were made in all fillets at two adjacent points at the front dorsal muscle. Results are the average of these two values. The probe was placed parallel to the muscle

fibers. The parameters hardness, springiness, cohesiveness, gumminess, chewiness, and resilience were calculated as described in Bourne [20].

#### 2.3.7. Fillet Color Determination

Flesh color was measured by L\*, a\*, and b\* system [21], using a Minolta Chroma meter CR400 device (Minolta, Osaka, Japan). The determinations were made at two adjacent points on the front dorsal part of each fillet. The parameters L\* (lightness, on a 0–100 point scale from black to white), a\* (estimates the position between red, positive values, and green, negative values), and b\* (estimates the position between yellow, positive values, and blue, negative values)) were recorded.

#### *2.4. Statistics*

Determinations on antioxidant activity of extracts were carried out in triplicate, and the effect of the categorical variables "algae species" and "water/ethanol ratio" were determined by analysis of variance (ANOVA). With regard to fillet determinations, the effect of the categorical variables "algae treatment" and "storage time", as well as their interactions, were determined for each numeric parameter studied by fitting a generalized lineal statistical model (GLM). Least squares means were tested for differences using Fisher's least significant difference (LSD) procedure. A significance level of 95% was considered to indicate statistical difference. Specific statistical software (SPSS 25, IBM Corporation Inc., Armonk, NY, USA) was used.

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

#### *3.1. Total Polyphenol (TPC) and Carotenoid (TCC) Contents and Antioxidant Activity*

The influence of solvent water to ethanol ratio on the characteristics of the algae extracts is shown in Table 1. The *Ap* water extract yielded the highest TPC value among the algae studied, followed by *Hp* and *Cc*, whereas *Uo* showed the lowest value. Previous studies have reported a wide variability in TPC for *Arthrospira* sp., likely as a result of both cultivation factors and extraction processes [22]. With regard to *H. pluvialis*, disparate TPC values have also been reported, attributable to different aspects, such as the culture growth phase and the extraction solvent used [23]. Similar TCP figures were obtained for both seaweeds (*Crassiphycus corneus* and *Ulva ohnoi*), which are within the range reported by other authors [24].

The efficiency of polyphenol extraction depended on the solvent water to ethanol ratio considered (Table 1). Roughly, water extraction was more efficient than the rest of mixtures, in agreement with previous studies [14,23], a fact that might be attributed to the partially polar nature of the phenolic compounds. However, opposite results were reported by Mazumder [22] for *A. platensis*, who observed higher phenolic yield, as well as higher antioxidant activity, after organic solvent extraction, such as 60% ethanol, hexane, or methanol.

With respect to carotenoids (TCC) in microalgae, aqueous extracts yielded higher values for *H. pluvialis* than for *A. platensis*, in line with previous studies [25]. Nevertheless, comparison among different studies should be made cautiously, given that microalgae composition could be influenced by several cultivation factors, mostly nutrient availability and light intensity. With the exception of *Ap*, increasing the proportion of ethanol in the extraction solutions yielded higher TCC in extracts.

With regard to the antioxidant activity of algae extracts, the relative contribution of TPC and TCC to antioxidant capacity has not been well established yet. In our study, FRAP activity was remarkable in *A. platensis* aqueous extracts, in agreement with their high phenolic and carotenoid contents. For seaweed extracts, those with superior TCC values have showed outstanding antioxidant activity (not least DPPH in *Cc* and FRAP in *Uo*). Other authors have reported that phenolics and carotenoids contributed similarly to the antioxidant activity in several microalgae species [23]. The study by Yarnpakdee et al. [26] found a correlation between phenolic content in extracts and FRAP and DPPH

values in the seaweed *Cladophora glomerata*. In our study, the aqueous extracts showed a correlation between phenolic contents and FRAP activity (R<sup>2</sup> = 0.727), as well as between total carotenoid and DPPH activity (R<sup>2</sup> = 0.990) considering all the algal extracts as a whole. These differences suggest a dissimilar mechanism of action for both groups of compounds.

Since no extensive but only preliminary research has been carried out in this respect in our work, further studies are required to optimize the extraction process of antioxidants for food use, not only in these, but also in many other microalgae species.

Even if the extraction efficiency was higher by using organic solvents in some of the algae species in this study (not least in macroalgae), when it comes to possible practical application, it should be borne in mind that certain organic solvents might not be considered as safe for direct food use, and thus, water extraction might well represent a clear advantage. Moreover, it is reasonable to presume that aqueous extracts would cause a lesser impact on fish fillet sensorial parameters than organic solvent-based extracts. Keeping in mind all the above, only aqueous extracts were considered in the following assays on fillets.

**Table 1.** Total phenolic and carotenoid contents and antioxidant activity of algae extracts prepared with different water to ethanol ratios.


W/EtOH: water to ethanol ratio in solvents. TPC: total phenolic content of extracts, expressed as mg of gallic acid equivalents per g. TCC: total carotenoid content of extracts, expressed as μmol equivalents Trolox per g. FRAP: ferric reducing antioxidant power, expressed as μmol equivalent of Trolox per g. DPPH: radical scavenging activity, expressed as μmol equivalents Trolox per g. Superscript uppercase letters indicate differences attributable to algae species for the same water/ethanol proportion. Superscript lowercase letters indicate differences attributable to water/ethanol proportion for the same algal species. Values are mean ± sd on dry weight basis.

#### *3.2. Effects of Extracts on Fillet Quality Parameters*

#### 3.2.1. Total Viable Counts

Initial psychrophilic bacterial count in control fillets was 2.2 log CFU g−1, and increased over storage time, exceeding the maximum acceptable limit for fish (6 log CFU g<sup>−</sup>1; ICMSF [27]) at 12 dpm. Compared to controls, all treatments inhibited bacterial growth in fillets to a greater or lesser degree (Figure 1, Table S1). ASC markedly delayed bacterial growth in trout fillets, in agreement with previous studies [28,29]. Indeed, this compound with acknowledged antimicrobial activity is, up until now, one of the very few additives authorized for unprocessed fresh fish in the EU (Regulation 1333/2008/EC) [30].

In this regard, it is remarkable that the inhibition of microbial growth caused by all the aqueous algae extracts was significantly more intense than that observed for ASC, not least during the first six days of the storage period. The effects of the *H. pluvialis* and *A. platensis* solutions outweighed the other extracts at the initial stages of cold storage. These

results agree with the antimicrobial efficiency reported for the seaweed *Fucus spiralis* on refrigerated hake [31].

**Figure 1.** Changes in psychrophilic bacterial counts (PBC) in rainbow trout fillets treated with distilled water (CONTROL); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are expressed as mean ± sd. CFU stands for colony-forming units.

3.2.2. Lipid Oxidation (TBARS)

The initial TBARS content (Figure 2) in fillets was within the values established for good quality fish products (1–2 mg MDA kg<sup>−</sup>1) [32]. The values increased during storage in all treatments, but ASC showed remarkably lower values than controls, confirming its powerful antioxidant effects [29,33]. Interestingly, the seaweed extracts assessed also displayed outstanding effectiveness in protecting muscle lipids from oxidative processes (Figure 2, Table S2), which is the same as described in previous studies on different fish products (*Durvillaea antartica*, *Ulva lactuca*, and *Pyropia columbina* on canned salmon, [3]; *Fucus spiralis* on fresh hake, [30]; *Cladophora glomerata* on sliced tuna, [26]). Moreover, it should be pointed out that the algae extracts not only exceeded the antioxidant capacity of ASC, but also the effects persisted longer than those of ASC up to the end of the storage period.

Surprisingly, despite their potent antioxidant activity, research on the use of microalgae with the purpose of preserving fish products is not extensive. Takyar et al. [34] found that ethanolic extracts of *Chlorella vulgaris* and *A. platensis* yielded significant antioxidant activity on rainbow trout fillets. Studies on other food products also reported remarkable antioxidant effects (*A. platensis* in olive oil, [35]; *H. pluvialis* in ground pork [36]).

**Figure 2.** Time course of lipid oxidation in rainbow trout fillets treated with distilled water (CONTROL); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are given as mean ± sd. TBARS stands for thiobarbituric acid reactive substances, expressed as mg malonyldialdehyde (MDA) kg<sup>−</sup>1.

#### 3.2.3. pH and Water Holding Capacity (WHC)

The postmortem changes in pH and WHC are shown in Figure 3 and Table S3. The values increased significantly throughout the 12-day trial for pH in all experimental batches, likely owing to the emergence of alkaline compounds from protein bacterial degradation [37]. The significantly lower pH values observed in all treatments compared to the control fillets (being the lowest those caused by *Ap* and *Hp* extracts) might well be attributed to the antimicrobial effects also observed (Figure 1), which is same as found in previous studies [29]. Nevertheless, doubts can arise regarding the possible influence on fillet pH of the dipping solutions themselves, and therefore, this was also assessed. Overall, the extracts prepared at the concentration assayed (3 g L<sup>−</sup>1) yielded pH values within half a pH unit from neutrality (Table 2), except *H. pluvialis* and, especially, ASC, which showed a clearly acidic pH. Not only the pH value, but also the buffering capacity was estimated, and the results indicated that, with the exception of ASC, this parameter can be considered as negligible.

**Table 2.** Values of pH and buffering capacity measured for the aqueous dipping solutions (0.3% *w/v*).


**\*** Acidic extracts (ASC, *U. ohnoi*, *C. corneus*, *H. pluvialis*) were neutralized with 0.1 N NaOH, whereas alkaline extracts (*A. platensis*) were neutralized with 0.1 N HCl.

No evidence pointing to any effect of dipping solutions on fillet pH was observed, not even for ASC, likely owing to the fact that pH was not measured on fillet surface, but in dorsal muscle depth by means of a penetration electrode.

With regard to WHC, this parameter decreased throughout storage time, with the remaining values being statistically higher for *Hp*-treated fillets during the entire trial compared to the rest of treatments. This fact could indicate significant contribution of this extract in maintaining muscle mechanical properties, which is in agreement with the improvement observed in the textural parameters as well.

**Figure 3.** Postmortem changes in pH (**A**) and water holding capacity (WHC; **B**) of rainbow trout fillets treated with distilled water (CONTROL); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are expressed as mean ± sd.

#### 3.2.4. Texture Profile Analysis (TPA)

The TPA parameters are shown in Table 3. The hardness decreased in all treatments during cold storage, although all treatments yielded consistently higher values for this parameter than the control fillets at any sampling time from day one onwards. The postmortem deterioration of the textural parameters (not least muscle softening) is caused by myofibrillar and connective tissue proteolysis, which leads to a relaxation of the muscle structure [38]. Not a single factor, but a complex constellation of them (both biochemical and microbiological) is responsible for the alterations of the muscle structures throughout storage time, this ending in the unacceptable softening of the fresh fish. Given that muscle hardness is crucial in terms of purchasing decision, any strategy aimed at preserving this parameter deserves attention.

**Table 3.** Changes in texture profile analysis parameters of rainbow trout fillets treated with algae aqueous extracts throughout a 12-day cold storage (4 ◦C) period.


dpm: days postmortem. C: control (distilled water). ASC: ascorbic acid. *Cc*: *Crassiphycus corneus*. *Uo*: *Ulva ohnoi*. *Ap*: *Arthrospira platensis*. *Hp*: *Haematococcus pluvialis*. Values are mean ± sd. Superscript uppercase letters indicate differences (*p* < 0.05) attributable to storage time within each additive treatment. Superscript lowercase letters indicate differences (*p* < 0.05) attributable to treatments within each storage time.

> The delayed softening in all treated fillets might well have been linked to lower microbial counts (Figure 1), which is also in agreement with a recent study on rainbow trout fillets [29]. In addition, the inhibition of muscle endogenous protease activity owing to the antioxidant effect of the additives might also have occurred [39]. Although all the algae treatments kept a firmness above that of control fillets, it is remarkable that *Ap* and *Hp* were especially effective in this regard, outweighing even the effects of ASC from day six onwards. This might be related to the higher antioxidant activity observed for those aqueous extracts (Table 1). On the other hand, no consistent trend was observed for the rest

of the texture parameters measured (gumminess, chewiness, cohesiveness and resilience) neither regarding storage time nor additive treatment.

#### 3.2.5. Instrumental Color

The influence of the experimental treatments on fillet color parameters is shown in Table 4. The lightness (L\*) values increased over time in all treatments. Such an increase has been attributed in salmonids to lipid oxidation and protein denaturation, both effects together leading to higher light refraction on the fillet surface [40]. Color-related quality loss in trout fillets during cold storage results from the combination of increased L\*, and decreased a\* and b\* parameters, together jeopardizing their market value [41]. Such loss of pigmentation has also been partly attributed to astaxanthin degradation [42], included routinely in finishing diets for farmed salmonids, and responsible for their distinctive color.

The most evident effect of algae extracts on L\* was the significant decrease in this parameter up to 12 dpm caused by *Ap*, compared to the controls and to the rest of the lots, which, roughly, were similar. The *Ap* extract also caused a clear detrimental effect on the a\* and b\* parameters (as also found by Takyar et al. [34]). *Uo* also decreased fillet redness throughout the complete storage period, and it is likely that this effect can be attributed to the richness of *Uo* and *Ap* in chlorophylls and phycocyanins, respectively.

At the other end, the *Hp* extract accounted not only for yielding the highest a\* and b\* values among treatments, but also for causing the most persistent effects during the entire storage period (Table 4), a very desirable effect from the point of view of market acceptability. It is likely that the outstanding astaxanthin content in *Hp* was responsible not only for these favorable effects on color properties, but also for the remarkably positive antimicrobial, antioxidant, and textural effects found in this work, which makes this microalga a promising candidate as a fish preservative in colored muscle fillets.

However, when it comes to practical utilization, there is a clear incompatibility between the intense antioxidant and antimicrobial activities of *Uo* and *Ap* extracts and their coloring properties, which could limit their applicability on fresh fillets of white muscle fish species.


**Table 4.** Changes in color parameters of rainbow trout fillets treated with algae aqueous extracts throughout a 12-day cold storage (4 ◦C) period.

dpm: days postmortem. C: control (distilled water). ASC: ascorbic acid. *Cc*: *Crassiphycus corneus*. *Uo*: *Ulva ohnoi*. *Ap*: *Arthrospira platensis*. *Hp*: *Haematococcus pluvialis*. Values are mean ± sd. Superscript uppercase letters indicate differences (*p* < 0.05) attributable to storage time within each additive treatment. Superscript lowercase letters indicate differences (*p* < 0.05) attributable to treatments within each storage time.

#### **4. Conclusions**

The results indicate that all the algae species tested showed valuable antioxidant and antimicrobial effects, which might well be linked to their richness in both carotenoid and phenolic compounds. Outstanding FRAP and DPPH activities were found in *A. platensis* aqueous extract, in agreement with the highest phenolic content. Furthermore, the seaweed extracts with superior carotenoid contents yielded the highest antioxidant capacity (*C. corneus* and *U. ohnoi*).

Not only the antioxidant, but also the antimicrobial effects of the aqueous extracts of all the algae species tested were noteworthy, outweighing even those caused by ascorbic acid, the most widely used authorized additive for fresh fish. Together with delayed firmness loss, all the algae extracts assayed were able to extend the shelf life of the trout fillets compared to untreated controls.

When the impact on trout fillet color is also included in the overall assessment, *H. pluvialis* exceeded the effects of the other species, given that this extract lacks the detrimental effects observed for *A. platensis* on color parameters, despite its powerful antioxidant and antimicrobial activity. In view of the results, extracts of the algae species tested could represent valuable alternative sources of natural food additives with remarkable effects on fresh fish objective quality parameters. In addition to favorable antioxidant and antimicrobial properties, the extracts did not cause negative impact on trout flesh color parameters, which are considered crucial quality attributes for this species.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/foods10050910/s1, Table S1: Changes in psychrophilic bacterial counts (PBC, log cfu g−1) in rainbow trout fillets treated with distilled water (C); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are given as mean ± standard deviation (n=4 fillets), Table S2: Changes in lipid oxidation (estimated as mg MDA kg−<sup>1</sup> content) in rainbow trout fillets treated with distilled water (C); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are given as mean ± standard deviation (n = 4 fillets), Table S3: Changes in pH and water holding capacity (WHC) in rainbow trout fillets treated with distilled water (C); ascorbic acid (ASC) and aqueous extracts of *Crassiphycus corneus* (*Cc*), *Ulva ohnoi* (*Uo*), *Arthrospira platensis* (*Ap*) and *Haematococcus pluvialis* (*Hp*) during a 12-day cold storage (4 ◦C) period. Values are given as mean ± standard deviation (n = 4 fillets).

**Author Contributions:** Conceptualization: T.F.M., M.I.S., M.D.S., F.J.A. Investigation: T.F.M., M.I.S., M.D.S., F.J.A. Formal analysis: M.I.S. and M.D.S. Data curation: M.I.S. and M.D.S. Writing—original draft preparation: M.D.S. Writing—review and editing T.F.M. Supervision, funding acquisition and project administration: T.F.M. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by SABANA project (the European Union's Horizon 2020 Research and Innovation program, grant # 727874), the Spanish Government's MCIU RTI2018-096625- B-C31; and AquaTech4Feed (grant # PCI2020-112204) granted by AEI within the ERA-NET BioBlue COFUND. APC was co-funded by MCIU RTI2018-096625-B-C31 and Universidad de Almería (Spain).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data available on request.

**Acknowledgments:** The authors thank Pablo Medina, from Piscifactorías Andaluzas, S.L. (Granada, Spain) for kindly providing the fish used in the study and LifeBioencapsulation S.L. (Almería, Spain).

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

#### **References**


### *Article* **Effect of Cross-Linked Alginate/Oil Nanoemulsion Coating on Cracking and Quality Parameters of Sweet Cherries**

**Camilo Gutiérrez-Jara 1, Cristina Bilbao-Sainz 2, Tara McHugh 2, Bor-Sen Chiou 2, Tina Williams <sup>2</sup> and Ricardo Villalobos-Carvajal 1,\***


**Abstract:** The cracking of sweet cherries causes significant crop losses. Sweet cherries (cv. Bing) were coated by electro-spraying with an edible nanoemulsion (NE) of alginate and soybean oil with or without a CaCl2 cross-linker to reduce cracking. Coated sweet cherries were stored at 4 ◦C for 28 d. The barrier and fruit quality properties and nutritional values of the coated cherries were evaluated and compared with those of uncoated sweet cherries. Sweet cherries coated with NE + CaCl2 increased cracking tolerance by 53% and increased firmness. However, coated sweet cherries exhibited a 10% increase in water loss after 28 d due to decreased resistance to water vapor transfer. Coated sweet cherries showed a higher soluble solid content, titratable acidity, antioxidant capacity, and total soluble phenolic content compared with uncoated sweet cherries. Therefore, the use of the NE + CaCl2 coating on sweet cherries can help reduce cracking and maintain their postharvest quality.

**Keywords:** sweet cherry; nanoemulsion coating; cracking; fruit quality, nutraceutical value; crosslinking

#### **1. Introduction**

The sweet cherry fruit has a high nutritional value, mainly due to its high antioxidant capacity associated with ascorbic acid, carotenoids, and phenolic compounds [1]. The phenolic compounds in sweet cherries play a protective role against oxidative stress, ultraviolet radiation, and free radical damage [2]. However, the rapid deterioration of sweet cherries after harvest often leads to quality loss. More research is needed to develop novel strategies to prevent or reduce postharvest deterioration [3].

The cracking of sweet cherries caused by rain during the harvest period is the most important source of crop loss in the industry [4]. Rainfall, high humidity, high temperature, rootstock type, crop load, soil moisture levels, and irrigation management are some of the main factors that affect sweet cherry cracking [5]. As for the development of cracks on the skin of the fruits, the main mechanism proposed is related to the increase in turgor pressure caused by water absorption during and after rain. The two main routes of water absorption occur through the fruit surface [6] and/or the roots of the tree [7]. Various strategies have been used to reduce cracking. These include the use of plastic rain shields; adequate irrigation management; the application of calcium salts; and, more recently, the use of protective waterproofing agents [5]. The use of these technologies can reduce the severity of the damage. However, the degree of effectiveness widely varies among seasons, cultivars, and geographic locations. Some strategies, such as plastic rain shields, have high implementation costs [8].

**Citation:** Gutiérrez-Jara, C.; Bilbao-Sainz, C.; McHugh, T.; Chiou, B.-S.; Williams, T.; Villalobos-Carvajal, R. Effect of Cross-Linked Alginate/Oil Nanoemulsion Coating on Cracking and Quality Parameters of Sweet Cherries. *Foods* **2021**, *10*, 449. https://doi.org/10.3390/foods1002 0449

Academic Editors: Matteo Alessandro Del Nobile and Amalia Conte Received: 21 January 2021 Accepted: 9 February 2021 Published: 18 February 2021

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**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

The use of edible coatings is an alternative strategy that has emerged in recent decades [9,10]. This approach could be used to decrease the cracking phenomena of sweet cherries and extend their postharvest shelf life. Some of these edible coatings include chitosan [11], Aloe vera gel [12], and Semperfresh [13,14].

Alginate is another film-forming material that has been used as a thickening agent, gelling agent, and stabilizer in a variety of food emulsions [15]. It is a natural polysaccharide that is extracted from brown seaweed (Phaeophyceae) and comprises the two uronic acids, β-D-manuronic and α-L-guluronic. Sodium alginate consists of block polymers of sodium poly-L-guluronate, sodium poly-D-mannuronate, and alternate sequences of both sugars [16]. It has been effective in maintaining the postharvest quality of tomatoes [17], peaches [18], and sweet cherries [1]. However, to the best of our knowledge there are no previous studies on using alginate-based coatings to prevent sweet cherry cracking.

The cross-linking of the alginate film surface can be used to improve its mechanical and barrier properties because the film disintegrates when subjected to high humidity conditions due to its hydrophilic nature [19]. Cross-linking methods usually include drying, heating, ultraviolet (UV) irradiation, and chemical methods [20]. The chemical cross-linking method for sodium alginate involves the ionic interaction between polymer chains and multivalent ions to form ionomers. This improves their water barrier properties, mechanical strength, cohesiveness, and rigidity [10,21].

Another way to improve the water vapor barrier properties of the film involves adding lipids and nanofillers to form composite or nanocomposite films [22–25]. Smaller lipid globules and a more homogeneous distribution of the oil droplets in the films generally lead to better water vapor barrier properties [26].

Therefore, the aim of this study was to apply a nanoemulsion (soybean oil with alginate solution) on sweet cherries with additional ionic cross-linking and evaluate its effect on the water barrier properties of the sweet cherry cuticle, such as fruit cracking, postharvest qualities, and nutraceutical values.

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

#### *2.1. Plant Material and Experimental Design*

Sweet cherries (*Prunus avium* L.) cv. Bing were randomly collected at the commercial maturity stage from the midsection of 10 trees grown under standard commercial practices on the same commercial farm located in Brentwood (CA, USA). Cultural practices were regularly implemented for all trees equally. Fruits were transported immediately to the laboratory and selected by color, size, and the absence of physical defects or decay. Subsequently, the fruits were randomly distributed in 3 batches of 111 fruits each prior to treatment. The first batch was treated with a nanoemulsion (NE). The second batch was treated with a nanoemulsion and CaCl2 solution (NE + CaCl2). The third batch was used as a control. The barrier properties were determined immediately after each coating treatment. The quality parameters of the fruits stored at 4 ◦C were evaluated weekly for 28 d.

#### *2.2. Nanoemulsion Preparation*

The alginate solution (1.0%, *w*/*v*) was prepared by dissolving sodium alginate in a 2.5% ethanol solution. Ethanol was used to decrease the surface tension of nanoemulsions [27]. Glycerol was added at 15% alginate mass and the alginate solution was stirred for 30 min. Tween 80 (1.0% *v*/*v*) and soybean oil (0.5% *v*/*v*) were added to the solution and homogenized at 11,000 rpm for 2 min with a rotor-stator homogenizer (Polytron 3000, Kinematica, Littau, Switzerland). These coarse emulsions were passed six times through a microfluidizer processor (model 110T, Microfluidics, Asheville, NC, USA) at 200 MPa to obtain the nanoemulsions. The composition of the nanoemulsion and the process variables were selected on the basis of a prior study [28].

#### Particle Size and Polydispersity Index (PdI) of Nanoemulsions

The average nanoemulsion particle size and polydispersity index (PdI) were determined by dynamic light scattering (DLS) with a Zetasizer Nano ZS laser diffractometer (Malvern Instruments Ltd., Westborough, MA, USA). Emulsion samples were diluted in ultrapure water to 10% of the original concentration, placed in a cuvette, and analyzed at 25 ◦C. The average particle size (z-average) value and PdI were recorded.

#### *2.3. Coating Application*

The nanoemulsion (NE) was sprayed for 30 s at 30 cm from the surface of the sweet cherries with a cordless 85 kV vector solo waterborne electrostatic gun applicator (ITW Ransburg, Toledo, OH, USA). A 3.0% calcium chloride solution was applied using the same spray system after a coating was formed on the sweet cherries.

#### *2.4. Microstructure*

The microstructure of sweet cherry cross-sections was observed with a JEOL 7900F field emission scanning electron microscope (SEM) (JEOL, Kyoto, Japan) with a Quorum PP3010T cryo-system. First, the sweet cherries were cut parallel to the longitudinal axis with a scalpel. The sample was placed in the SEM sample holder and plunged into subcooled nitrogen (−210 ◦C). Afterward, the frozen sample was transferred to the cryo stage and freeze-fractured and gold-coated. The samples were viewed and photographed at 5 kV in the SEM.

#### *2.5. Barrier Properties*

#### 2.5.1. Cracking Index (CI)

The CI was determined using the method reported by Christensen [29]. For this purpose, sweet cherries harvested on the same day were selected based on size (22.3–24.9 mm), total soluble solids (20.03–20.24◦ Bx), firmness (3.70–3.78 N), water activity (0.964–0.978), and color (a \* = 10.16–13.83). Thirty fruits with stems from each batch were submerged in distilled water at 20 ◦C for 5 h to induce cracking. The number of cracked fruits was counted at 1 h intervals. The CI was calculated using the formula expressed in Equation (1).

$$\text{CI} = ((5\text{a} + 4\text{b} + 3\text{c} + 2\text{d} + 1\text{e})/(\text{MPV})) \times 100,\tag{1}$$

where a, b, c, d, and e represent the number of cracked samples at each time interval and MPV is the maximum possible value (30 fruits × 5 h = 150).

#### 2.5.2. Resistance to Water Vapor Transfer (RWVT)

During the same day of harvest, coated and uncoated sweet cherries were placed in a desiccator at 75.65% relative humidity using a saturated sodium chloride solution. Fans were used to ensure a uniform relative humidity throughout the desiccator. The desiccator was placed in a thermostatic chamber maintained at 4 ± 1 ◦C. Sweet cherries were weighed at 2 h intervals at 0.0001 g accuracy. The Resistance to Water Vapor Transfer (RWVT) was estimated by the equation of the first modified Fick law as established by different authors [30,31]. Weight loss data were used under stationary conditions. The RWVT was calculated by Equation (2).

$$\text{RPV}\text{'T} = \left[ \left( \left( \mathfrak{a}\_{\text{W}} - \text{@RH}/100 \right) \text{PWV} \right) / \text{RT} \right] \times \left( \text{A} / \text{I} \right), \tag{2}$$

where RWVT is the resistance to water vapor transfer (s/cm), aw is the sweet cherry water activity determined with a water activity meter (Aqua LAB 4TE, Pullman, WA, USA), %RH is the relative humidity inside the desiccator, PWV is the water vapor pressure at the chamber temperature (mm Hg), R is the universal gas constant (3,464,629 mm Hg cm3/g K), T is the storage chamber temperature (K), A is the sweet cherry surface area at the beginning of the test (cm2), and J is the slope of the weight loss curve under stationary conditions (g/s).

#### *2.6. Fruit Quality Parameters*

#### 2.6.1. Weight Loss

Fruit weight loss was evaluated with a digital balance (Precisa XB 320M, Dietikon, Switzerland). Sweet cherries were individually weighed at the beginning of the experiment and on each sampling day (7, 14, 21, and 28). Weight loss was expressed as a percentage of the initial weight and calculated by Equation (3).

$$\text{Weight loss (\%)} = \text{((W}\_{\text{o}} - \text{W}\_{\text{f}}) / \text{W}\_{\text{o}}) \times 100 \tag{3}$$

where Wo is the initial weight and Wf is the weight on the sampling day.

#### 2.6.2. Optical Properties

Color measurements were performed with a CR-300 colorimeter (Minolta Camera Co., Ltd., Osaka, Japan). The CIELAB parameters *a\**, *b\**, and *L\** were obtained with the D65 light source and an observation angle of 10◦ using the reflectance specular mode. The *L\** coordinate represented the lightness of the color (*L\** = 0 denoted black and *L\** = 100 denoted white), *a\** indicated the position between green and red (*a\** varied from −80 to +100), and *b\** was the extent of blueness/yellowness (*b\** varied from −50 to +70).

The hue angle (h◦) was calculated by Equation (4) as:

$$\text{Hue angle} = \arctan\left(b^\*/a^\*\right). \tag{4}$$

#### 2.6.3. Fruit Firmness

Mechanical tests were performed with a Texture Analyzer (TA-XT2i, Stable Microsystems Ltd., Surrey, UK) at room temperature using a puncture test. A probe (3 mm diameter stainless steel cylinder) with a trigger force of 5 N penetrated the sample to a depth of 8 mm at a speed of 1 mm s−1. Fruit firmness was measured as the maximum penetration force, and the results were expressed in newtons.

#### 2.6.4. Determination of Total Soluble Solids (TSS) and Titratable Acidity (TA)

For the TSS and TA tests, 5 g of sweet cherry tissue was homogenized in 25 mL of distilled water and filtered. The TSS was determined in the juice at 20 ◦C with a temperature-compensated LR-01 laboratory refractometer (Maselli Measurements Inc., Stockton, CA, USA). The TA was determined by titrating with 0.025 N of NaOH to pH 8.2 with a semi-automated titrator (Hanna Instruments, Woonsocket, RI, USA).

#### 2.6.5. Total Soluble Phenolic (TSP) Content

The TSP content was determined by the Folin–Ciocalteu method as described by Bilbao-Sainz et al. [32] with slight modifications. Samples (5 g) were homogenized with 20 mL of methanol with a Waring Laboratory Blender (Waring Commercial, Torrington, CT, USA) surrounded with dry ice for 1 min at medium speed. Samples were placed in tubes and stored for 20 to 72 h at 4 ◦C. Homogenates were centrifuged (rotor SA-600, Sorvall RC 5C Plus, Kendro Laboratory Products, Newtown, CT, USA) at 29,000× *g* for 15 min at 4 ◦C. Duplicate samples from each extract were used for the final analysis. A 150 μL aliquot of methanol extract was taken from the clear supernatant, diluted with 2400 μL of ultrapure water and 150 μL 0.125 mol L−<sup>1</sup> Folin–Ciocalteu reagent, and incubated for 3 min at room temperature. The reaction was stopped by adding 300 μL of 0.5 mol L−<sup>1</sup> Na2CO3 and the mixture was incubated for 25 min. Absorbance readings at 725 nm of clear supernatant samples were measured with a Shimadzu PharmaSpec UV-1700 spectrophotometer (Shimadzu Scientific Instruments, Inc., Columbia, MD, USA). A blank sample prepared with methanol was used as a control. The total amount of phenols was determined using a gallic acid standard curve and the results were expressed as milligrams of gallic acid equivalent (GAE) per gram of fresh weight (FW) of cherry purée. Three replicates were performed for each sample.

#### 2.6.6. Antioxidant Capacity (AC) Analysis

The AC analysis was adapted from Bilbao-Sainz et al. [32] with slight modifications. The same methanol extract from the TSP analysis was used for the AC analysis. Sample aliquots of 50 μL were taken from the clear supernatant (equivalent methanol volume as a control) and reacted with 2950 μL of 2,2-diphenyl-1-picrylhydrazyl (DPPH, 103.2 μmol L−<sup>1</sup> in methanol; absorbance approximately 1.2 at 515 nm) in a covered shaker at room temperature. The samples were allowed to react until steady-state conditions were reached. The AC was calculated with the PharmaSpec UV-1700 spectrophotometer by measuring the decrease in sample absorbance at 515 nm compared with the blank methanol sample. The AC was reported as μg Trolox equivalent from a standard curve developed with Trolox (0–750 μg mL<sup>−</sup>1) and expressed as mg Trolox g−<sup>1</sup> FW. Three replicates were performed for each sample.

#### 2.6.7. Total Anthocyanin Content

The total anthocyanin content was determined in duplicate with a PharmaSpec UV-1700 spectrophotometer (Shimadzu Scientific Instruments, Inc., Columbia, MD, USA) following the method reported by Serrano et al. [33]. Results were calculated by Equation (5) and expressed as milligrams 100 g−<sup>1</sup> FW.

$$\text{Total antibocycles} = \frac{\left(\frac{\Delta \text{BS}}{\varepsilon \times l} \times \text{MW} \times 1000\right) \times \left(\frac{V + W \times \rho}{1000}\right)}{W} \times 100\tag{5}$$

where ABS = absorbance of sample; ε = molar absorption coefficient (23,900 L mol−<sup>1</sup> cm−<sup>1</sup> for cyanidin-3-glucoside (cyd-3-glu)); l = path length in cm; MW = molecular weight (449.2 g mol−<sup>1</sup> for cyd-3-glu); V = volume of dilution in mL; W = sample weight in g; ρ = specific weight (0.83 in mL g<sup>−</sup>1); and 100 = 100 g of FW.

#### *2.7. Statistical Analysis*

A completely randomized design was used in the experiments. Statistical analysis was performed with the Statgraphics Centurion XVII software (version 17.1 12, Statgraphics Technologies Inc., The Plains, VA, USA) by a one-way analysis of variance. Significant differences between means were determined by the least significant difference (LSD) test at the 5% significance level (*p* < 0.05).

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

#### *3.1. Particle Size and Polydispersity Index (PdI) of Nanoemulsions*

After six passes through the microfluidizer, an emulsion was obtained with an average droplet size of 376.89 ± 2.73 nm and a PdI of 0.36 ± 0.04. These results concur with findings reported by other authors [34,35], who indicate that the increased number of passes through a homogenization system produces a reduced particle size and a more homogeneous particle size distribution. Similar results have been found by Artiga-Artigas et al. [36]. They achieved an emulsion with a 261 nm particle size and 0.25 PdI by mixing sodium alginate with an oil-in-water emulsion before the homogenization process (five passes). The smallest droplet size found in that study could be related to the different emulsion compositions because they used Tween 20 as an emulsifier and did not add a plasticizer.

#### *3.2. Microstructural Analysis*

Figure 1 shows the cross-sections of fresh sweet cherry (A), the NE coating (B), and NE + CaCl2 coating (C) on the sweet cherry surfaces. The micrograph of the uncoated sweet cherry surface shows a layer of the cuticle membrane over a layer of the regular epidermal cells, similar to the findings reported by Bargel et al. [37]. The layer of epidermal cells can be observed in all three micrographs. Subepidermal cells increased in size below the layer of epidermal cells because they were located farther away from the surface.

**Figure 1.** Cross-section micrographs of dried coatings on sweet cherry fruits. (**A**) Control, (**B**) nanoemulsion (NE), and (**C**) NE + CaCl2.

There was a continuous layer of NE coating on the sweet cherry surface (Figure 1B) as a result of the good adhesion of the NE. This adhesion could be attributed to the low surface tension of the coating formation solution because Tween 80 [38] and ethanol [27] were added. Meanwhile, the sweet cherries coated with NE + CaCl2 showed two layers. One layer was the NE on the cuticular membrane and the other was more compact and corresponded to the alginate cross-linked with calcium ions (arrows in Figure 1C). This second layer could reinforce the barrier properties of the cuticular membrane in sweet cherries.

#### *3.3. Barrier Properties*

#### 3.3.1. Cracking Index (CI)

Figure 2 shows the CI of coated and uncoated sweet cherries. The NE, which was expected to provide protection against water absorption by cherries and reduce their cracking, had the opposite effect and its application significantly increased the percentage of cracked sweet cherries (71.1%) compared with uncoated sweet cherries (65.6%). This effect could be due to the dissolution of the cuticular waxes by the Tween 80 emulsifier in the NE formulation, resulting in the increased water permeability of the cuticle [39]. This higher cuticle permeability could have induced the water diffusion inside the fruit, causing a localized burst of the cells that led to cracking [6].

**Figure 2.** Effect of coating sweet cherries with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) on the laboratory-induced cracking of sweet cherries. (a–c) Different letters indicate significant differences between treatments (*p* < 0.05).

Adding calcium ions to the NE coating dramatically increased the cracking tolerance and the percentage of cracked sweet cherries decreased from 65.5% to 12.2% (53.3% reduction). This result could be attributed to three different combined mechanisms. The first could be related to the decrease in osmotic potential on the fruit surface produced by the presence of calcium ions that did not react with the alginate [40]. Other authors mention that incorporating calcium ions on the surface of sweet cherries increased their cracking tolerance because of the decreased osmotic potential [41,42]. The second mechanism could involve strengthening the cuticular wax layer by Ca2+, hardening the cell walls to tolerate greater osmotic pressure before rupturing [43]. The third mechanism could be associated with the formation of a more compact and insoluble cross-linked alginate layer (Figure 1C) that increased the hydrophobicity of the sweet cherry surface [10,21]. This could have decreased the water diffusion from the outside to the inside of the sweet cherries, thus reducing the cracking percentage.

Other researchers have studied the application of hydrophobic coatings in the preharvest stage to reduce rain cracking in sweet cherries. Torres et al. [44] applied RainGard (mixture of fatty acids and vegetable oil) three times on cherry trees and reported 40.5%, 40.0%, and 52.0% reductions in rain cracking at harvest for Bing, Sweetheart, and Van cherries, respectively. They indicated that the coating waterproofed the surface of the cherries and acted as a filler for the micro-cracks in the cuticle. The application of cellulose nanofiber-based hydrophobic coatings (Innofresh) to Sweetheart cherry trees decreased the rain cracking between 31.18% and 44.60%, depending on the level of the surfactant mixture (Tween 80 and Span 80 at a 1:1 ratio) used in the coating [45]. The surfactant mixture was the most critical factor affecting the wettability, hydrophilicity, and elasticity of the coatings [45]. However, in other study an opposite result was found when spraying an anti-transpirant (Vaporgard) on Royal Ann sweet cherry trees 7 d before harvest [46]. They revealed that applying Vaporgard produced more cracking than in the controls by increasing the overall turgor in the trees and causing the cherries to exceed the strength of the cuticle or wall against rupture with minimal water absorption.

#### 3.3.2. Resistance to Water Vapor Transfer (RWVT)

Figure 3 shows the RWVT of coated and uncoated sweet cherries. All the coated fruits were less resistant to water loss than the control (Figure 3). The significant decrease in RWVT in the coated sweet cherries could be due to the emulsifier (Tween 80) that altered some epicuticle sites by changing, partially damaging, or extracting wax from the cuticle. These alterations could cause the dilation of the hydrophilic pores and lead to greater cuticle permeability [39,47]. Crosslinking with CaCl2 did not increase the resistance to

water loss despite the presence of an additional insoluble cross-linked alginate layer. This behavior could be associated with the swelling of the NE layer produced by water vapor transfer from the inside to the outside of the fruit. This increased volume of the NE layer can cause mechanical damage in the outer layer of cross-linked alginate, thus reducing its water vapor barrier capacity.

**Figure 3.** Effect of coating sweet cherries with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) on the resistance to water vapor transfer. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

#### *3.4. Fruit Quality Parameters*

#### 3.4.1. Weight Loss

Fruit weight loss during postharvest is due to the gradient of water vapor pressure between the fruit and the surrounding air [48]. Both the layer of epidermal cells and the cuticle are responsible for controlling this weight loss. Sweet cherries are characterized by rapid senescence and a cuticle with low resistance to water vapor diffusion, which promotes rapid water loss from the fruit and stem [49]. In the present study, the weight loss of coated and uncoated sweet cherries progressively increased with storage time (Figure 4). Contrary to the expected effect, coated sweet cherries experienced greater weight loss (16% and 14% after 28 d at 4 ◦C for cherries coated with NE and NE + CaCl2, respectively) than uncoated cherries (4% after 28 d at 4 ◦C).

**Figure 4.** Effect of coting sweet cherries with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) on weight loss during cold storage. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

Similar results have been reported by Chiabrando and Giacalone [50] when applying alginate coatings at 1%, 3%, and 5%. These authors found that applying these coatings was not effective to reduce the expected weight loss. When using Big Lory sweet cherries, they obtained 8.15%, 7.40%, and 8.25% weight loss, respectively, after 21 d at 2 ◦C, while uncoated cherries reached 7.35%. In Grace sweet cherries, the control fruits and those coated with 1% alginate achieved a 10% weight loss, while cherries coated with 3% and 5% alginate reached 12%. However, Díaz-Mula et al. [1] obtained weight losses of 5.93%, 4.88%, and 3.71% in Sweetheart cherries when was applying an alginate coating at 1%, 3%, and 5%, respectively, after 16 d at 2 ◦C. Uncoated cherries had a weight loss of 6.81%. Some previous studies have indicated that gum arabic, almond gum [51], and chitosan [52] coatings can reduce the weight loss of sweet cherries.

In the present study, the higher weight loss of coated sweet cherries compared with uncoated sweet cherries can be attributed to the low RWVT of coated sweet cherries. This lower barrier capacity was caused by the emulsifier present in the NE coating, which increased the permeability of the sweet cherry cuticle, as described in Section 3.3.2. On the other hand, the crosslinking of the NE coating with CaCl2 only slightly reduced the weight loss of the sweet cherries, due to the increase in the water vapor permeability of the alginate layer caused by its swelling.

#### 3.4.2. Color Attributes

Changes in the skin color parameters of uncoated and coated sweet cherries during storage are shown in Figure 5. The hue angle was correlated with the anthocyanin content and the lowest hue angle values corresponded to high anthocyanin contents [53]. Hue angle values slightly decreased during storage in all the samples; the reduction was more pronounced from day 21 onward, especially in coated fruits (Figure 5). The coated fruits had lower hue values than uncoated fruits, and there were no significant differences between NE and NE + CaCl2. Decreased hue values represent the progress of the fruit ripening process, reaching darker red colors in more advanced stages of maturity. This decrease in hue values during storage has also been described for other sweet cherry cultivars [54] and in sweet cherries coated with alginate [1] and Semperfresh (sucrose esters and mono-diglycerides of fatty acids and sodium carboxymethyl cellulose) [14].

**Figure 5.** Color evolution (hue angle) of uncoated sweet cherries (control), sweet cherries coated with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) during cold storage. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

The lower hue values of the coated fruits compared with the uncoated fruits can be related to the greater water loss experienced by the coated fruits during storage and promoted by the emulsifier, as discussed in Section 3.4.1. As a consequence, the anthocyanin content in these fruits increased, thus producing a darker red color. In turn, no significant differences were observed in the hue values in the fruits coated with NE and NE + CaCl2, mainly due to the fact that the sweet cherries with these coatings presented similar weight losses (Figure 4).

#### 3.4.3. Fruit Firmness

Changes in postharvest firmness can be produced by moisture loss and enzymatic changes [55]. All the coated and uncoated fruits showed decreased firmness during storage (Figure 6). The coated sweet cherries exhibited higher firmness values than the uncoated fruits; however, no significant differences were observed between them as of day 21 of storage.

**Figure 6.** Changes in the firmness of uncoated sweet cherries (control), cherries coated with nanoemulsion (NE), and cherries coated with nanoemulsion and the application of CaCl2 (NE + CaCl2) during cold storage. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

Similar results have been described in several studies applying different edible coatings on sweet cherries such as Semperfresh [14], alginate [1,50], almond gum, gum arabic [51], and guar gum with ginseng extract [56]. In these studies, the greater firmness retention of coated sweet cherries has been explained by the delayed enzymatic degradation of the components responsible for fruit structural rigidity caused by a decreased respiratory rate and cold temperature; it is also associated with reduced fruit moisture loss or maintained cellular turgor pressure.

No significant differences in firmness were observed between the fruits coated with NE and NE + CaCl2. Even when a crosslinked alginate layer was formed on the surface of sweet cherries, it was not able to improve its water barrier capacity, obtaining a similar weight loss levels as those coated with NE and producing the same mechanical behavior.

#### 3.4.4. Determination of Titratable Acidity (TA) and Total Soluble Solids (TSS)

The TA and TSS of coated and uncoated sweet cherries are shown in Figure 7. The TA value at harvest was 1.18 ± 0.1% malic acid equivalent, which decreased during storage for all cherries reaching at the end of the storage period values of 0.97 ± 0.02%, 1.02 ± 0.03%, and 1.03 ± 0.04% for control, NE, and NE + CaCl2-coated sweet cherries, respectively (Figure 7A). The TA value decreased over time (Figure 7A) because organic acids are substrates for the enzymatic reactions of respiration [57]. From day 7 onwards, the uncoated sweet cherries showed a greater reduction in TA than the coated sweet cherries; however, these differences were not significant. These results indicate that the coatings used were not able to significantly reduce the respiratory rate of fruits because they did not delay the use of organic acids, which are used in the enzymatic reactions of respiration [14]. Similar results have also been described when using 1% alginate coatings [50] and coatings of different types of 1% chitosan [58] in sweet cherries.

**Figure 7.** Effects of coating sweet cherries with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) on (**A**) the titratable acidity and (**B**) total soluble solids during cold storage. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

Additionally, no significant differences were observed between the NE and NE + CaCl2 coatings. These results would also indicate that the formation of a cross-linked layer of alginate fails to improve the gas barrier capacity of the coating and thus reduce the respiratory rate of the coated fruits. Possibly, the nanoemulsified coating and its cross-linked layer undergo a plasticization process as water vapor is lost to the environment, reducing the gas barrier capacity of these coatings.

The initial TSS value was 20.0◦ Brix, but it increased during the storage period (Figure 7B). The coated sweet cherries exhibited a greater increase in TSS than the uncoated fruits, reaching values of 24.1 and 21.20◦ Brix, respectively, at the end of storage. The application of edible coatings on sweet cherries usually produces a delayed ripening process and a low increase in TSS compared with uncoated sweet cherries [54,55]. However, the opposite result was obtained in the present work. The higher TSS values for coated sweet cherries can be largely explained by the higher water loss in these fruits. Several authors point out that the loss of water during storage produces the concentration of sugars in coated fruits [55,59]. In the present study, the greater weight loss shown by the coated fruits compared to the uncoated fruits (Figure 4), due to the presence of emulsifier in the NE coatings as described in Sections 3.3.2 and 3.4.1, would be the main cause of these higher TSS values. This is also the reason why no significant differences were detected between

the fruits coated with NE and NE + CaCl2, since the fruits with both coatings showed similar levels of weight loss. As previously mentioned, the formation of an additional layer of cross-linked alginate on the sweet cherries was not able to reduce their water loss, because the nanoemulsified coating undergoes a swelling process during storage, reducing its water vapor barrier capacity.

The sourness and sweetness of sweet cherries are important for consumer acceptance. The TSS can be used to measure sweetness and TA to measure sourness [60]. Crisosto et al. [61] indicated that consumer acceptance and the level of satisfaction with sweet cherries increased with higher acidity (TA > 0.80%) and sweetness (TSS > 20.0%). According to our results, coated sweet cherries could have better consumer acceptance and level of satisfaction than the uncoated sweet cherries after 14 d because they had higher TA and TSS values.

#### *3.5. Total Soluble Phenolics (TSP), Anthocyanin Content, and Antioxidant Capacity (AC)*

The consumption of fruit and vegetables with high phytochemical contents such as anthocyanins and other polyphenolics, carotenoids, and vitamins C and D have been associated with the prevention of different chronic diseases [62]. Phenolic compounds also contribute to the sensory and organoleptic quality of sweet cherries, such as flavor and astringency [2] and their antioxidant potential [54].

Figure 8 shows that coatings affected the TSP content and AC of sweet cherries but not their anthocyanin content. The initial TSP content in uncoated sweet cherries was 1.62 ± 0.25 mg GAE/g (Figure 8A). The TSP values of uncoated and coated sweet cherries progressively increased over time. Coated sweet cherries had higher TSP values than uncoated sweet cherries. The TSP value after 28 d for uncoated sweet cherries was 2.11 ± 0.10 mg GAE/g, whereas the TSP values for sweet cherries coated with NE and NE + CaCl2 were 2.47 ± 0.08 and 2.56 ± 0.22 mg GAE/g, respectively.

The increase in the total polyphenol content in sweet cherries in the present study was contrary to the expected results. Several studies have reported a decrease in the total polyphenol content in sweet cherries during storage as a result of peroxidase and polyphenol oxidase enzyme activity during the ripening process. They have also mentioned that applying edible coatings on sweet cherries has achieved a higher total polyphenol retention compared with uncoated fruits due to the formation of a protective barrier to gases on their surface [49,50,63].

In our study, the increased total polyphenol content can be explained by the development of two phenomena. First, the formation of a high gas barrier coating might have reduced phenol enzymatic oxidation. Second, the significant water loss in the coated fruits during storage due to the emulsifier in the coatings might have produced a concentration of the soluble polyphenolic compounds.

Anthocyanins are responsible for the red color in sweet cherries; their content increases during postharvest storage because the ripening process progresses and they are used as a quality indicator of cherries [64]. The anthocyanin content for all sweet cherries progressively increased during the storage period, and there were no significant differences between the coated and uncoated fruits (Figure 8B). These results were consistent with the decrease in hue angles observed in sweet cherries during the same period (Figure 5), which indicates a color change in sweet cherries from reddish red to more violet red [53,54]. However, even when the coated fruits had lower hue values than the uncoated fruits, it was not possible to detect these differences in anthocyanin content, which likely occurred because of the increased anthocyanin concentration in these coated fruits caused by significant water loss during storage.

**Figure 8.** Effects of coating sweet cherries with nanoemulsion (NE) and nanoemulsion plus CaCl2 (NE + CaCl2) on (**A**) total soluble phenolics, (**B**) total anthocyanins, and (**C**) antioxidant capacity (DDPH) during cold storage. (a–b) Different letters indicate significant differences between treatments (*p* < 0.05).

The AC values of stored uncoated and coated sweet cherries progressively increased over time (Figure 8C). In general, sweet cherries coated with NE + CaCl2 exhibited a higher antioxidant activity than uncoated sweet cherries and those coated with NE. At the end of storage, the inhibition of DPPH radicals was 7.13 ± 0.74, 6.50 ± 0.24, and 5.83 ± 0.37 g Trolox/g for NE + CaCl2, NE, and uncoated sweet cherries, respectively. The higher CA of the coated fruits compared with the uncoated fruits could be explained by the increase in the TSP due to the modified internal atmosphere caused by the coatings [63] and the water loss indicated in Section 3.4.1.

#### **4. Conclusions**

The results presented in this study showed that the nanoemulsified coatings based on alginate and soybean oil presented different effects on the barrier properties and quality parameters of sweet cherries. The NE + CaCl2 coatings were able to significantly reduce the cracking of sweet cherries, achieving a 53.3% reduction compared to the control fruits, due to the formation of a cross-linked layer on the surface of the coatings caused by the addition of CaCl2 as a cross-linking agent.

The Ne coatings had a limited effect on the delay of the ripening process and the quality parameters of the cherries. The presence of an emulsifier in these coating could have altered the cuticle waxes and caused increased weight loss in sweet cherries coated, reducing their potential effect on the quality parameters of the cherries. This behavior was not improved either with the formation of the cross-linked alginate layer (NE + CaCl2). However, a higher retention of total polyphenols and antioxidant capacity of the sweet cherries coated with NE + CaCl2 was verified. Future studies should focus on optimizing the amount of emulsifier in nanoemulsified coatings to improve the barrier properties and make them more effective in delaying the ripening of sweet cherries.

**Author Contributions:** Conceptualization, R.V.-C. and C.G.-J.; methodology, R.V.-C., B.-S.C., C.B.-S., and T.W.; software, B.-S.C.; validation, C.G.-J.; formal analysis, C.G.-J. and C.B.-S.; investigation, C.G.-J. and T.W.; resources, T.M.; data curation, B.-S.C.; writing—original draft preparation, C.G.-J. and C.B.-S.; writing—review and editing, R.V.-C., C.B.-S., T.M., and B.-S.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Western Regional Research Center (WRRC) of the United States Department of Agriculture (USDA), CONICYT-Chile (Doctoral Scholarship PFCHA/DOCTORADO BECAS CHILE/2016-21161275) and Universidad del Bío-Bío.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** The authors thank CONICYT for the Scholarship provided to the Camilo Gutierrez-Jara (PFCHA/DOCTORADO BECAS CHILE/2017—21170948) and the Graduate School of the Universidad del Bío-Bío, Chile for a research grant. The authors thank the Healthy Processed Foods Research Unit (HPFR) in the Western Regional Research Center (WRRC) of the United States Department of Agriculture (USDA), Albany, California, USA, for providing the infrastructure to carry out the research.

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

#### **References**

