**1. Introduction**

Microgreens are high-value crops considered the latest innovation in the vegetable sector [1]. Their supply and demand are highly influenced by emerging gastronomic trends and consumer familiarisation with the sensory attributes [2]. However, industrial production and marketing are limited since this highly respiring produce has a very short shelf life at ambient temperature [3,4]. Microgreens are known to respire during the germination process, metabolising stored carbohydrates in the cotyledonary leaves [5]. Once the carbohydrate sources are depleted, degradation of the microgreens occurs. Thus, modification of the plant metabolic activity and extending their shelf life by even a few days could be advantageous.

The main objectives of any postharvest technology are quality optimisation and loss reduction in fresh produce. Modification of the package atmosphere is one of the important methods in extending the shelf life. Passive modified atmosphere packaging (MAP) with relatively high oxygen transmission rate (OTR) films or perforated packaging is suggested to favour postharvest performance in microgreens [2]. High OTR polyethylene bags were preferred for the storage of radish microgreens over laser microperforated polypropylene packaging [6]. In another study, better postharvest quality of "Tah Tasai" Chinese cabbage microgreens was maintained in polyethylene bags compared to polypropylene packaging [3]. Sunflower microgreens have been reported to have a better shelf life and nutritional quality when packed in polystyrene trays compared to LDPE bags [7]. However, at the commercial front, microgreens are mostly packaged in plastic clamshell containers. To the best of our knowledge, there are no scientific reports comparing the e fficacy of such commercial packaging with polyethylene packaging on the postharvest quality and shelf life of microgreens. Therefore, such comparison warrants further studies.

Macro-perforated packaging, commonly preferred in commercial set-ups, is known to provide additional gaseous di ffusion and is beneficial in reducing o ff-flavour of fresh produce [8,9]. In addition, our earlier observations have shown that it minimises surface condensation on the packaging used for highly respiring produce, such as microgreens. However, it is often accompanied by higher weight loss, and the content is potentially exposed to outside contaminants. These limitations could be addressed using natural polymeric coating materials as primary packaging on the surface of fresh produce.

Edible coatings represent new packaging strategies in the postharvest managemen<sup>t</sup> of fresh produce. They are reported to create a micro-modified atmosphere around the produce by acting as a gas and water vapour barrier [10]. This helps in retarding food deterioration and enhancing its quality. Edible coatings are made up of natural polymers such as carbohydrates, proteins, and lipids. Edible coating applications have previously been reported to improve postharvest quality of fresh-cut produce such as celery sticks [11], and minimally processed lettuce [12,13]. However, to date, there are no published reports on the use of edible coating on microgreens.

In the last decade, there has been increased interest in using *Aloe vera* gel, as an edible coating on fruits and vegetables, due to its film-forming, antimicrobial, biodegradable and biochemical properties [14]. Benítez et al. [15] reported *Aloe vera* gel to be a better coating compared to chitosan and alginate coatings to extend the postharvest quality and shelf life of kiwi slices. *Aloe vera* gel (50%) was reported to reduce enzymatic browning in fresh-cut lotus roots and conserve the overall quality when stored at 5 ◦C [16].

The successful application of the edible coating on foods is dependent on several factors, including the method of application and its cost [17]. Dip coating technique is one of the age-old methods used commercially to coat fresh, whole, and minimally processed fruits and vegetables [18]. In earlier studies conducted in our laboratory, *Aloe vera* gel dip-coating gave promising results by reducing weight loss, minimising changes in the physicochemical parameters, reducing decay and extending the shelf life of papaya [19], figs [20] and litchi fruits [21]. A similar but less pronounced e ffect was observed in fenugreek and sunflower microgreens (unpublished data) using Aloe gel dip-coating. Dip-coating was found to be a little harsh on delicate and tender microgreens. There are also other drawbacks of dip-coating, such as the requirement of a large quantity of dip solution and quality deterioration of dip solution. Powder-coating was successfully used in our lab instead of dip coating for carrot shreds [22] and radish shreds [23]. However, this is not suitable for microgreens.

The spray-coating technique, which has recently attracted considerable industrial interest [24], was hence considered as an alternative technique. Chitosan postharvest dip-coating alone or combined with preharvest chitosan spray has been reported to enhance fruit quality and lower decay incidence in table grapes [25]. Recently, pre-harvest CaCl2 spray has been used to delay senescence in broccoli microgreens [26,27]. In another study, preharvest calcium spray displayed better overall quality and longer shelf life in broccoli microgreens than postharvest dip treatment [28]. However, to the best of our knowledge, there are no published scientific studies evaluating the e fficacy of preharvest spray treatment using a bio-based coating such as *Aloe vera* gel on fresh-cut leafy produce or microgreens.

In the present study, radish and roselle microgreens belonging to the Brassicaceae and Malvaceae families, respectively, with di fferent leaf morphologies, were selected. Radish microgreens are characterised by succulent cotyledonary leaves, while roselle have broad, thin, and flat leaves. Based on our nutritional evaluation studies among ten microgreens, these two microgreens were also found to be among the nutrient-rich ones [29], hence the need to optimise their postharvest quality. In the first phase of this study, the postharvest quality of these microgreens was assessed in two macro-perforated packaging, PET clamshell containers, and LDPE self-seal bags, commonly used for packaging fresh-cut produce, salad mixes and microgreens at the commercial and household levels. In the second phase of the study, spray- and dip-coating techniques were compared to study the e ffect of *Aloe vera* gel as an eco-friendly treatment on the postharvest quality and shelf life of radish and roselle microgreens.

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

#### *2.1. Plant Material Cultivation*

Good quality seeds (germination rate > 90%) of radish (RaS; *Raphanus sativus* L.) and roselle (HbS; *Hibiscus sabdari*ff*a* L.) were purchased from government-approved outlets of seed corporations (Anantapur, India). Seeds were broadcast in plastic trays (L: 24 × W: 17 × D: 4 cm) containing cocopeat in triplicates. The seeded trays were germinated in darkness at a relative humidity of 95% ± 5%. After two to three days of germination, they were exposed to sunlight (photoperiod 11.5 h; light intensity 2500–4400 lux) with an average air temperature of 25 ± 5 ◦C and relative humidity of 65% ± 10%. Seven-day old RaS and HbS microgreens were harvested by cutting the stem ends with sharp and sterile scissors. Microgreens were inspected prior to storage, and plants with defects or discoloured leaves were discarded.

#### *2.2. Experimental Design*

The study was conducted in two phases—Phase I determined the e ffect of packaging, and Phase II determined the e ffect of edible coating techniques on the postharvest quality and shelf life of RaS and HbS microgreens in the packaging which maintained better postharvest quality. The summary of the experimental design is represented in Figure 1.

**Figure 1.** Experimental design and sample coding. RaS: radish microgreens; HbS: roselle microgreens; PET–CS: PET clamshell container; LDPE–SSB: LDPE self-seal bag.

#### 2.2.1. Postharvest Packaging—Phase I

Fifteen grams of freshly harvested microgreens were packaged in clear macro-perforated (4 perforations of 3 mm diameter) polyethylene terephthalate clamshell containers (PET–CS) with hinged lid (dimension: 12.5 × 10 × 3.5 cm; thickness: 0.2 mm) or low-density polyethylene self-seal bags (LDPE–SSB) (dimension: 12.5 × 12.5 cm; thickness: 0.14 mm). The number of perforations was optimised in earlier experiments in our laboratory in order to minimise condensation on the inner package surface and, at the same time, retain the fresh weight of the produce (unpublished data). Samples were stored at 5 ◦C for 8 days. Quality evaluations were performed on 0, 4 and 8 days of storage, except for physiological loss in weight (PLW), which was measured every two days. Three replicates of each packaging were prepared for every analysis. A total of 84 packaged samples were obtained in Phase I (two microgreens × two packaging × seven parameters × three replicates).

#### 2.2.2. Edible Coating Techniques and Application—Phase II

Medium-sized and freshly harvested *Aloe vera* leaves were used to extract the gel according to a previously standardised protocol [20], and suitable dilutions were prepared for application on RaS and HbS microgreens. Two common techniques viz. spray coating and dip coating, were adopted in this study. The uncoated control (C) comprised of microgreens sprayed with water, prior to harvest. Preliminary trials were conducted to optimise the concentration of *Aloe vera* gel for application (unpublished data). The edible coating comprised of Aloe gel in an amount ranging from about 25 to 50 wt %, with the Aloe gel dip-coating (AGDC) having double the concentration of Aloe gel spray coating (AGSC). Prior to the harvest of microgreens, three trays were randomly selected for the spraying of Aloe gel. The AGSC was applied as a fine mist in the early hours of the morning as multilayers, with intermittent drying periods between the coating application. Harvesting was done upon complete drying of the Aloe gel coating on the surface of microgreens, and they were packaged in PET–CS. In the AGDC treatment, the microgreens were harvested from 3 random trays and dipped in Aloe gel and fan-dried for 5–10 min without allowing wilting to take place. Quality evaluations were performed on 0, 4, 8 and 12 days of storage, except for PLW, which was measured every two days. Three replicates of each treatment were selected for quality evaluations on every sampling day. The final number of packaged samples in Phase II was 126 (two microgreens × three treatments × seven parameters × three replicates).

#### *2.3. Quality Evaluations*

#### 2.3.1. Physiological Loss in Weight

The physiological loss in weight (PLW) was determined by accurately weighing the bagged samples at the beginning of storage and during storage at regular intervals (every two days). Results were expressed as a percentage of weight loss relative to the initial fresh weight of the microgreens [6].

#### 2.3.2. Respiration Rate

Respiration rates of the microgreens were determined in a closed system every 4 days during the storage period. Preliminary trials were conducted to determine the optimal incubation time for the studied microgreens. Gas samples were taken from the microgreens container every 15 min for a period of 1 h and evaluated until the CO2 level reaches a steady point. Thirty minutes was found to be the time when equilibrium was reached. Hence, 30 min was selected as the incubation time. In the case of LDPE–SSB, the macro-perforated package was placed inside a rigid container of a known volume containing ambient air as the initial atmosphere and incubated for 30 mins at 5 ◦C. This was carried out to minimise handling of the greens and ensure an air-tight atmosphere for gas sampling. In PET–CS containers, the macro-perforations were sealed during the incubation period. Gas composition (O2 and CO2%) in the headspace of the packaged sample was measured using a needle connected to the CO2/O2 gas analyser (PBI Dansensor, Checkmate II, Ringsted Denmark). The needle was inserted through a septum (silicone sealant) placed on the rigid container (in the case

of LDPE–SSB package) and directly through the septum of the PET–CS containers. The change in the concentration of CO2 evolved during the incubation period was used in the calculation of respiration rate using the following Equation (1) [30]:

$$\text{Resprimation rate} \left( \text{uL CO}\_2 \text{ g}^{-1} h^{-1} \right) = \frac{\left( \% \text{ CO}\_2 \right)\_{\text{Final}} - \left( \% \text{ CO}\_2 \right)\_{\text{initial}}}{\text{Sample weight} \times \text{incident time} \times 100} \times \text{headspace volume} \quad \text{(1)}$$

where (% CO2) final is the CO2 concentration after 30 min; (% CO2) initial is the CO2 concentration at the beginning of the incubation period; headspace volume is the volume of the container minus the volume occupied by the microgreens, expressed in μL; sample weight is the weight of microgreens on the evaluation day in g; incubation time is expressed in h.

#### 2.3.3. Electrolyte Leakage

Tissue electrolyte leakage was measured following the procedure given by Xiao et al. [31]. Samples (5 g) were submerged in 150 mL deionized water at 20 ◦C and shaken for 30 min. The electrolyte of the solution was measured using a conductivity meter (ELICO CM-180, India). Total electrolytes were obtained after freezing the samples at −20 ◦C for 24 h and subsequent thawing. Tissue electrolyte leakage was expressed as a percentage of the total electrolyte.

#### 2.3.4. Instrumental Colour

The instrumental colour of samples was measured with a Konica Minolta colour reader CR-10 (Minolta Co. Ltd., Osaka, Japan), equipped with an 8 mm aperture and calibrated with a white tile before the measurement was performed. The instrumental colour was measured in the form of CIELAB colour coordinates. The colour coordinate *L*\*, which denotes lightness, was measured in both microgreens. To trace the degradation of chlorophyll in the microgreens, *a\** (−) corresponding to greenness was recorded. The coordinate *b\** (+) denoting yellowness was measured in RaS as leaf yellowing was observed. In the case of HbS microgreens, since browning and not yellowing was a problem, chroma value, which denotes the overall chromacity, was calculated using the formula (a<sup>2</sup> + b2) 1/2. Leaves were plucked and placed in a 3-inch petri plate until filled with the sample. The probe of the colour reader was placed onto the adaxial surface of the leaves in the dish, and the reflectance spectra were measured by the instrument directly at three di fferent locations and the mean was calculated.

#### 2.3.5. Ascorbic Acid

The extraction and estimation of free, dehydro- and total ascorbic acid were performed according to the method given by Kampfenkel et al. [32], and the DHA/FAA ratio was computed and expressed. The concentration of ascorbic acid was calculated based on values obtained from the L-ascorbic acid standard curve (100–500 μg/mL). Results were expressed as mg/100 g fresh weight.

#### 2.3.6. Microbial Enumeration

To assess the microbial quality of microgreens, total aerobic mesophilic bacterial count (APC), and total yeas<sup>t</sup> and mold count (YMC) were determined. Aseptically weighed sample (1 g) was homogenised in a sterilised diluent (0.1% peptone water). The extract was centrifuged, filtered under sterile conditions, and volume was made up to 10 mL. The filtrate was serially diluted (10−<sup>1</sup> to <sup>10</sup>−5), and 100 μL of the appropriate dilution was spread on the agar plate using a spiral plater. The APC was determined by plating samples on the plate count agar, while YMC was determined by culturing on the potato dextrose agar. The incubation time was 24 and 48 h for APC and YMC, respectively. Microbial colonies were counted using a digital colony counter (Scan100 Interscience, St Nom, France), and results were reported as log CFU/g of sample.

#### 2.3.7. Overall Acceptability and Marketability

Microgreens were evaluated for overall acceptability by a group of 25 female panel members (selected from the authors' department). The panel members were familiarised with the samples and scoring system, but not specifically trained as they were to reflect consumer acceptability. Samples were coded and presented to the panelists immediately after opening the containers, in a randomised manner. The panelists were asked to rate the samples based on their degree of liking, using a 9-point hedonic scale.

End of shelf life was determined based on marketability score derived from the percentage loss of saleability (Equations (2)–(6); Table 1). The latter was a composite value calculated as a sum of 40% of the degree of wilting, 40% of the degree of yellowing/browning and 20% loss of overall acceptability. The degree of wilting/discoloration was determined by counting the number of wilted/discoloured leaves and expressed as a percentage of the total number of leaves in the package. The parameters were determined on duplicate samples. The overall acceptability (OA), as determined by the sensory panel on a 9-point hedonic scale, was first converted to a percentage. Hundred minus the % OA was the percent loss of acceptability. The equations used to derive the loss of overall saleability are given below:

$$Degree\ of\ writing(\text{\textquotedblleft}\text{\textquotedblright}) = \frac{\text{Number of leaves rolled}}{\text{Total number of leaves in package}}\tag{2}$$

$$\text{Degree of discoluration (\%)} = \frac{\text{Number of discoloured leaves}}{\text{Total number of leaves in package}} \tag{3}$$

$$\text{Overall acceptability } \left( \% \right) = \lfloor \frac{OA \text{ score}}{9} \rfloor \times 100 \tag{4}$$

$$\text{Loss of overall acceptable } (\%) = 100 - \% \text{ OA} \tag{5}$$

$$\text{Loss of overall salicylicity (\%)} = 40\% \text{ within} \lg + 40\% \text{ dissociation} + 20\% \text{ loss of OA} \tag{6}$$

Marketability scores were assigned as follows:

**Table 1.** Percentage loss of saleability and marketability score for shelf life quality assessment.


A loss of saleability of 20–29%, corresponding to a marketability score of ≤3, denoted loss of marketable shelf life of the produce.

#### 2.3.8. Scanning Electron Microscopy

The surface morphologies of coated (AGSC and AGDC) and uncoated (C) HbS microgreens were observed using an environmental scanning electron microscope (ESEM/VP–SEM–JEOL IT-300, Tokyo, Japan) operated in high vacuum mode. Samples were mounted on an aluminium stub using a double-sided adhesive carbon tape, sputter-coated with a thin layer of platinum and observed.

#### 2.3.9. Statistical Analysis

Three replications per treatment were employed, and results were expressed as means along with their standard deviation. All statistical analyses were performed using SPSS software (IBM SPSS Statistics 25, New York, NY, USA). Data obtained for the seven parameters (PLW, RR, EL, colour, ascorbic acid, microbial quality, and sensory acceptability) across the storage period were subjected to analysis

of variance (ANOVA) using the generalised linear model. This was followed by a posthoc Tukey HSD test at *p* ≤ 0.05 to determine significantly different groups. Graphical representations were performed using OriginPro® 2020 Graphing and Analysis software (OriginLab, Northampton, MA, USA).
