Physiological and Biochemical Changes During the Growth of Custard Apple (Annona squamosa L.) Fruit Cultivated in Vietnam

Custard apple fruits (‘Na Dai’ cultivar) were collected from the four-year-old orchard in Dong Tan commune, Huu Lung district, Lang Son province, Vietnam. Experiments in this research were analyzed at the Physiological Plant and Application Laboratory, Faculty of Biology, Hanoi National University of Education, Vietnam.

Thirty healthy trees were selected from a total of 200 trees in a custard apple garden. Custard apple fruits were harvested in nine stages, beginning from week 3 to 16. Samples were collected according to the mixed sampling method. After collecting, samples were put into plastic bags with small ventilation holes, labeled, and transported to the laboratory for analysis.

The number of samples collected and used for the analysis of physiological and biochemical indicators in one stage was 30.

Determination of pigment content in the peel by the spectral method was made as per Ma et al. (2013). Took the green part of the fresh fruit peel with a sharp knife and accurately weighed 2 g into a porcelain mortar, crushed it and then added 80% acetone and centrifuged to collect the solution, and measured the solution on the spectrophotometer at the corresponding wavelengths. Chlorophyll content was calculated by the formula: C_a (mg·L⁻¹) = 9.784 × E₆₆₂ − 0.990 × E₆₄₄; C_b (mg·L⁻¹) = 21.426 × E₆₄₄ − 4.650 × E₆₆₂; C_(a+b) (mg·L⁻¹) = 5.134 × E₆₆₂ + 20.436 × E₆₄₄. Carotenoids content was calculated by the formula: C_car (mg·L⁻¹) = 4.695 × E_440.5 − 0.268 × C_(a+b). Then the pigment content per 1 g of fresh fruit peel was calculated by the formula: A=C×VP×1000; {\rm{A}} = {{{\rm{C}} \times {\rm{V}}} \over {{\rm{P}} \times 1000}}; where E₆₆₂, E₆₄₄, and E_440.5 are the results of measuring chlorophyll color at wavelengths of 662 nm, 644 nm, and 440.5 nm; C_a, C_b, and C_(a+b) are respectively chlorophyll contents a, b, and total; A (mg·g⁻¹) is the content of chlorophyll in 1 g of fresh fruit peel; C is the chlorophyll content of the pigment extract (mg·L⁻¹); V is the volume of pigment extract (10 mL); and P is the sample mass (g).

Vitamin C content was determined by the iodine titration method (Arya et al. 2000), in which 5 g of fresh fruit flesh was triturated with 5 mL of 5% HCl in a ceramic bowl. It was ground and put in a volumetric flask where distilled water was added to the 50 mL mark and well stirred. Then, 20 mL of the solution was put in a 100 mL conical flask and titrated by I₂ solution with starch as a color indicator until a blue color appeared, thereby determining the vitamin C content. The result was calculated based on the formula: X=V×V1×0.00088×100V2×g; {\rm{X}} = {{{\rm{V}} \times {\rm{V}}1 \times 0.00088 \times 100} \over {{\rm{V}}2 \times {\rm{g}}}}; where X is the content of vitamin C in the materials (%); V is the volume of diluted sample solution (mL); V1 is the volume of 0.01 N I₂ solution (mL); V2 is the volume of analyzed solution (mL); g is the weight of the sample (g); 0.00088 is the weight (g) of vitamin C, which was equivalent to 1 mL of 0.01 N I₂; and 100 is the conversion factor to percent.

Reducing sugar content and starch content was determined by the Bertrand method that was described by Mui (2001). Reducing sugar content was calculated using the following formula: X=a×V1×100V×b×1000; {\rm{X}} = {{{\rm{a}} \times {\rm{V}}1 \times 100} \over {{\rm{V}} \times {\rm{b}} \times 1000}}; where X is the reducing sugar content (%); a is the weight (mg) of glucose obtained when examining the table for volume KMnO₄ 1/30 N (mL) used for titration of the laboratory sample minus the volume KMnO₄ 1/30 N (mL) titration in the control sample; V is the volume of the diluted sample solution (mL); V1 is the volume of the analyzed sample solution (mL); b is the weight of the test sample (g); 100 is the conversion factor to percent; and 1000 is the coefficient converts grams to milligrams.

The starch content was calculated by the formula: Y=a×V1×100×0.9V2×b; {\rm{Y}} = {{{\rm{a}} \times {\rm{V}}1 \times 100 \times 0.9} \over {{\rm{V}}2 \times {\rm{b}}}}; where Y is the content of starch (%); a is the amount of reducing sugar; V1 is the volume of analyzed sample solution (mL); V2 is the volume of diluted sample solution (mL); b is the weight of the analyzed sample (g); 100 is the conversion factor to percent; and 0.9 is the coefficient of converting glucose into starch.

Determination of tannin content was by the Leventhal method (Chau et al. 1998), in which 2 g of crushed dried fruit pulp was added to a heat-resistant conical flask, and tannin was extracted by adding 100 ml of distilled water to the flask. It was boiled for 40 minutes in a water bath, then filtered to collect the solution. Extraction was continued until the filtrate was no longer reactive with tannins. The extract was then cooled and quantified through titration with KMnO₄. The tannin content was calculated by the formula: X=(a-b)×V×k×100V1; {\rm{X}} = {{({\rm{a}} - {\rm{b}}) \times {\rm{V}} \times {\rm{k}} \times 100} \over {{\rm{V}}1}}; where X is the tannin content (%) in dried weight; a is the volume of KMnO₄ 0.1 N used for titration in the flask (mL); b is the volume of KMnO₄ 0.1 N used for titration in the control vessel (mL); V is the total volume of extract (mL); V1 is the volume of the analyzed extract (mL); b is the weight of the analyzed sample (g); and k is the tannin coefficient and is 0.00582 (every 1 mL KMnO₄ 0.1 N is equivalent to 0.00582 g tannin).

Determination of total organic acid content by titration method is described by Ermakov et al. (1972). The total organic acid content in fresh fruit flesh was calculated by the formula: X=a×V1×100V2×p; {\rm{X}} = {{{\rm{a}} \times {\rm{V}}1 \times 100} \over {{\rm{V}}2 \times {\rm{p}}}}; where X is the amount of total organic acid present in the extract (mg·100 g⁻¹ fresh fruit); P is the amount of analytical sample (g); V1 is the total volume of extract (mL); V2 is the volume to be titrated (mL); and a is the amount of 0.1 N NaOH titration (mL).

Protein content was estimated by the micro-Kjeldahl method (Mui 2001) A=V×Va×0.142×5.595×100Vc×g; {\rm{A}} = {{{\rm{V}} \times {\rm{Va}} \times 0.142 \times 5.595 \times 100} \over {{\rm{Vc}} \times {\rm{g}}}}; where A is the protein content (%); Va is the volume of H₂SO₄ 0.01 N used to titrate BO₂; V is the total volume of enzyme extract (mL); Vc is the volume of NH₃ in analytical extract (mL); g is the weight of the crushed sample (g); 0.142 mg N is equivalent to 1 ml H₂SO₄ 0.01 N; 5.595 is the conversion factor to indicate the result of protein; and 100 is the conversion factor to percent.

Lipid content was determined by the Soxhlet method (Mui 2001). The fruit flesh was crushed, dried to constant weight, then the sample was put in a paper bag and placed in the extraction cylinder. The lipid content was calculated by the formula: X=Gm−GcG×100; {\rm{X}} = {{{\rm{Gm}} - {\rm{Gc}}} \over {\rm{G}}} \times 100; where X is the lipid content (%); Gm is the mass of the dry sample pack before extraction (g); Gc is the mass of lipid-extracted dry sample pack (g); and G is the mass of the initial sample taken for analysis (g).

The enzyme activity was determined using a spectrophotometer at 656 nm wavelength (Mui 2001). α-amylase enzyme activity was calculated by the formula: A=6.889×c−0.089388W {\rm{A}} = {{6.889 \times {\rm{c}} - 0.089388} \over {\rm{W}}} and c was calculated by the formula: c=OD1×OD2OD1×0.1; {\rm{c}} = {{{\rm{OD}}1 \times {\rm{OD}}2} \over {{\rm{OD}}1}} \times 0.1; where A is α-amylase enzyme activity (UI·g⁻¹·h⁻¹); C is the amount of starch hydrolyzed (g); OD1 is the optical density at the control vessel; OD2 is the optical density at the experimental flask; 0.1 is the amount of starch analyzed; W is the amount of analytical enzyme composition (g); and 6.889 and 0.089388 are the coefficients of the equation for the activity obtained by the mathematical processing method.

The activity was estimated by the A.N. Bac and A.I. Oparin method (Mui 2001). Catalase enzyme activity was calculated by the formula: X=(V1−V2)×1.7×VxVc×30×0.034×a×0.1; {\rm{X}} = {{({\rm{V}}1 - {\rm{V}}2) \times 1.7 \times {\rm{Vx}}} \over {{\rm{Vc}} \times 30 \times 0.034 \times {\rm{a}}}} \times 0.1; where X (µM H₂O₂·g⁻¹·min⁻¹) is the catalase activity calculated by the number of micromol H₂O₂ resolved in 1 minute under the action of catalase enzyme in 1 g sample at 30 °C; V1 is the volume of KMnO₄ 0.1 N used to titrate H₂O₂ in the control vessel (mL); V2 is the volume of KMnO₄ 0.1 N used to titrate H₂O₂ in the experimental flask (mL); Vx is the total volume of enzyme extract (mL); Vc is the volume of analytical extract (mL); a is the weight of the crushed sample (g); 1.7 is the conversion coefficient from the titrant KMnO₄ 0.1 N to mg H₂O₂ resolved; 30 is the duration of enzyme action (min); and 0.034 is the conversion factor of milligrams to micromoles.

Peroxidase activity was determined using the A.N. Boiarkin method on a spectrophotometer (Mui 2001): Peroxidase enzyme activity was calculated by the formula: A=E×a×bp×d×t; {\rm{A}} = {{{\rm{E}} \times {\rm{a}} \times {\rm{b}}} \over {{\rm{p}} \times {\rm{d}} \times {\rm{t}}}}; where A (UI·g⁻¹·s⁻¹) is the peroxidase activity in 1 g of sample; E is the selected optical density; a is the total volume of extract (mL); b is the degree of extract dilution; p is the weight of the plant sample (g); d is the cup thickness (cm); and t is the time (s).

All experiments were conducted three times independently. The results are expressed as mean values and standard deviation (SD). The results were subjected to an analysis of variance. Data were compared according to Tukey’s test using IRRISTAT software (version 5.0) for Windows computers (IRRI 2005).

During maturation, unlike other fruits, such as mango, persimmon, and tomato, although there is no noticeable discoloration in the custard apple, slight changes in color on the peel are also a sign of ripening.

Results in Table 1 show that the content of chlorophyll a, b, and total in fruit peel increase from week 3 to 13. After that, chlorophyll content decreased gradually from week 14 to 16. The chlorophyll content reached the highest value at week 13, specifically, the content of chlorophyll a was 0.046 mg·g⁻¹ fresh peel, chlorophyll b was 0.095 mg·g⁻¹, and total chlorophyll was 0.141 mg·g⁻¹ fresh peel.

The carotenoid content of the peel increased from fruit formation to maturity. Specifically, in the fruit development stage from week 3 to 13, the content of carotenoid increased slowly from 0.014 to 0.027 mg·g⁻¹ fresh peel, doubling after week 10 of development. Meanwhile, with only 2 weeks of growth, from week 13 to 15, carotenoid content in the peel increased nearly 2.5 times, reaching a value of 0.065 mg·g⁻¹. When the fruit was ripe, at week 16, this content tended to decrease slightly to a value of 0.063 mg·g⁻¹.

This change in pigment content is consistent with the change of color observed on the pods during the fruit growth. The newly formed fruit is pale green, then turns dark green (Gundewadi et al. 2018). When the fruit is nearly ripe, the green color gradually fades, turning white to yellow (Pal & Sampath Kumar 1995). This result is consistent with some studies that chlorophyll decomposition is related to the maturity of fruit (Du et al. 2014).

For most fruits, the reducing sugar content increases as the fruit enters the ripening stage (Paull et al. 1983). Some previous reports on mango or persimmon showed that the reducing sugars content tends to increase from the early stage to the maturity stage of the fruit (Trong et al. 2020a, b). In the present study on the custard apple, we did not observe a similar pattern of change. The data in Table 2 show that the content of reducing sugars in custard apple decreased gradually from week 3 to 7 by about 50%. The content of these sugars remained constant in the fruits until week 9, then increased sharply from week 14 to a peak at week 16, when it was 8.3 times higher than at week 3 (Table 2). This increase is higher than that announced in tomatoes (2 times) and mangoes (3.5 times) (Trong et al. 2019, 2020b).

Similar to the results of reducing sugar content analysis, the starch content in the custard apple decreased gradually from week 3 to 7, then increased and reached the maximum value at week 13, before decreasing gradually due to ripening fruit. The starch content of fruit at week 16 was 1.795%, only equal to 17.9% of the starch content at the week 13. The starch content decreased rapidly when the fruit was ripe following the increase in reducing sugar content of the custard apple as analyzed above.

The tannins in custard apple fruit had a relatively high content, reaching 11.410% at week 3. The high content of tannin in the early period made the fruit acrid and hard. The tannin content gradually decreased with the fruit development and dropped sharply until week 13. This decline is due to the breakdown of the hydrolytic tannins into pyrogallol and CO₂, which causes the fruit to ripen (Del Bubba et al. 2009). In the stage of fruit maturity, tannin content had dropped to only 0.582% at week 16; this result made the custard apple soft, not sour.

The results in Table 3 show that the contents of proteins, lipids, and total organic acids tend to decrease gradually depending on the development stage of custard apple. The vitamin C content fluctuates more complicatedly. It decreased from week 3 to 7, and then increased again from week 9 to 15, while at week 16 it began to decrease. The protein content of the fruit was highest at week 3, reaching 11.175%; this content was almost halved to 6.15% when the fruit ripened at week 16. Likewise, the lipid content also decreased from 11.45% (week 3) to 4.588% (week 16). As the fruit enters the ripening stage, vigorous action of pro-teases and lipases can lead to rapid hydrolysis of proteins and lipids in the flesh. The proteins content of the fruit decreased during growth and development because the proteins in fruit mainly act as enzymes and not reserves (Wills et al. 1998).

In persimmons and mangoes, the total organic acids contents usually increased gradually and reached the maximum value when the fruit entered the ripening stage (Trong et al. 2020a, b). Unlike these fruits, the total organic acids content was highest during the early stage of custard apple fruit development. Total organic acids content obtained at this stage (week 3) was 97.917 mg·100 g⁻¹ fresh fruit. During subsequent fruit development, the total organic acids content decreased gradually, and decreased most strongly at the ripening stage. In comparison with week 3, at week 16, the organic acid content decreased more than four times, and was 22.860 mg·100 g⁻¹ fresh fruit. This content sharp drop may be related to the participation of organic acids in energy respiration. On the other hand, energy continues to be needed for the biosynthesis of fruit-specific ripening substances, such as enzymes for hydrolysis, esters to create an aroma, and synthesis of sugars to create sweetness of fruits, resulting in a decrease in total acids content (Prasanna et al. 2007).

Vitamin C is an important nutritional indicator to evaluate the quality of many fruits. The amount of vitamin C in the custard apple varied quite complicatedly according to the development of the fruit. The analytical results showed that the vitamin C content in the flesh was relatively high when the fruit was newly formed, reaching 32.467 mg·100 g⁻¹ fresh fruit, then gradually decreasing to 18.667 mg·100 g⁻¹ fresh fruit at week 7. From week 7 to week 15, vitamin C content increased again. The highest vitamin C content was measured at week 15, reaching 36.2 mg·100 g⁻¹ fresh fruit. After that, vitamin C content decreased slightly to 35.5 mg·100 g⁻¹ fresh fruit when the fruit was fully ripe at the week 16. Vitamin C decreased due to its participation in the biosynthesis of ethylene, oxalate, and tartrate in the fruit (Singh et al. 2011).

Table 4 shows that the α-amylase activity of custard apple is lowest at week 3 and highest at week 15. Amylase enzyme activity increased slowly from week 3 to week 11, while this activity increased sharply between week 11 and 13, more than four times after week 2, from 2.460 to 9.958 UI·g⁻¹·h⁻¹. Then, amylase activity continued to increase and peaked at week 15 (12.208 UI·g⁻¹·h⁻¹). After that, it tended to decrease slightly to 11.167 UI·g⁻¹·h⁻¹ at week 16. The α-amylase activity change during the fruit development is consistent with the change in the sugar and starch content of the fruit when the fruit was ripe (Jain et al. 2001). This result was similar to the result of α-amylase activity analysis in some other fruits such as persimmons, tomatoes, and mangoes (Trong et al. 2019, 2020a, b).

In the early stages of fruit development, the activity of catalase was relatively high (2.083 µM H₂O₂·g⁻¹·min⁻¹ at week 3). This value increased significantly from the fruiting stage of week 3 to 11 and then gradually decreased. The highest catalase activity – 11.542 µM H₂O₂·g⁻¹·min⁻¹ was measured at week 11. The variability of the activity of antioxidant enzymes during ripening depends on the fruit species. For example, an increase in catalase activity was found in tomatoes during ripening (Andrews et al. 2004), while a decrease in oranges (Huang et al. 2007) and mangoes (Singh & Dwivedi 2008). In guavas, the activities of antioxidant enzymes, including catalase, reached maxima at the stage of color change and decreased at later stages (Mondal et al. 2009). This variability in the behavior of antioxidant enzymes can be attributed to differences in the activity of isoenzymes (Resende et al. 2012).

Both catalases and peroxidases play a role in the destruction of H₂O₂ but they use different mechanisms that are competitive. When the concentration of H₂O₂ in the cell is high, the catalase is highly active. Conversely, when the H₂O₂ concentration is low and the reducing agent is present, the peroxidase is highly active (Mui 2001). Accordingly, unlike the change in the activity of the catalase, the activity of the peroxidase increased continuously during the fruit growth and development. From week 3 to week 11, peroxidase activity increased slowly (from 0.097 UI·g⁻¹·s⁻¹ to 0.226 UI·g⁻¹·s⁻¹); this was a period when the high concentration of H₂O₂ in the cell was lowered mainly by catalase activity. From week 11 to soft ripening (week 16), the peroxidase activity, reaching 0.858 UI·g⁻¹·s⁻¹, was about four times higher compared to the data obtained at week 11. This result may be related to a rapid tannin degradation, synthesis of several aromatic ring compounds and ethylene during the fruit ripening, which are catalyzed by peroxidase (Pandey et al. 2017).