*2.6. In Vivo Study*

#### 2.6.1. Bioefficacy in Pot House Conditions

To determine the bioefficacy of polymeric conjugated nanoparticles in controlling early blight and leaf spots in tomatoes (*Lycopersicon esculentum* L.), the study was conducted in pots filled with sandy soil in a glasshouse in natural light and temperature.

#### 2.6.2. Treatment of Seeds and Disease Detection

Seeds were properly rinsed and treated with 4% sodium hypochlorite for 10 min before being thoroughly cleaned. The seeds were dipped in CMC (carboxymethyl cellulose, 5.0 g in 100 mL DDW) for 10 min and air-dried. The seeds were then air-dried after being treated with conjugated nanoparticles (10 ppm) for 2 h and 30 min. Five tomato seeds per pot were planted in pots filled with soil (pH 7.7 at 20 ◦C) infected with pathogenic fungus [13]. The forty-day-old plants were sprayed with 15 mL of aqueous conidial solution (3.1 × 107 CFU/mL) of specific pathogens and covered with clear plastic bags to maintain the humidity essential for disease outbreaks. Following the disease outbreak, a foliar spray of CSGA NPs (10 ppm and 15 mL/pot) was used to test the bioefficacy. As a positive control, commercial mancozeb was employed.

Disease severity (DS) was recorded randomly in the standard grade of 0–5 before the polymeric NP spray.

```
Disease severity (%DS) =
(Sum of all disease ratings)/(Total plants assessed × maximum rating scle) × 100 (3)
```
The disease control efficacy (% DCE) was calculated after the polymeric NP spray using the formula in [25].

For the overall health and vitality of the test plant, bioefficacy was measured using plant development characteristics, such as plant height, root–shoot ratio, and dry biomass. The dry weight of each tomato per plant was determined by placing the entire plant with roots in a brown envelope (3 plants per envelope) and drying it for seven days at 40 ◦C in a hot air oven.

## *2.7. Statistical Treatment of Data*

All experiments were executed in triplicate, and the results are presented as mean ± standard deviation (SD). Statistical variances among sets were determined using one-way ANOVA. The statistical significance was accepted at a level of *p*-value ≤ 0.05 by a *t*-test. To handle statistical data, Microsoft Office Excel 2013 was utilized (Microsoft Corporation, Albuquerque, NM, USA).

## **3. Results and Discussion**

#### *3.1. Nanoparticle Size Optimization, Stability, and Physicochemical Characterization*

The quantity of chitosan and gum acacia used in the experiment affected the particle sizes of the NPs in the initial trials. The optimization graph shows that the particle size increases with increasing chitosan and gum acacia concentrations (Figure 1a). In the case of chitosan, however, the impact is more pronounced.

**Figure 1.** (**a**) Optimization of the concentration of gum acacia and chitosan for the particle size by response surface methodology (RSM); (**b**) mancozeb-loaded NP size; (**c**) mancozeb-loaded NP zeta potential.

The average diameter of blank NPs was 322.2 ± 0.9 nm, a 1.00 ± 0.1 PDI, and a zeta potential of −23.2 ± 0.08 mV. The size of NPs grew in response to increasing mancozeb concentrations (Table 1).


**Table 1.** CSGA NP size, PDI, and zeta potential at native pH; freshly prepared, and their storage stability after 20 days in DDW at 4 ◦C.

Mean ± standard deviation in replication of three.

The molecular weight, degree of deacetylation of the chitosan employed, the stirring speed, and time determine the size of the NPs [26]. A single strong peak at 403.7 ± 0.7 nm was detected for CSGA-1.0 (NPs containing 1.0 mg/mL mancozeb) with a zeta potential of −6.99 ± 0.5 mV (Figure 1b,c). Zeta potential values show the stability of NPs up to ±30 mV [27]. The nanocomposites formed were noticed in the synthesis mixture as a white, foggy haziness that settled at the bottom of the flask.
