S. tuberosum periderm biomass was obtained from the canteen of Delhi Technological University (DTU) (Figure 1). The processing involved washing the periderm and subsequently drying them at 40°C. A constant weight was achieved using Matrix Scientific Instrument. To achieve biomass size reduction in the range of 0.5–1 mm, a physical pretreatment involving grinding was applied.

The bacterium used in this research, Pseudomonas putida 2475, was procured from the Microbial Type Culture Collection (MTCC) in Chandigarh, India. Upon procurement, a mother culture was established on nutrient agar medium composed of the following components per liter: beef extract (0.1 g), yeast extract (2 g), peptone (5 g), NaCl (5 g), and agar (15 g), The pH of the medium maintained to be 7.

S. tuberosum periderm biomass is composed of various components, including starch, cellulose, lignin, protein, cellulose, and hemicellulose. To release fermentable sugars from these polysaccharides, an appropriate treatment strategy is necessary before subjecting the biomass to fermentation. Thermochemical treatment involving dilute acid at high temperatures is employed to convert the carbohydrates into monomer units and enhance sugar yield. For the optimization of sugar yield, a chemical treatment was carried out using varying HCl concentrations (0.25, 1.15, and 2%), and times (15, 55, and 95 min) at autoclave condition (121°C/15psi). For acid hydrolysis stock solution of 10% HCl (3.2 M) was prepared and then working solutions of 0.25%, 1.15%, and 2% of acid concentrations were prepared from it. Reaction volume was taken 1L and biomass loading was 100 g in 1,000 mL of aqueous phase in every single combination. After cooling centrifugation was done to separate liquid hydrolysate from unhydrolyzed solid part. After centrifugation liquid hydrolysate was subjected for detoxification with dry calcium carbonate until the pH of sample reaches up to neutral value (6.8–7).

The chemical composition of S. tuberosum periderm, including proximate analysis (total solid, moisture, and ash), was determined using techniques developed by the National Renewable Energy Laboratory (NREL) (Fatmawati et al., 2023). Briefly for total solid analysis 1 g biomass was kept in oven at 105°C till constant weight achieved. For the ash content analysis, the sample was subjected to a temperature of 575°C in a muffle furnace for a duration of 6 hours. Subsequently, the crucibles were taken out directly from the furnace and transferred into a desiccator, where they were allowed to cool for a designated period. Following this, the crucibles and ash were weighed and analyzed gravimetrically using prescribed formula %   A s h = w e i g h t   c r u c i b l e   p l u s   a s h − w e i g h t   c r u c i b l e ÷ O v e n   d r y   w e i g h t   s a m p l e × 100 .

Starch and total phenolics were quantified spectroscopically, following methods found in the literature (Lafont-Mendoza et al., 2018). In the starch analysis method 1 mL sample was taken in a test tube and 0.33 mL of potassium iodide (KI) solution was subsequently added. The blue color intensity was measured at 600 nm using spectrophotometer against KI blank. The amount of starch in the sample was determined by the standard curve (1 mg/mL). For total phenolics 1 gm sample was subjected to methanol extraction (1:10) and then subjected to incubation with Folin reagent (0.5 mL) and sodium carbonate (1 mL) for 2 h at room temperature and then subjected to spectrophotometric assay at 765 nm against the gallic acid standard (1 mg/mL) (Samarin et al., 2012). Cellulose and hemicellulose content was estimated using the chlorite method. Briefly holocellulose content was determined using treatment with sodium chlorite. 17.5% NaOH was used to extract the cellulose compound from the holocellulose. Hemicellulose was determined by subtracting the value of cellulose from holocellulose (Zhou et al., 2015).

After dilute acid treatment, Hydrolysate was used for reducing sugar analysis by the Dinitro salicylic acid method briefly 1 mL of extracted hydrolysate with equal amount of DNS solution (3,5 dinitro salicylic acid 1 gm, Na₂SO₃ 0.05 g, sodium potassium tartarate 18.2 g, NaOH 1 gm, and phenol 0.2 gm per 100 mL of H₂O) were subjected to heating for 10 min at 100°C and optical density at 540 nm was recorded upon cooling using standard glucose curve (1 mg/mL). Total reducing sugars (glucose, fructose, and xylose) present in the biomass has been considered for estimation (Shangdiar et al., 2023). S u g a r   r e c o v e r e d   m g m L = V o l u m e   o f l i q u i d   h y d r o l y s a t e   r e c o v e r e d ÷ A m o u n t   o f   s a m p l e   t a k e n   f o r   h y d r o l y s i s × 100

Functional group analysis of both raw and acid hydrolyzed S. tuberosum periderm waste was conducted using Fourier-transform infrared (FTIR) spectroscopy with KBr pelleting (PEIR SUBTECH SPECTRUM ASCII PEDS 4.00). Thermogravimetric analysis (TGA) of the S. tuberosum periderm was carried out to observe decomposition peaks at various heating rates under an inert nitrogen environment, using a PerkinElmer analyzer. The analysis covered temperatures ranging from 30°C to 900°C with a heating rate of 10°C min⁻¹. The surface structure of both raw and treated S. tuberosum periderm biomass was observed using a Scanning Electron Microscope (ZEISS EVO 18) at a beam accelerating voltage of 20 kV.

Pseudomonas putida (strain 2475) was procured from MTCC, Chandigarh, India. The culture was periodically revived by transferring it to fresh Luria Bertani media (comprising 5 g/L yeast extract, 10 g/L NaCl, 15 g/L agar, and 10 g/L tryptone) and storing it at a low temperature of 4°C. To develop the inoculum, bacterial colonies from the pure culture were transferred to 150 mL of nutrient broth and incubated at 30°C with continuous shaking at 180 rpm for 24 h. Initially, we assessed the growth pattern of P. putida by measuring its optical density at 600 nm using a GENESYS 50 UV-visible spectrophotometer from Thermo Fisher. This assessment was conducted in mineral salt media containing Na₂HPO₄·7H₂O (30 g/L), KH₂PO₄ (15 g/L), NaCl (5 g/L), NH₄Cl (1 g/L), MgSO₄ (2 g/L), CaCl₂ (0.1 g/L), and synthetic glucose (10 g/L), as the carbon source. For nitrogen limitation, the quantity of NH₄Cl was reduced to 1 g/L. In media containing hydrolysates, 0.1% tryptone was used as nitrogen limitation.

For inoculation, 1% of the freshly prepared seed culture, prepared in Luria Bertani media with an optical density of 0.8 and a concentration of 312 × 10⁶ colony-forming units (cfu) was employed. The study involved three different media conditions for PHA production, each conducted in 2 L conical flasks containing 1 L of modified medium: (A) Modified medium comprised a specified volume (1%) of acid treated liquid hydrolyzed S. tuberosum periderm hydrolysate at a concentration of 10 g/L (after neutralization with calcium carbonate), 5 g/L NaCl, and 1 g/L tryptone. Incubation was carried out in a shaking incubator (180 rpm) at 30°C for 24, 36, 48, and 72 h. (B) Pure hydrolysate medium at a concentration of 10 g/L. (C) Mineral salt medium, as described in the literature (Sriyapai et al., 2022). PHA production values were averaged after each experiment, which was conducted in duplicate.

After incubation, the culture broth was subjected to centrifugation at 10,000 rpm using an Eppendorf 5810 centrifuge for 15 min at 4°C. Following centrifugation, the supernatant was carefully drained, and the cell pellet was recovered. Recovered cell pellet was subsequently washed sequentially with 25 mL deionized water, 25 mL acetone, and 25 mL ethanol, and then dried at 40°C until a constant weight was achieved. This recovered cell biomass, referred to as cell dry mass, was determined gravimetrically, as per the method outlined in the literature (Tanikkul et al., 2020b). Polymer extraction was carried out using the chloroform extraction method with slight modifications, following the procedure described by Filippi et al. (2021). 2.1 g cell pellet was crushed with the help of mortar pestle and then incubated in 20-fold chloroform (43.8 mL) at 60°C in a water bath for 2 h. The resulting mixture was filtered using Whatman filter paper (1), and the liquid was concentrated through evaporation. PHA was obtained by precipitation of whole recovered solution obtained after heating with 22 mL ice-chilled methanol and subsequently analyzed using GC-MS, following the previously established protocol (de Souza Reis et al., 2020). Briefly For the analysis of monomer composition, we performed GC MS-MS. First, 20 mg of dry cell biomass was subjected to acidic methanolysis. Methyl esters were prepared by combining 15% sulfuric acid in 85% methanol (2 mL) with chloroform (2 mL) in a screw-capped culture tube and heating it at 100°C for 2 h and 20 min (de Souza Reis et al., 2020).

After cooling, 2 mL of water was added to separate the organic layer from the aqueous layer. The GC-MS-MS analysis was conducted using the Triple quadrupole 7000D GC/TQ Agilent, equipped with a triple-axis detector. For the analysis, PHA was used in the form of hydroxy alkanoic acid methyl esters, following the method reported by Mahato et al. (2021). A sample of 2 μL at a split ratio of 1:50 was automatically injected into the GC and injection temperature was 200°C. Helium served as the carrier gas at 48 mL⁻¹min and 0.42 bar pressure (Hierro-iglesias et al., 2023). Monomer identification was performed using the NIST 17 library.

The extracted PHA film was prepared in a fume hood by dissolving 200 mg extracted PHA in 20 mL of chloroform at room temperature and subsequently pouring the solution into Petri dish till complete evaporation of solvent. To determine the melting temperature of the PHA film, we conducted Differential Scanning Calorimetry (DSC) using a temperature-regulated system (DSC 8000, Perkin Elmer). We took 5 mg of extracted PHA in an aluminum pan and subjected it to a nitrogen flux rate of 10 mL/min, heating it from 30°C to 250°C, following the procedure outlined in the literature (Kacanski et al., 2023) with some modifications.

For the analysis of functional groups, we recorded Infrared (IR) spectra in the range of 4,000 to 450 cm⁻¹. This was done by preparing KBr pellets and using a (Perkin Elmer Frontier Shelton CT08484). In brief potassium bromide (KBr) was subjected to drying at 110°C for 2 h for moisture removal. 5 mg sample was mixed with 200 mg of powdered KBr. After pulverization, mixture (sample and KBr) placed in to pellet forming machine and resulting pellet was taken for IR analysis (Cerrone et al., 2023). To analyze the chemical shift of the PHA, we utilized 1H Nuclear Magnetic Resonance (NMR) by dissolving a 5 mg sample in 600 μL of CDCl₃, employing a (BRUKER Proton NMR 500 MHz), as described by Kacanski et al. (2023). The chemical shift of PHA was recorded in parts per million (ppm).

The composition analysis of S. tuberosum periderm collected during winter was conducted, and the findings are presented in Table 1. One gram of the sample was used for composition analysis, and the results were reported in triplicate.

The composition of the S. tuberosum periderm was as follows: starch 64.44%, ash 14%, total lignin 6.5%, phenolics at 0.6% of S. tuberosum periderm, cellulose 9.01%, hemicellulose 1.89%, and the total solid content was found to be 90% (w/w).

In comparison, Malakar et al. (2020) reported 76% moisture and 8% ash in S. tuberosum periderm, Arapoglou et al. (2010) reported starch content at 52.14%, Lima et al. (2021) reported cellulose and hemicellulose content ranging from 10% to 30%, and Samarin et al. (2012) reported a total phenolic content of 522 µg GAE/g dry weight. These findings support the results of our study. However, it is worth noting that the composition of S. tuberosum periderm can vary among different Solanum tuberosum species (Sampaio et al., 2020).

Different concentrations of dilute acid and time variation for hydrolysate development were used in this study which resulted in a varied amount of reducing sugar and their combined severity factor (LogR_o). Based on the above parameters, results presented in Table 2. From the obtained results it can be concluded that very low acid concentration (0.25), is unable to liberate high sugar at any time. Briefly 0.25% acid at 15 min extracted 0.05% reducing sugar. However, at 55 and 95 min sugar yield was 5.49% and 16.45%, respectively. Center point (1.12% acid and 55 min time) was obtained through the software and the sugar yield in all the five runs was between 59% and 63%. While High acid concentration (2%) releases the highest amount of sugar such as 76% at 15 min, and 70% at 95 min. From the above observations it can be said that sugar yield increases with the increase in acid concentration. However, time is not very significant factor in terms of sugar removal as 2% acid at 15 min can extract more sugar as compare to 95 min. The reason behind the diversity in sugar removal is high acid concentration and longtime combinedly generate toxic metabolites which leads to degrade sugar molecule. Therefore, a comparatively low acid concentration was optimized from response surface methodology the for maximum sugar with low toxicity. Further the optimized condition was used for acid hydrolysis.

This study was designed to establish a linear equation based on thirteen experimental runs, including five center points, all subjected to analysis of variance (ANOVA). Two factors, time and acid concentration, were optimized through Central Composite Design (CCD) and further analyzed using regression analysis and analysis of variance. The independent variables and their respective values are presented in Table 3.

Following the completion of thirteen trial runs, the optimal conditions for achieving the highest yield of reducing sugars were determined using Minitab software and graphically represented in Figure 2. The Pareto chart illustrates the standardized effects of individual factors: (A) acid concentration, (B) time, and the combined effects of both factors. The Pareto chart indicates that factor (A) exerts the most significant influence on sugar extraction, while the impact of time is relatively negligible. Furthermore, the combined influence of acid and time explains a smaller portion of sugar extraction.

The contour plot displays the region of maximum sugar yield, falling within the ranges of 1.5%–2% for acid concentration and 50–90 min for time. Based on above observations, this study was aimed to exact the maximum sugar at mild acid concentration. At 2% acid concentration and after 95 min of hydrolysis time, sugar was observed to be 70.80 ± 0.27, however, the acid concentration and time was a bit higher. While at 2% acid and 15 min of time sugar yield was maximum (76%) but acid concentration was higher. Hence, the maximum sugar liberation and milder condition were established to obtain optimized condition. The predicted optimal conditions from the model were determined for 1.75% acid concentration and 50 min of reaction time, resulting in a predicted sugar yield of 70.47%. To validate the accuracy of the predicted response, a hydrolysis experiment was conducted under the specified parameters, and the trials were conducted in triplicate. The average sugar yield obtained was 69.34% ± 0.25%, which closely matched the predicted value. Further this extracted sugar used as a carbon source for PHA production by P. putida in through this study.

The combined severity factor (LogR_o) under the optimized conditions was calculated to be 1.99. In a study, 5% hydrochloric acid, yielding 62 g L⁻¹ of sugars (Khanal et al., 2023). In a two stage hydrolysis study, 32 g of potato peel in 400 mL of 1% w/w H₂SO₄ at 180°C for 60 min was performed, For second stage hydrolysis with cellulase, α-amylase and glucoamylase 36 g L⁻¹ glucose was obtained (Abedini et al., 2020). In another two-step hydrolysis process of potato peels, 141 g L⁻¹ of sugar obtained with 3% w/w H₂SO₄ in an autoclave condition for 15 min, The second stage enzymatic hydrolysis was done with the crude enzyme complex produced by Aspergillus sp. For 72 h at 50°C and 150 rpm (Soni et al., 2023). In an experiment, potato peel waste underwent pretreatment with 1% H₂SO₄ at 121°C for 30 min, with a substrate loading of 10%. In the subsequent hydrolysis step, 10 g of the pretreated substrate were exposed to enzymatic hydrolysis using cellulase, hemicellulose and amylase (at concentration of 30 U, 5 U, and 70 U/g of substrate, respectively). This hydrolysis process occurred at 50°C in a rotatory shaker incubator, with agitation at 700 rpm for 48 h. The outcome of this process revealed the presence of 77 g/L of reducing sugar (Kalafat et al., 2018).

In the current study, we achieved a considerably high yield of reducing sugars at 1.75% HCl, which is milder in terms of the generation of toxic byproducts such as furfural and hydroxymethylfurfural. Details regarding the responses after the trial runs and their Coefficient of Standard Deviation (CSD) are provided in Table 2.

RSM (Response Surface Methodology) was employed to assess the combined impact of two independent variables: Acid concentration (X ₁ ) and Time (X ₂ ), From the numerical data obtained in this study, the simplified linear regression model for sugar yield from S. tuberosum periderm can be expressed as follow ‐ 24.19 + 93.8 X 1 + 0.1758   X 2   –   25.54   X 1 × X 1 where X ₁ = Acid concentration (%) and X ₂ = Time (min).

The regression analysis of the trial runs yielded a statistically significant result, with an R-squared value of 93.14%, indicating a strong fit of the model. Additionally, the lack-of-fit for the model was found to be negligible. The output of the regression analysis for glucose yield is presented in Table 2, where a p-value greater than 0.05 indicates the significance of the model.

To create the best-fit model and eliminate irrelevant conditions (those with p-values greater than 0.05), a linear equation for regression was applied. So, a model for the optimization of reducing sugar yield from S. tuberosum periderm was successfully generated. Based on obtained values, the sequence of relevance for the two factors on acidic hydrolysis was acid concentration > time.

The counterplots and Pareto chart were used to visually depict the effects of individual variables and the combined interactions of acid concentration (X₁) and Time (X₂) on sugar yield.

These statistical analyses and graphical representations are valuable tools for understanding the impact of the variables on the hydrolysis process and optimizing the yield of reducing sugars from S. tuberosum periderm waste.

Based on the GC-MS-MS analysis, using S. tuberosum periderm waste hydrolysate as the sole carbon source resulted in the formation of medium-chain length polyhydroxyalkanoates (mcl PHA) containing three distinct monomers: 3-hydroxydodecanoate (3HDD) with a retention time (RT) of 6.99 min and a peak area of 1.75%., 3-hydroxydecanoate (3HD) with an RT of 10.34 min and a peak area of 1.72% and 3-hydroxytetradecanoate (3HTD) with an RT of 8.52 min and a peak area of 19.2% (Figure 5B).

These identified monomers demonstrate the feasibility of producing industrially important mcl PHA with carbon sources of low cost, such as S. tuberosum periderm hydrolysate. This approach enables the construction of mcl PHA with chain lengths ranging from 5 to 14 carbons, making it an economically attractive option.

This finding aligns with the work of Mahato et al. (2021), who reported the presence of beta-3-hydroxybutyric acid (7.41%), 3-hydroxytetradecanoic acid methyl esters, and 3-hydroxyhexadecanoate (5.03%) in PHA produced by Pseudomonas aeruginosa. However, it is important to note that their PHA production was observed in mineral salt media containing different oils as carbon sources, which may not be as sustainable and could potentially incur higher costs compared to the approach used in the present study. In a study Ciesielska et al. (2017) reported mcl PHA monomers such as 3- hydroxyhexadecanoate and 3- hydroxyoctadecanoate from Pseudomonas putida KT2440 using oleic acid as a carbon source.

By comparing PHA monomers produced by Pseudomonas from different studies it is evident that PHA extracted from P. putida MTCC 2475 (present study) is mcl PHA.

The PHA film was created by adding extracted PHA in solvent (chloroform) and stored securely under aseptic conditions at room temperature (Figure 5A). The Differential Scanning Calorimetry (DSC) thermogram presented in Figure 5D reveals the melting temperature (T_m) of the produced Polyhydroxyalkanoates (PHA), which was found to be approximately 170°C. This temperature closely aligns with the melting temperature of 3-hydroxydecanoic acid (166°C), as reported in the literature (Sharma et al., 2017).

The identification of functional groups within the extracted PHA was accomplished through Fourier Transform Infrared Spectroscopy (FTIR), as illustrated in Figure 5C. The spectral pattern of functional groups closely resembled findings from Mahato et al. (2021), who identified different forms of decanoic acid methyl esters, including Methyl 3-hydroxytetradecanoate and hexadecanoic acid methyl ester, from Pseudomonas aeruginosa. Notable absorption bands included: 1) A band at 3,437 cm⁻¹, attributed to the O-H stretching of the hydroxyl group in PHA. 2) The band at 2,929 cm⁻¹, assigned to the CH₂ group. 3) An absorption band at 1,733 cm⁻¹, indicating C=O stretching vibration, confirming the presence of an ester bond in the PHA monomer. 4) A band at 1,277 cm⁻¹, associated with the asymmetric C–O–C stretching vibration.

Furthermore, the Proton Nuclear Magnetic Resonance (NMR) spectrum of the PHA film is depicted in Figure 5E. The complex multiple resonance bands between 1.26 and 1.33 ppm (peak no. 1) suggest the presence of methyl groups (–CH₃) in the PHA film. Multiple resonance spectra observed between 2.17 and 2.60 ppm (peak no. 2) confirm the existence of methylene groups (–CH₂). The band position ranging from 5.22 to 5.27 ppm (peak no. 3) indicates the presence of methane (–CH) in the PHA polymer. Peak no. 4 corresponds to the solvent CDCl₃. These observed peaks align with prior studies on medium-chain length PHA produced by Pseudomonas strains, as reported by Gumel et al. (2014). Comparing the results of the present study with findings from various previous investigations, it can be confidently affirmed that the extracted polymer is indeed medium-chain length Polyhydroxyalkanoates (PHA).