Changes in the Chemical and Physical Properties of Untreated and Finished Polyamide 6.6 Fabrics Buried in Different Soil Matrices, from the Lab-Scale to a House Garden

Abstract

This paper aims to analyze the biodegradation behavior of a common synthetic fiber, well-known for its environmental recalcitrance: polyamide 6.6. In particular, polyamide 6.6 fabrics finished with chitosan to impart antibacterial properties and the natural red dye carmine were studied. Fabrics of standard polyamide 6.6 served as references. Some specimens were buried in compost-enriched soil for 1, 2 and 3 months and kept in the laboratory; simultaneously, others were placed in an outdoor house garden to simulate landfill conditions. After each sample withdrawal, various characterization techniques were employed to assess the status of the fibers. The first evidence was that, in general, there were no weight changes or significant macroscopic damage within three months, except for white stains as an index of microorganism colonization, which was confirmed by microscopic analyses, where bacteria and fungi could be clearly seen. On the one hand, some effects were revealed during the burial in the house garden that impacted the fabrics’ surface characteristics in terms of interaction with soil derivatives (susceptibility to adsorption of water and soil-derived substances). On the other hand, the samples buried under laboratory conditions showed a weak antibacterial efficacy, leading to the hypothesis that more aggressive degradation may have occurred at the expense of chitosan. Still, three months of burial led to mild surface deterioration, opening possibilities for further research.

1. Introduction

Polyamides, also known as nylons, are linear synthetic polymers made of monomers connected with amide bonds (-CO-NH-) via a polycondensation reaction [1]. Together with several other polymers (polyesters, acrylics, etc.), constituting 60% of global textile manufacturing, polyamides are widely produced synthetic textile materials, reaching a production of about 7 million tons per year [2,3].

Polyamides can be classified into diverse types depending on the number of carbon atoms between the amide bonds. The most common are nylon 6 and 6.6 [4,5,6,7], but nylons 4, 6.9, 6.10, 11, and 12 also find applications in various sectors [8,9], like the automotive, electronics, and medical fields [10,11,12,13]. Actually, polyamides are thermostable matrices that exhibit good wear durability and fatigue resistance, but, on the other hand, they undergo biodegradation very slowly [14,15]. Polyamide 6.6, in particular, is a semicrystalline polymer synthesized by the reaction between a diamine (hexamethylenediamine) and a diacid (adipic acid) [16,17], and it is so named because of the six carbon atoms in the diamine and diacid that characterize its chemical structure [18]. Since it possesses high mechanical strength and thermal resistance, flexibility, durability, good dyeability, and resistance to shrinkage and chemicals, it is largely employed in the production of synthetic apparel, especially technical textiles such as sportswear, and in furnishings [19,20,21,22]. For these multiple uses, polyamide fibers are subjected to finishing treatments, such as dyeing. With acid dyes, for instance, the reaction between the amino-terminal groups of the fibers and the sulfonic groups of the dye can take place [23], but reactive dyes can also be employed to establish covalent bonds, while disperse dyes can exploit hydrophobic interactions [24,25]. Despite an important recourse to synthetic dyes, eco-friendly and natural-based substances are objects of research in the field of sustainable finishing [26,27,28,29]. Among others, a successful finishing was previously discussed by this research group in [30], where polyamide 6.6 fabrics were functionalized with chitosan, a biopolymer able to increase the dyeability degree of the substrate with a natural dye, Carmine Red (hereafter referred to as “carmine”). This textile treatment based on chitosan also brought about effective antibacterial activity against both Gram-positive and Gram-negative bacterial strains.

Given polyamide 6.6 products’ widespread use and their recalcitrance in the environment due to the high intermolecular strength of hydrogen bonds and resultant difficult degradability [31], a current problem is developing strategies for limiting their permanence and impact on the ecosystem after disposal [32,33], especially given the exponential growth of the fast-fashion model, which generates an unmanageable amount of synthetic textile waste [34]. Among the possibilities to achieve this critical goal, there is the exploration of alternative compounds that can replace the most common polyamide precursors, often petroleum-derived, with bio-based ones [35]. For instance, polyamide 5.6 can be prepared with a diamine monomer derived from biological raw materials of plant origin. Gülel and Güvenilir [36] deepened the research on this compound by seeking a method to accelerate its biodegradation pathway. They incorporated olive stone powder at variable concentrations into the synthesized polyamide and evaluated its biodegradability by carrying out a six-month soil burial test. The sample filled with the concentration of 10 wt% of olive stone powder registered the lowest tensile strength and a high elastic modulus due to changes in the crystalline structure promoted by the bio-based additive, and the soil burial test indicated a weight loss of 5.24%, providing evidence of some biodegradability.

However, the present concern is the management of polyamides that have already been fabricated but were not designed with a “from cradle to grave” approach, and therefore, are not biodegradable. In this direction, Gashti et al. tested the degradation of polyamide 6.6 by soaking the fibers in solutions formed by lipolytic and proteolytic enzymes at diverse concentrations. The combination of both enzymes determined a relevant sample weight loss [37]. Meng and coauthors proposed a new study focused on the decomposition of polyamide 6.6 in supercritical water [38] using a specific apparatus and setting the temperature (380 °C), pressure (28 MPa), and time (30 min) to achieve the desired reaction conditions. A complete decomposition of polyamide 6.6 was observed. The acidic features of water promoted the hydrolysis of polyamide; in this way, the starting material was converted into monomers and other products.

Another strategy can be represented by the exploitation of microorganisms of fungal and bacterial origin to initiate the break process of polyamide substrates. In this direction, different studies have focused on nylon 4 since it can be more easily degraded in a natural environment compared to other polyamides [13]. Yamano and colleagues [14] discovered that a specific bacterial strain of Pseudomonas sp. was capable of producing extracellular hydrolytic enzymes that promote the degradation of polyamide 4. They also investigated a new method to monitor the in vivo biodegradation process of polyamide 4 by implanting variable samples (non-woven fabrics, film, and mold) in the backs of rats [39]. A recent article [40] analyzed the microbial degradation of polyamide 4 in different types of soil and how the bacterial community changes during the degradation process. The most effective degradation occurred in composted soil, where a single bacterium strain called NR4 was able to cause a degradation rate of around 65% of polyamide 4 within 15 days. Similarly, Tachibana et al. [41] identified, through the analysis of the DNA of the soil microbial community, two diverse microbial strains responsible for polyamide 4 degradation in the composted soil: a bacterium species (Stenotrophomonas) and a fungal species (Fusarium). The hypothetical mechanism was based on their ability to recognize the acyllactam or carboxy-type chain end of polyamide 4 and start the degradation process.

In [42], polyamide 6 fibers were used as the unique source of nitrogen to evaluate the behavior of a filamentous and lignolytic fungus species: Phanerochaete chrysosporium. The physical and chemical changes in the specimens were monitored for 5 months. After only 3 months, the molecular weight had decreased by 50%, and notable fiber damage was visible, progressing from an intrinsic smooth surface to deep grooves. Another approach was adopted in the work of Shi et al. [43]: polyamide 6 pellets were exposed to variable outdoor conditions (in terms of temperature, humidity, and rainfall) and collected after different time intervals (1, 3, 6, 12, 18, 24, 30, and 36 months). The outdoor aging behavior revealed that some factors, such as UV irradiation, heat, ozone, and microorganisms, had a major impact on accelerating the samples’ aging.

A significant aspect to consider is the scale of the specimens that can undergo microbial biodegradation, ranging from relatively large samples to microplastics. The latter are a source of environmental and health concerns due to their small size (<0.5 cm) and resistance to degradation. It must be considered that annually, 3.45 million tons of nylon 6.6 microplastics are produced and released into the environment [44]. In the paper by Chauhan et al. [45], polyamide 6.6 particles 100–200 nm in size were subjected to the action of a bacterial consortium, thus implying a combination of diverse microorganisms. Specifically, three strains, Brevibacillus brevis, Achromobacter xylosoxidans, and Acinetobacter baumannii synergistically produced laccase and peroxidase, which were able to break down the polymer into smaller oligomers and monomers. The same enzymes responsible for reducing polymer chain length were also found in another study [46], where the degradation of nylon 6.6 microplastics was studied through bacterial cultures isolated from a landfill site.

To make the research work more realistic and reproducible in an actual context, one way is to compare the obtained results in a lab-scale condition with the same experimental design translated into a real environment. In this regard, a survey focused on the degradation of polyamide 4 has been conducted in both a closed setting (using sampled seawater) and in an open-air habitat (the sea) [47], since the accumulation of synthetic fabrics can also occur in aquatic environments. The findings suggested a biodegradation activity promoted by the hydrolysis of a specific MND-1 bacterial strain belonging to the family Alteromonadaceae. A study on the degradation of polyamide 4 reported its transformation into gamma-aminobutyric acid oligomers mediated by the action of marine bacterial enzymes [48]. Moreover, a comparative analysis [49] was performed to investigate the degradation of nylons 6 and 6.6 in the marine habitat after 1 month and 3 months. In particular, marine microbes such as Bacillus cereus, Vibrio furnissii, Bacillus sphericus, and Brevundimonas vesicularis promoted the polymers’ biodegradation under marine salt medium. After 90 days, a decrease in molecular weight of 42% and a weight loss of 7% for nylon 6.6 were registered, while for nylon 6 these parameters resulted in values of 31% and 2%, respectively.

Here, following a previous work focused on the biodegradation behavior of finished cotton in compost-enriched soil [50], the aim was to assess the ability of polyamide 6.6 to undergo biodegradation both in lab-scale conditions and in a real outdoor environment (soil house garden). The specimens involved in this research were a bare polyamide 6.6 fabric and a sample finished with chitosan and carmine. As previously mentioned, the latter is frequently used as a natural dye in textile applications because it is easily obtained by crushing the Cochineal insect, whereas chitosan is a biopolymer with intrinsic biodegradability and antimicrobial properties. The rationale behind the biodegradation tests on this kind of finished synthetic fabric was related to a possible higher interaction and favorable attachment of microorganisms to a bio-based coating, despite the intrinsic non-biodegradability of the polyamide substrate.

2. Materials and Methods

2.1. Reactants and Fabric Preparation

Adjacent polyamide 6.6 fabric (plain weave fabric suitable for ISO 105-F03, mass per unit area 130.0 g/m² determined in accordance with ISO 3801) was supplied by Testfabrics Inc. (West Pittston, PA, USA). The polyamide fibers’ diameter, measured with the Optical Fibre Diameter Analyzer OFDA 2000 instrumentation (OFDA, Ardross, Australia), was 13.98 ± 1.66 μm. Before the analysis, the yarns were withdrawn from both the warp and weft directions and cut with a microtome. Four samples of each fabric were analyzed, and the mean value was then obtained.

Low-molecular-weight chitosan (50–190 kDa) and glacial acetic acid (99.8–100.5%) were purchased from Sigma-Aldrich (Milan, Italy). Carmine dye (E120, C.I. Natural Red 4, C.I. 75470) was kindly supplied by Aromata Group srl, Bresso, Italy.

Polyamide 6.6 fabrics were left in their original form or finished with chitosan plus the dye carmine, following a previously described method [30]. In brief, polyamide 6.6 fabrics were dipped into an acid solution of chitosan (2% w/w chitosan solution in 2% w/v glacial acetic acid); after an overnight contact period, they were manually padded to reach a 90% wet pick-up. The fabrics were annealed to favor the reaction between the chitosan and the fabric. The chitosan-coated fabrics were dyed with the natural colorant carmine, using a 4% on weight of fiber (owf) dye concentration and a fabric:bath ratio of 1:20. The equipment used for dyeing was an Ahiba Nuance Top Speed II machine (Datacolor Italia srl, Giussano, Italy) operating at 100 °C. The amount of coating was measured by the dry weight of the polyamide samples before and after chitosan functionalization and dyeing treatment. The weight of the fabrics was evaluated after conditioning the samples at 105 °C for 2 h. The coating amount was calculated as the mean of three measurements per sample, yielding 6.5 ± 0.6 wt% chitosan+dye deposited on the fabrics.

2.2. Burial Tests

Samples of 3 × 3 cm were cut, and each fabric piece was buried about 2–3 cm deep below the soil surface and then withdrawn after 1, 2, and 3 months. The first medium used for the biodegradation process was commercial soil containing neutral sphagnum, composted green soil improver, pumice, guano, and mineral fertilizer (COMPO SANA^®, COMPO Italia Srl, Cesano Maderno, Italy) [51]. The pH(H₂O) of such soil was reported as 7. The second medium was the same soil, but sterilized at 105 °C for 24 h to reset the biota loading [51]. For the first and second media, the storage during the burial period took place in a conditioned laboratory (20 °C and 65% relative humidity). To avoid total dryness, both sterilized and non-sterilized soil were moistened twice a week with 10 mL of MilliQ^® water (MilliQ^® system from Merck KGaA, Darmstadt, Germany).

The third medium, simulating a real environment, was a house garden placed in Biella (North-West Italy). To monitor the environmental variables as much as possible, the weather conditions were noted in terms of rain intensity (mm), humidity, and temperature (

Table 1. Weather data during the three-month experiment in the house garden.

Parameter/Period	23 April/22 May	23 May/22 June	23 June/21 July
Mean Tmax (°C)	11.2	15.7	19.3
Mean Tmin (°C)	17.5	22.2	27.4
Mean humidity (%)	69.6	68.3	69.4
Rainfall (mm)	281	139	179.3

, data taken from Ilmeteo.it). Further details on the investigation of soil characteristics are available in Section 2.5.

In each case, after being withdrawn from the soil, excess soil matter from the fabrics was gently removed before characterization.

The samples studied in this work are labeled using “P” for Polyamide 6.6, while “1/2/3” stands for the period (month) of burial, “T” represents the samples dyed with carmine, “S” is for samples buried in sterilized soil, and “G” identifies fabrics kept in the house garden.

2.3. Fabric Physical–Chemical Characterization

The fabrics after burial were first weighed with an analytical balance and multi-analytically characterized, as in [50]. Morphological investigations were performed on gold-coated samples using a Scanning Electron Microscope (SEM, EVO 10 Carl Zeiss AG, Oberkochen, Germany). Differential Scanning Calorimetry (DSC) in the range 30–300 °C was carried out in N₂ (100 mL min⁻¹, 10 °C/min) with the Mettler Toledo 821e calorimeter. Fourier-transform infrared (FTIR) spectra were acquired in the attenuated total reflection (ATR) mode (4 cm⁻¹ resolution, 64 scans) by means of a ZnSe crystal-equipped Thermo Nicolet iZ10 spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). Contact angle and drop absorption time were obtained by the EasyDrop instrument (Krüss Scientific GmbH, Hamburg, Germany) to evaluate the fabric hydrophilicity/hydrophobicity.

The thickness of the samples was evaluated in triplicate through a Digital Dial Indicator (Fowler, Canton, MA, USA), comparing the untreated polyamide and polyamide 6.6+Chitosan+Dye fabrics both before and after burial.

Colorimetric analysis was performed with a Datacolor SF 600 X Spectralflash (Datacolor Italia srl, Italy) with CIE standard illuminant D65, 10°, determining the CIELab ∆E values from Equations (1) and (2) [52]:

$$∆E=\sqrt{{∆L}^2+{∆a}^2+{∆b}^2}$$

(1)

$$∆E=\sqrt{{∆L}^2+{∆C}^2+{∆H}^2}$$

(2)

According to the CIELab color space and Equation (1), “L” denotes lightness, “a” represents the value from green (negative) to red (positive), and “b” goes from blue (negative) to yellow (positive). Therefore, the ∆s refers to the differences between the measured colorimetric parameters and the related L₀, a₀, and b₀ of the reference non-buried samples. Similarly, ∆L, ∆C, and ∆H in Equation (2) represent the changes in lightness, saturation, and hue, respectively, again with respect to the references. The global colorimetric data (∆E) are reported as the average values of ≥3 measurements for each sample. Additional measurements were performed to assess the homogeneity of the coating before burial, testing polyamide 6.6+Chitosan and polyamide 6.6+Chitosan+Dye against bare polyamide 6.6.

2.4. Antibacterial Tests

Since polyamide fabrics finished with chitosan have been previously proven to possess relevant antibacterial properties, in the present work, the eventual depletion of the chitosan layer and, thus, the decrease in antimicrobial action due to burial were evaluated on the samples PT3, PT3S, and PT3G. To this aim, antibacterial trials were performed against the Gram-positive bacterium Staphylococcus aureus (strain ATCC 6538) following the method ASTM E 2149-2013 “Standard test method for determining the antimicrobial activity of antimicrobial agents under dynamic contact conditions” [53]. Briefly, Staphylococcus aureus was grown in a nutrient broth (Buffered peptone water for microbiology, VWR Chemicals) at 37 °C for 24 h. The bacterial concentration was set at approximately 1.5–3.0 × 10⁵ CFU mL⁻¹ working dilution. This inoculum was brought into contact with each antibacterial sample (fabric: inoculum ratio of 1 g: 50 mL) under shaking at room temperature for 1 h. After this time, 1 mL of inoculum was diluted 1000 times and plated on Petri dishes containing yeast extract agar (Sigma-Aldrich, Milan, Italy), which were subsequently incubated at 37 °C for 24 h. After this period, the surviving bacterial colonies were counted and compared to the initial bacterial concentration to determine the percentage of bacterial reduction, according to Equation (3).

$$Reduction \left(\%\right)={\displaystyle \frac{\left(A-B\right)\times 100}{A}}$$

(3)

where A = number of viable microorganisms before treatment, and B = number of viable microorganisms after treatment.

The fabrics that underwent biodegradation required pretreatment to eliminate biota contamination, which could invalidate the antibacterial tests. For this reason, they were sterilized under UV radiation (Osram Ultra Vitalux UV-A 300 W Lamp, OSRAM GmbH, Germany) for 4 h, and a non-buried polyamide 6.6+Chitosan+Dye was subjected to the same treatment to serve as a reference. After UV sterilization, the samples were analyzed using FTIR-ATR to verify that such a treatment did not alter the principal chemical functionalities.

2.5. Characterization of House Garden Soil

As reported in Section 2.2, the commercial compost-rich soil has known compositional characteristics indicated by its label. To overcome the total lack of knowledge of the house garden soil, a survey of its features (moisture, pH, and principal chemical groups) was made. Moisture and pH were determined following the modified Italian law methods [54]. In detail, a 0.5 mm sieved sample (ca. 200 g)—previously stabilized at room temperature to prevent excess moisture due to previous rains—was put in an oven at 105 °C and left overnight. The moisture content was calculated as the difference between the soil weights before and after drying. The pH of the 0.5 mm sieved dried soil was evaluated by adding deionized water to obtain a final ratio of soil to water of 1:2.5. The suspension was left to stir overnight to ensure homogeneity, and the next day, after deposition, the pH was measured in the solution by means of a pH meter.

Moreover, the estimation of inorganic matter was conducted: 80 g of soil were put in a muffle furnace at 800 °C for 5 h to burn the organic matrix (ignition method [55]). Again, the inorganic portion was calculated as the difference between the initial and final weights. Then, both the residues at 105 °C and 800 °C were analyzed by FTIR-ATR (with the same setup described in Section 2.3) to acquire general chemical information.

3. Results and Discussion

3.1. Characteristics of House Garden Soil

The outcomes of soil analysis are summarized in

Table 2. Results related to the parameters used to characterize the house garden soil.

Type of Soil Subjected to Analysis	Parameter	Results
0.5 mm sieved sample	Humidity	12%
0.5 mm sieved sample dried at 105 °C	Inorganic residue estimated after heating at 800 °C	92.5% (resulting in a red ash powder)
0.5 mm sieved sample dried at 105 °C	pH	6.2
0.5 mm sieved sample dried at 105 °C	Functional groups (FTIR-ATR)	Main vibrations at (cm−1): 3695, 3618, 3420, 1634, 1410, 1350 (shoulder), 1024 (shoulder), 995, 910, 795, 776, 747, 690.
0.5 mm sieved sample after treatment at 800 °C	Functional groups (FTIR-ATR)	Main vibrations at (cm−1): 1000, 777, 719, 694.

. First, the pH (slightly acidic) falls within a range considered normal, for instance, for plant growth [56]; thus, this acidity level could hardly have affected the chemical degradation of the polyamide fabrics considered in this study or have caused an important coating leaching in the case of polyamide 6.6+Chitosan+Dye samples.

Regarding this latter aspect, indeed, it was previously proven that the adopted finishing is resistant to washing with detergents [30].

The FTIR-ATR spectra of the soil dried at 105 °C showed two sharp peaks at 3695 and 3618 cm⁻¹ attributable to ν(O–H) and ν(Si)O–H of clay minerals, while the adjacent broad band at 3420 cm⁻¹ subtended the ν(O–H) vibrations of phenolic substances together with entrapped H₂O [57,58,59,60]. The presence of water was also suggested by a significant signal at 1634 cm⁻¹, namely δ(H–O–H). Nevertheless, superimposed with this peak, the contribution at 1634 cm⁻¹ may also belong to organic moieties and can be imputed to the stretching of C=C aromatics and/or amide C=O [57,58,59,60]. The vibration at 1410 cm⁻¹, coupled with the shoulder at ca. 1350 cm⁻¹, represents carbonates (inorganic portion) and δ(C–H) of aliphatic chains and ν(C–O) of aromatics (organic part) [57,58,61]. Despite the δ(C–H) observation, the absence of distinct peaks of ν(C–H) between 2800 and 3000 cm⁻¹ must be registered, which can be reasonably explained by their coverage due to the extension of the complex band of ν(O–H) described at the beginning of this paragraph. Around 1000 cm⁻¹, a multimodal band was visible and was assigned to ν(Si–O) and δ(Al–OH–Al) of aluminosilicate minerals and C–O stretching of polysaccharides [58,60,61]. At lower wavenumbers in the region of fingerprints, a series of peaks related to Me–OH and/or Me–O modes (where Me = Si, Al, Mg, Ti, Fe) were detected [58,59,60]. After calcination at 800 °C, the soil spectrum was devoid of different signals, as evident in Table 2. The region of ν(O–H) was flattened, indicating a surface depauperation of organic, inorganic, and water-derived hydroxyls. The strong reduction in intensity in the zone around 1400 cm⁻¹ also corroborates the disappearance of organic moieties and a portion of carbonates. The signals that stood out were those at 1000 cm⁻¹, related to minerals (which simultaneously became smoother), and those below 800 cm⁻¹, corresponding to Me–OH and/or Me–O vibrations. In particular, the red aspect of the soil ashes led to the hypothesis of a non-negligible concentration of iron oxide polymorphs that turned into hematite.

3.2. Preliminary Evaluation of Burial Effects on Fabrics

The first attempt to determine the effects of different soil types on buried fabrics was the visual inspection of the just withdrawn samples, as shown in Figure 1 and Figure 2.

It can be seen that the just withdrawn fabrics with the highest degree of attached soil (both in Figure 1 and Figure 2) were those buried in sterilized soil, followed by, in order, the samples in the non-sterilized soil and in the house garden. The main reason, at least when comparing the samples buried in the laboratory within the two different media, seems to be the different packing of soil particles. Indeed, the non-sterilized soil remained always hydrated, leading the water used to avoid dryness to flow uniformly. In contrast, the soil after sterilization was initially dry, so the subsequent moistening (20 mL_(water)/week) presumably created more-compacted aggregates that deposited on the fabric surface. Another observation regarding the sterilized medium is that, after one month, the restoration of microorganism communities from the environment occurred with the formation of mold in some points, as was already pointed out by this group [50,51].

Despite these considerations and the fact that white-stained spots were evidence of microorganism presence, no fabrics showed macroscopic erosions or damage, and indeed, no weight losses were verified. Moreover, the thickness values of the bare polyamide 6.6 and the polyamide 6.6+Chitosan+Dye were, respectively, 0.34 ± 0.02 and 0.41 ± 0.02 mm and did not vary after burial. These results are not particularly surprising considering the well-known resistance of polyamide to breakdown in environmental contexts.

3.3. Color Analysis and Wettability

Color is an illustrative parameter of surface and surface modifications. The colorimetric tests conducted on the finished non-buried fabrics showed that the coating of chitosan alone could be considered slightly dishomogeneous (∆E referred to original polyamide 0.25 ± 0.04), while with carmine subsequent addition, the surface color homogeneity was practically restored (∆E referred to original polyamide 7.63 ± 0.01). Furthermore, in this work, the color was expected to vary due to carmine dye fading or yellow-brown transformation during biodegradation. In this regard,

Table 3. ∆E CIELab values of fabrics buried in various soil types, using polyamide 6.6 and the polyamide 6.6+Chitosan+Dye as references for the related sample series.

Sample	∆E a	Sample	∆E b
P1	4.1 ± 1.0	PT1	1.9 ± 0.3
P2	4.3 ± 1.1	PT2	4.2 ± 0.5
P3	8.1 ± 0.7	PT3	6.0 ± 1.0
P1S	8.7 ± 1.8	PT1S	3.9 ± 0.5
P2S	10 ± 1	PT2S	7.1 ± 0.1
P3S	8.6 ± 0.1	PT3S	5.7 ± 0.5
P1G	26 ± 4	PT1G	6.1 ± 0.2
P2G	21 ± 0.4	PT2G	2.2 ± 0.8
P3G	14 ± 2	PT3G	5.5 ± 0.6

^a Calculated on polyamide 6.6 reference, ^b calculated on polyamide 6.6+Chitosan+Dye reference.

reports the ∆E CIElab color difference between all the buried fabrics and the correlated reference (polyamide 6.6 and polyamide 6.6+Chitosan+Dye). No univocal trends are visible from the obtained values, but a couple of results are worth attention. The first is that the undyed samples overall showed greater ∆E, as the soil adhesion introduced the contribution of colored components after burial, and this also led to an increase in saturation values (ΔC). Dyed samples suffered less from such an effect, being intrinsically colored. The second noteworthy result concerns the high ∆E reached by P1G, P2G, and P3G due to yellowing, demonstrating a substantial impact of the garden soil burial.

Contact angle and drop absorption time were measured to integrate information on changes to the fabric surface. Polyamide 6.6 and polyamide 6.6+Chitosan+Dye resulted in contact angles of 115 ± 3.5° and 117.5 ± 0.2°, respectively, and an absorption time higher than 30 s, thus showing a hydrophobic behavior. For the samples P1, PT1, P1G and PT1G, burial caused a decrease in the contact angle around the limit of 90° and a measurable drop absorption time between 0.5 and 7 s, indicating a certain hydrophilization, probably due to abiotic and biotic deterioration of the surface [62]. Indeed, although ponderal measurement did not reveal relevant degradation, the burial may have turned some exposed chemical groups into polar moieties or introduced adsorbed hydrophilic substances from the surrounding soil. After more than 30 days, the values for the same series (samples in non-sterilized soil and in the house garden) oscillated between the 90° threshold and higher values. These fluctuating outcomes can be attributed to a greater amount of irremovable soil-derived substances/particles covering the surface and, in some measure, altering the analysis [50]. In contrast, the contact angles of fabrics maintained in the sterilized soil never varied remarkably from the untreated references.

3.4. Morphological Studies

SEM analyses of the samples buried for one month did not reveal any particular signs of degradation, except for the detection of adhered soil on the surface, which was significantly abundant in the samples left in the garden (in accordance with the data color results). The situation drastically changed after two and three months, as is visible in Figure 3 and Figure 4, respectively. After two months, the typical smoothness of polyamides was largely maintained. Still, filaments ascribable to bacterial extracellular excretions and/or fungal hyphae were detected (Figure 3A,D) and, consequently, a biofilm seemed to have initially deposited (Figure 3B) [63,64,65,66]. The dyed samples PT2S and PT2G, in particular, showed intense colonization of various microorganisms, resembling, in shape and size, spirochete bacteria [67] (helix in Figure 3C), fungal spores of different origins [68] (depending on the soil involved, Figure 3D,E), and other rod-shaped, chained bacteria [69,70].

After three months, the microbial effects were exacerbated with the increase in biological components. In Figure 4A, for P3, a strong colonization, bringing about an extended biofilm, can be evidenced, together with the presence of irregular black growths (<5 μm-sized). These newly observed formations (zoomed in Figure 4B) have been encountered in all samples buried under laboratory conditions and not detected in the garden-related specimens, which were subjected to a completely different surrounding environment and also to different natural biota competition [71]. It is not straightforward to clearly define their origin, since in the several areas analyzed for each sample, they appear as relieved bulges or detached fiber portions, and even indicate zones with huge erosion signs. Something similar was found by Gashti et al. [37], who subjected polyamide 6.6 to the hydrolysis operated by proteolytic and lipolytic enzymes. The important three-month action of microorganisms is undeniable and also emerges in Figure 4C,D, which shows other events of biofilm formation and fiber damage for P3S and PT3. Figure 4E displays an example of another peculiar phenomenon, in correspondence with bacterial and fungal spore presence, that resembles fiber fusion. Figure 4F–H represents some events that occurred in the samples buried in the house garden, revealing important colonization of fungi and bacteria, although with apparently less severe damage than in other samples.

In general, the above-described degradation phenomena simultaneously caused a further significant loss of smoothness for fibers that often appeared wrinkled or scratched (see Figure 4B,F).

3.5. FTIR-ATR Characterization

Figure 5 displays illustrative results of infrared spectroscopy analyses. The samples of both polyamide 6.6 and polyamide 6.6 after finishing were included as references and showed practically indistinguishable spectra.

The principal absorption peaks at around 3500, 3300 (3070), 2930, 2855, 1630, and 1530 cm⁻¹ are assigned, respectively, to stretching vibrations of surfacial O–H, N–H (plus the related bending overtone), asymmetric and symmetric stretching vibration peaks of C–H of methylene, stretching of C=O (amide I) and the combination of N–H bending and C–N stretching vibrations (amide II) [8,72,73]. The Amide III complex contribution (involving C–N, C–H, and N–H vibrations) [74,75] can be found from 1200 to 1370 cm⁻¹ together with other vibrations linked to methylene and hydrocarbon skeleton (up to 1500 cm⁻¹) [72,73,75]. The signals appearing at 936 and 1140 cm⁻¹ have been previously associated with the crystalline and amorphous structures of polyamide 6.6, respectively [76,77]. With respect to other buried fabrics, the specimens taken from the house garden exhibited the most significant absorption changes. Indeed, the band centered at 1015 cm⁻¹ arose from the first month (magenta rectangle), superimposing the small band at about 1040 cm⁻¹ of the C–C skeleton [78]. This new absorption is concurrent with small peaks at 3700 and 3616 cm⁻¹ (gray arrows in Figure 5) and, together, these signals suggest the presence of soil-derived compounds, especially given the good correspondence with the FTIR-ATR results of the soil analysis (Section 3.1, Table 2). A relative increment of hydroxyl groups’ bands (around 3500 cm⁻¹) was seen as well, after prolonged burial times (ascribable to both organic soil-derived compounds and/or adsorbed moisture), while, contrary to previous findings [50,51], the protein absorptions related to microorganisms were not detected, since they fall in the same range as amide functionalities of polyamide.

3.6. Study of Thermal Properties Through DSC Measurements

In Figure 6, selected DSC results are shown. First, bare polyamide 6.6 and the as-prepared polyamide 6.6+Chitosan+Dye had predictable and comparable peaks around 259 °C and a crystallinity of 44% [79]. Independent of the variables adopted in this research, the analyzed fabrics did not change significantly in the thermogram trend or the melting position (Figure 6A). However, the peak shape (Figure 6B) varied slightly with burial time and treatment type, for instance, in terms of sharpness, such as for curves PT3S and PT3G, which become smoother. Despite the expectation of the recognized mechanism by which chemical agents and microorganisms degrade the amorphous portion and, subsequently, the crystalline part in semicrystalline polymers [80,81], here, the polyamide seemed resistant to substantial structural modification. Indeed, the crystallinity was always around the value of 44%. In a similar attempt, Sudhakar et al. [49] used DSC to characterize a 3-month-biodegraded polyamide 6.6, finding exiguous changes in the melting peak. On that occasion, the transition temperature (Tg) around 50 °C was taken into account to assess the biodegradation. However, in the present work, such differences were not visible since the moisture made a contribution with a broad signal (water evaporation up to 90 °C, Figure 6A). Nevertheless, the increased presence of moisture itself in the buried materials (especially those kept in the house garden) may have been an indication of some deeper phenomenon, such as partial hydrolysis caused by microorganisms or exposure to a humid environment [31,82]. This hypothesis seems consistent with contact angle measurements (hydrophilicity increment within 30 days), but it is not supported by spectroscopic analysis, except for the slight ν(O–H) band modification. Indeed, no clear FTIR-ATR signals of hydrolyzed groups, traducible in C=O and N–H band intensity variations [62,83], were revealed. Similar conclusions regarding higher water absorbency, while maintaining bulk properties, were found in the work of Kanelli et al. [62].

3.7. Evaluation of Antibacterial Activity of Samples Buried for 3 Months

Since the bare polyamide 6.6 fibers do not possess intrinsic antibacterial activity [84], the tests were conducted on UV-sterilized polyamide 6.6+Chitosan+Dye (reference), PT3, PT3S, and PT3G. In previous experiments [30], a bacterial reduction of 95% was registered for polyamide 6.6+Chitosan+Dye after 1 h of contact with S. aureus, while the tests performed in this work showed a lower antibacterial activity (64%) due to a slight heterogeneity of the chitosan coating (see datacolor results) that must be taken into consideration for further data interpretation. PT3 and PT3S were the buried samples that lost the highest degree of antibacterial power (−35–40% of bacterial reduction with respect to the polyamide 6.6+Chitosan+Dye), leading to the hypothesis that, considering the uncertainty given by the small dimension of the samples used and the related heterogeneity, the substantial microbiota colonization seen in SEM analyses in the samples kept in the laboratory had an impact on chitosan efficacy.

3.8. Important Factors Affecting the Biodegradation of Polyamide Fabrics

The combination of the evidence resulting from the materials’ characterizations overall suggests the burial experiments brought about on the polyamide substrate (i) colonization of different microorganisms that verifies the existence of a slow bioactivity, (ii) chemical changes at a minor/localized extent that hinder univocal interpretations on biodegradation phenomena, and (iii) modification of the surface due to adsorbed soil-derived substances.

Figure 7 schematizes the example of a non-finished polyamide 6.6 fabric placed in the garden house. In that case, the atmospheric conditions (UV light, but especially rainfall), combined with mineral-derived compounds and organic substances leaching from soil, seem to be the most impactful variables on the polyamide surface. However, in the case of finished samples, the burial may have slightly affected the antibacterial chitosan functionality. In the literature, chitosan biodegradation is reported to involve enzymes such as chitosanase, lysozyme, and proteases that initiate the process through depolymerization and deacetylation/or cleavage of chitosan’s functional groups, thereby progressively decreasing the molecular weight [85,86,87,88].

4. Summary of Results, Conclusions and Perspectives

The biodegradation of a bare polyamide 6.6 fabric, as well as one finished with chitosan and dyed with the natural carmine colorant, was followed over 3 months, burying fabric pieces in different media and environments: (i) commercial compost-enriched soil as purchased, (ii) the same soil previously sterilized at 105 °C/24 h held in a conditioned laboratory, or (iii) in an open-air house garden in the North West of Italy. This last soil, being completely unknown, was studied for its main features (moisture, pH and nature of chemical groups).

The extent and degradation signs on fabrics were investigated through a preliminary visual inspection and weight assessment, followed by the application of various physical-chemical characterization techniques, including contact angle measurements, color determinations, FTIR-ATR, SEM, and DSC. The characterization outcomes suggested that, within 90 days, all the polyamide 6.6 materials under examination were colonized by different microorganisms, albeit with mild surface deterioration. In particular, the samples buried in the house garden showed a higher interaction with the complex composition of this real soil matrix which contributed to the adsorption/deposition of both organic and inorganic substances that, in turn, brought about (i) non-finished polyamide 6.6 yellowing; (ii) greater FTIR-ATR spectra modifications; and (iii) a higher propensity to water adsorption (from DSC curves). Except for the color, no other differences emerged when comparing the bare or finished fabrics. Similarly, due to the overall exiguous entity of fabric disruption, no particular behavioral discrepancies were highlighted between the samples buried in sterilized and non-sterilized soil at the lab scale.

As main conclusions, despite the evidenced interaction between fabrics and soil (see Section 3.8), a rapid and substantial biota stimulation of the bio-based substances chitosan and carmine to degrade the polyamide 6.6 may be ruled out, in contrast to what was hypothesized when the fabric substrate was cotton. However, observing the finished samples, particularly PT3 and PT3S (lab-scale), a slight reduction in antibacterial action against S. aureus operated by chitosan was revealed. It cannot, therefore, be excluded that microbiota have started a degradation process at the expense of this macromolecule before damaging the polyamide substrate. Thus, on the one hand, the results obtained in this work insightfully delineate the crucial role of burial environment (soil type, weather conditions, etc.) for synthetic fabrics, together with the limited influence of biodegradable finishing. On the other hand, the design of biodegradation experiments over longer periods and employing differently finished synthetic fabrics will be of paramount interest to better quantify the major effects.