Use of Hydroxyapatite Nanoparticles to Reduce Cd Contamination in Agricultural Soils: Effects on Growth and Development of Chenopodium quinoa Willd

Abstract

Soil contamination and degradation have prompted extensive research into remediation techniques. A promising approach involves the use of nanoparticles, which can mitigate heavy metal contamination, such as cadmium (Cd), without adversely affecting crop development. This study evaluated the effects of hydroxyapatite nanoparticles (HANPs) on the growth of Chenopodium quinoa Willd. in soils contaminated with varying Cd contents (0, 5, 10, 25, and 50 mg/kg). The results indicated that HANPs enhanced early shoot development, particularly in soils with Cd concentrations ≥10 mg/kg, while significantly reducing Cd accumulation in plant tissues. HANPs also decreased Cd mobility in soil, retaining it in fractions less available for plant uptake. Germination rates differed between pot experiments and phytotoxicity assays, although the first demonstrated greater Cd immobilization in HANP-treated soils, resulting in reduced Cd translocation to quinoa shoots. These findings highlight the potential of HANPs as an effective tool for remediating Cd-contaminated soils, thereby improving soil pollution, crop safety, and quality for human consumption.

1. Introduction

Soil is the main environmental compartment of terrestrial ecosystems, which provide fundamental functions for human and environmental well-being. Soil is the primary medium for food production, supporting approximately 95% of the global food supply [1]. Beyond its agricultural function, soil acts as a natural filter, retaining and detoxifying pollutants, and plays a key role in the biogeochemical cycles of essential nutrients such as nitrogen, phosphorus, and carbon. These processes are facilitated by the diverse microbial and faunal communities inhabiting the soil, which enhance organic matter decomposition and nutrient recycling [2]. However, human activities, such as industrialization, urbanization, and intensive agricultural practices, have significantly altered soil properties and functions. This has led to widespread degradation and contamination, threatening soil productivity and ecosystem services [3].

One of the most persistent causes of soil pollution is contamination by heavy metals. Although heavy metals occur naturally in soils in trace concentrations, human activities have greatly aggravated their presence. Cadmium (Cd), in particular, is a highly toxic and mobile heavy metal with no known biological function. Its anthropogenic sources include mining, smelting, industrial discharges, and the widespread use of phosphate fertilizers containing Cd as an impurity [4,5,6]. Cadmium contamination in agricultural soils is of particular concern due to its mobility, which facilitates its uptake by plants and its entry into the food chain [7]. Once absorbed, Cd impairs plant physiological functions, causing chlorosis, reduced photosynthetic activity, and stunted growth [8]. In humans, chronic exposure to Cd is associated with kidney damage, osteoporosis, carcinogenesis and the infamous “itai-itai” disease, which emerged in Japan due to the long-term consumption of Cd-contaminated rice [9,10].

The global extent of heavy metal contamination is alarming, with over than 22 million hectares of agricultural soils affected, a figure likely underestimated given the intensification of industrial and agricultural activities worldwide [11]. Addressing this contamination is vital for ensuring food safety and environmental health. Conventional remediation methods, such as soil excavation, washing, and chemical stabilization, are often costly and environmentally damaging. For instance, soil washing generates large volumes of contaminated leachates, while excavation and dumping permanently deplete soil resources and disrupt ecosystems [12]. These limitations have prompted the search for innovative, sustainable, and cost-effective approaches to soil remediation, including nanotechnology-based interventions.

Nanotechnology offers a promising alternative due to the unique physicochemical properties of nanoparticles, such as high surface area, increased reactivity, and the ability to interact with contaminants at molecular levels [13]. Among several nanoparticles, hydroxyapatite nanoparticles (HANPs) have garnered significant consideration for their effectiveness in immobilizing heavy metals [14,15]. Hydroxyapatite (HA), a calcium phosphate mineral, occurs naturally in rocks and soils and has been recognized for its capacity to immobilize heavy metals through mechanisms such as adsorption, ion exchange, and co-precipitation [16]. The HA has improved reactivity and surface area in its nanoparticulate form, making HANPs particularly effective for environmental applications. In addition, HANPs act as slow-release phosphorus fertilizers, improving soil fertility and favoring plant growth, which increases their applicability in agriculture [17]. Studies have shown that HANPs significantly reduce the mobility and bioavailability of Cd, mitigating its toxic effects on plants. For instance, the application of HANPs to Cd-contaminated soils reduced Cd concentrations in rice tissues by 50% while enhancing root and shoot biomass [18]. In addition, the degradation nutrients of HANPs, i.e., calcium and phosphate ions, make them an environmentally friendly alternative to conventional amendments [19]. Despite these advantages, integrating HANPs with biological remediation approaches, such as phytoremediation, remains practically unexplored.

Phytoremediation is a sustainable approach to soil remediation that has gained attention for its environmental compatibility and cost effectiveness. However, its effectiveness is often limited by the toxicity of contaminants, which can impair plant growth and reduce biomass [20,21]. Combining phytoremediation with HANPs may offer a synergistic solution, where nanoparticles reduce the bioavailability of heavy metals, allowing plants to thrive and enhance soil recovery [19,22,23].

Among the potential phytoremediation crops, quinoa (Chenopodium quinoa Willd.) stands out for its resilience, adaptability, and nutritional value. Quinoa, a native species from the Andean region, is able to grow in extreme environmental conditions, including saline, nutrient-poor, and contaminated soils [24,25,26]. Its genetic diversity and robust root system make it an excellent candidate for phytoremediation in heavy metal-contaminated soils. Previous research has shown that quinoa has a moderate capacity to tolerate Cd contamination, with some genotypes capable of restricting Cd translocation to aerial parts, thereby reducing the risk of Cd entering the food chain [27,28]. In addition, the low nutrient requirements and high biomass production of quinoa increase its suitability for remediation strategies in degraded soils.

This study hypothesizes that HANPs can effectively immobilize Cd in soils with high concentrations of this metal, promoting the adaptation and growth of quinoa in Cd-contaminated soils. The main objective of this research is to determine whether the combined use of HANPs and Chenopodium quinoa Willd. is effective for the remediation of Cd-contaminated soils, thereby improving the development of this species in such environmental conditions. Additionally, the study evaluates the effects of both Cd and HANPs on the germination and early growth of quinoa. The specific objectives are (i) to evaluate the capacity of HANPs to reduce the mobility and bioavailability of Cd in soil; (ii) to investigate the sorption and desorption processes of Cd in soil, as well as its distribution among different geochemical fractions with or without the presence of HANPs; (iii) to assess the toxicity of Cd to quinoa through phytotoxicity assays in soil; and (iv) to study the effects of Cd and HANPs on the germination and early growth of quinoa.

2. Materials and Methods

2.1. Soil Sampling

An agricultural soil developed over granite and without fertilizer applications for more than 10 years in the district of Parada (Nigrán, Pontevedra, Galicia, Spain) (42°8′9″ N, 8°46′34″ W) was selected (Supplementary Materials). A total of six sub-samples were taken from different points within the selected plot to ensure representative sampling. The soil sub-samples were collected from a depth of 0–30 cm (topsoil) using an Eijkelkamp sampler and stored in polyethylene bags. These sub-samples were pooled, air-dried, sieved through a 2 mm mesh, and homogenized into a single composite soil sample.

2.2. Soil Characterization

The soil samples were analyzed for pH, organic matter, N and P contents, particle size distribution, effective cation exchange capacity (ECEC), oxide contents, and pseudototal metal content according to the following methodologies. The pH was measured in two suspensions: one prepared with deionized water and another with 0.1 M KCl, both in a 1:2.5 (w/v) ratio [29]. Organic matter content was assessed by loss on ignition, heating 2 g of soil at 550 °C for 5 h in a muffle furnace according to ISO 10694:1995 [30]. Nitrogen contents were determined in an elemental analyzer (Thermo Flash EA 1112, Waltham, MA, USA), by complete combustion of the sample inside a reactor with a high-temperature (900 °C) oxidizing catalyst. The phosphorus content was determined using the Olsen method, which estimates the available phosphorus in soils through extraction with sodium bicarbonate (0.5 M, pH 8.5) [31]. Particle size distribution was analyzed by wet sieving for fractions >50 µm (sand) and using the Robinson pipette method for silt and clay fractions, based on Stokes’ law. The soil was pretreated with hydrogen peroxide to remove organic matter and dispersed with sodium hexametaphosphate [29]. The ECEC was determined by the sequential extraction of exchangeable cations (Na⁺, K⁺, Mg²⁺, and Ca²⁺) with 1 M NH₄Cl, quantified by ICP-OES (Perkin Elmer Optima 4300 DV, Perkin Elemer, Waltham, MA, USA), and exchangeable Al³⁺ was extracted with 1 M KCl and titrated with 0.01 M NaOH [29]. Oxides of Fe, Al, and Mn were extracted using the dithionite–citrate method [29] and quantified by ICP-OES (Perkin Elmer Optima 4300 DV). The pseudototal metal content was determined by the acid digestion of 0.2 g of soil using concentrated HNO₃ and HCl in a microwave oven according to EPA Method 3051A [32]. After filtration, metal concentrations were measured by ICP-OES (Perkin Elmer Optima 4300 DV). All results were standardized to the dry soil weight.

2.3. Plant Selection

Quinoa (Chenopodium quinoa Willd.) seeds were selected for toxicity assays on soils with and without nanoparticles due to their well-known resilience to harsh environmental conditions, including nutrient-poor and contaminated soils. The seeds used in this study were supplied by Semillas Vivas S.L. (Extremadura, Spain). To ensure high germination potential, only certified seeds were selected based on preliminary germination tests. Quinoa was chosen for its remarkable adaptability to a wide range of climatic conditions, tolerating temperatures from −4 °C to 38 °C and relative humidities between 40% and 90%.

2.4. Nanoparticles

The HANPs with a needle-like morphology and a nominal particle size of 20 nm (purity: 99%) were used in the experiments. The HANPs were supplied by MKnano (M.K. Impex Corp., Toronto, ON, Canada; product reference: MKN-HXAP-020P, CAS number: 12167-74-7).

According to the Certificate of Analysis of MKnano, the HANPs exhibited additional physicochemical properties worthy of consideration, such as: a white color; sulfates of 0.025% (max: 0.048%); chlorides of 0.02% (max: 0.05%); other heavy metals of 7 ppm (max: 10 ppm); a loss on drying of 0.75% (max: 1.0%); a density of 3.14 g/cm³; a pH stable under neutral and slightly acidic conditions; water insolubility; and a melting point ≈1100 °C. The HANPs were selected for their high chemical stability and reactivity under environmental conditions.

2.5. Soil Treatments with Cd and/or Nanoparticles

Ninety grams of soil were placed in glass jars, and HANPs were added at 1% (w/w). The decision to use 1% (w/w) HANPs in the assays was based on both previous studies and preliminary experiments conducted by our research team. In earlier studies, doses with 2% and 3% of HANPs were tested in different experimental settings, demonstrating that higher concentrations may offer increased metal immobilization but could also pose potential risks to plant development due to their high reactivity [13,33]. For untreated soils, this step was made without nanoparticles. An aqueous Cd(NO₃)₂ solution was added to achieve final Cd concentrations of 0, 5, 10, 25, and 50 mg kg⁻¹ (

Table 1. Treatments applied for the different soil analyses with Cd and/or HANPs.

Cd Concentration Added	Nanoparticles
0 mg kg−1	Without nanoparticles
0 mg kg−1	1% HANPs
5 mg kg−1	Without nanoparticles
5 mg kg−1	1% HANPs
10 mg kg−1	Without nanoparticles
10 mg kg−1	1% HANPs
25 mg kg−1	Without nanoparticles
25 mg kg−1	1% HANPs
50 mg kg−1	Without nanoparticles
50 mg kg−1	1% HANPs

). The selected Cd concentrations were determined based on the established Generic Reference Levels (NGRs) for Galicia and other international guidelines [34]. These levels represent increasing degrees of contamination, from uncontaminated soils (0 mg/kg) to highly contaminated scenarios (50 mg/kg). The inclusion of these concentrations simulates realistic contamination scenarios that may occur due to agricultural or industrial activities. The samples were adjusted to field capacity and incubated at room temperature on a rotary shaker for 96 h. No fertilizers were applied during the experiments. The only amendments used were HANPs and Cd(NO₃)₂, added before planting quinoa seeds. Treated and untreated soils were used for phytotoxicity assays with quinoa.

2.6. Phytotoxicity Assays

Phytotoxicity assays were conducted using quinoa (Chenopodium quinoa Willd.) to evaluate the effects of cadmium (Cd) and HANPs on seed germination and early plant growth. The procedure followed the standardized protocol provided by Phytotoxkit (MicroBioTests Inc., Montreal, QC, Canada) [35], which measures the inhibition of germination and the growth of roots and shoots under the different soil treatments (Table 1).

The phytotoxicity assays were conducted in transparent plates to evaluate seed germination and measure seedling length (shoot and root) (Supplementary Materials). They were carried out following the procedure below: (i) Soil samples (untreated and treated with Cd and/or nanoparticles) were placed at the bottom of transparent plates (90 cm³) at field capacity. The soil was then leveled in each plate, compacted, and covered with black filter paper. (ii) Ten quinoa seeds were evenly placed on the black filter paper on each plate. The plates were sealed, closed, and positioned vertically. The plates were incubated for 3 days in darkness at 25 °C. After incubation, seeds with radicles ≥ 1 mm were considered germinated and recorded. (iii) Finally, the plates were photographed with a digital camera, and the images were analyzed using the ImageJ software (version 1.54f, National Institutes of Health) to obtain root and shoot length data. These data were used to determine and compare germination inhibition (%) and root growth inhibition (%) across all soil samples (treated and untreated with Cd and/or nanoparticles).

To assess the phytotoxicity of Cd and the mitigating effects of HANPs, the germination index (G_Index) was calculated following [36]:

$${\mathrm{G}}_{\mathrm{I}\mathrm{n}\mathrm{d}\mathrm{e}\mathrm{x}}\left(\mathrm{\%}\right)={\displaystyle \frac{{\mathrm{G}}_{\mathrm{s}}·{\mathrm{L}}_{\mathrm{s}}}{{\mathrm{G}}_{\mathrm{c}}·{\mathrm{L}}_{\mathrm{c}}}}·100$$

where G_s and L_s, represent the seed germination percentage and average root length in Cd-treated soils (with or without HANPs), and G_c and L_c are the corresponding values in control soils. G_Index values were categorized as follows [36]:

• G_Index: values between 90 and 110% are classified as non-toxic to the species under Cd exposure.
• G_Index: values < 90% are classified as having an inhibitory effect.
• G_Index: values > 110% are classified as having a stimulatory effect.

The aerial part influence index (Ap_Index) was calculated based on the G_Index equation:

$${\mathrm{A}\mathrm{p}}_{\mathrm{I}\mathrm{n}\mathrm{d}\mathrm{e}\mathrm{x}}\left(\mathrm{\%}\right)={\displaystyle \frac{{\mathrm{G}}_{\mathrm{s}}·{\mathrm{A}\mathrm{p}}_{\mathrm{s}}}{{\mathrm{G}}_{\mathrm{c}}·{\mathrm{A}\mathrm{p}}_{\mathrm{c}}}}·100$$

where G_s and Ap_as are the seed germination percentage (%) and the average shoot length (mm), respectively, in soils treated with Cd and/or nanoparticles. Similarly, G_c and Ap_c, are the corresponding values for plants grown in untreated soils. Ap_Index was classified as follows:

• Ap_Index: values between 90 and 110% are classified as non-toxic to the species for shoot growth under Cd exposure.
• Ap_Index: values < 90% are classified as having an inhibitory effect on shoot growth.
• Ap_Index: values > 110% are classified as having a stimulatory effect on shoot growth.

2.7. Pot Experiments

Pot experiments were conducted to study the medium-term effects of Cd and HANPs on the growth of quinoa. The treatments applied were identical to those used in the phytotoxicity assays (Table 1).

Soil samples (1.5 kg per pot) were homogenized and mixed with HANPs at a concentration of 1% (w/w) for treatments involving nanoparticles. Aqueous solutions of Cd(NO₃)₂ were added to achieve final Cd concentrations of 0, 5, 10, 25, and 50 mg kg⁻¹ in the soil. The pots were incubated for 96 h at room temperature before planting. Ten quinoa seeds were sown in each pot. The pots were maintained in a greenhouse at the Campus of Ourense (Universidade de Vigo, Galicia, Spain) under controlled environmental conditions. They were irrigated regularly with deionized water to maintain field capacity for 21 days. After this period, the seedlings were thinned to two plants per pot, selecting those with the greatest development potential to minimize competition. Irrigation with deionized water continued for 69 more days, allowing the plants to grow for a total of 90 days.

At the end of the growth period, the plants were harvested, and their shoots and roots were separated. Plant height and fresh biomass were recorded. Roots and shoots were washed with tap water, followed by distilled water, and then subjected to ultrasonic cleaning for 30 min to remove residual soil particles. Shoot samples were further divided into leaves and stems. Leaves were photographed with a digital camera, and their images were analyzed using ImageJ software (version 1.54f, National Institutes of Health) to measure leaf area.

Both stems and leaves were oven-dried at 40 °C, and their dry biomass was determined. Subsequently, 0.2 g of leaf and stem samples were digested with 98% HNO₃ in a microwave digester to determine Cd content in plant tissues. The plant material was digested in 7.5 mL of concentrated HNO₃ for 12 h at room temperature. The samples were then heated in a digestion block for 2 h at 105 °C. After cooling, the digests were filtered and diluted to 10 mL with distilled water. The Cd concentrations were measured by ICP-OES (Perkin Elmer Optima 4300 DV).

2.8. Sequential Chemical Extraction

Sequential chemical extraction was performed to determine the distribution of Cd among different geochemical fractions of the soil. Soil samples were collected from the pots after the 90-day growth period. The extraction method was based on the protocol described by Almås et al. [37], with modifications made by Salbu et al. [38], and originally developed by Tessier et al. [39]. The process involved the use of the following reagents in a stepwise manner:

• F1 (water-soluble and exchangeable): 0.001 M KNO₃ for 24 h at room temperature.
• F2 (exchangeable): 1 M NH₄OAc at soil pH for 2 h at room temperature.
• F3 (specifically adsorbed): 1 M NH₄OAc at pH 5 for 2 h at room temperature.
• F4 (oxide-bound): 0.04 M NH₂OH·HCl in 25% HAc for 6 h at 80 °C.
• F5 (organic matter-bound): 30% H₂O₂ at pH 2 for 5.5 h at 80 °C, followed by 3.2 M NH₄OAc in 20% HNO₃ for 0.5 h at room temperature.
• F6 (irreversibly adsorbed): 7 M HNO₃ for 6 h at 80 °C.
• F7 (residual): the remaining solid fraction after previous extractions.

After each extraction step, the samples were centrifuged, and the supernatant was filtered and stored for analysis. The solid fraction was washed with 10 mL of distilled water, centrifuged again, and the wash solution was combined with the previous extract for further analysis. Cd concentrations in each fraction were measured using ICP-OES (Perkin Elmer Optima 4300 DV). This procedure allowed for the quantification of Cd in the following soil fractions:

• F1 and F2: physical sorption (reversible).
• F3: electrostatic adsorption (exchangeable).
• F4, F5, F6: chemisorption (irreversible).
• F7: residual Cd.

The recovery percentage was calculated as the ratio of the sum of the six fractions (F1–F6) to the total amount of Cd initially added to each pot.

2.9. Statistical Analysis

All experiments were performed in triplicate, including soil characterization, phytotoxicity assays, and pot experiments. Statistical analyses were conducted using IBM-SPSS v. 28.0 software for Windows. Descriptive statistics were calculated for all measurements, including mean values and standard deviations. Differences between treatments (with and without Cd and/or HANPs) were evaluated using analysis of variance (ANOVA) followed by Duncan’s multiple range tests for pairwise comparisons. Fisher’s least significant difference (LSD) test was applied at a 5% significance level to compare mean values. Data normality was verified using the Kolmogorov–Smirnov test, and homogeneity of variances was assessed using Levene’s test to confirm data homoscedasticity.

3. Results and Discussion

3.1. Soil Analysis

The soil was exhaustively characterized, and its main characteristics and pseudototal metal contents are summarized in

Table 2. Soil characteristic analysis.

Characteristic	Unit	Value	Element	Units	Concentration
pH(H2O)	-	5.75 ± 0.10	Al	g kg−1	57.71 ± 8.69
pH(KCl)	-	4.85 ± 0.10	As	mg kg−1	14.58 ± 1.18
Organic Matter	%	2.01 ± 0.35	Ca	g kg−1	1.76 ± 0.05
N	%	0.13 ± 0.01
P	mg kg−1	28.60 ± 1.04
Texture	-	sandy-loam	Cd	mg kg−1	nd
			Co	mg kg−1	17.92 ± 0.38
Ca2+	cmol(+)kg−1	1.69 ± 0.34	Cr	mg kg−1	68.75 ± 2.18
K+		18.11 ± 4.84	Cu	mg kg−1	77.92 ± 1.84
Mg2+		0.50 ± 0.10	Fe	g kg−1	48.81 ± 3.08
Na+		1.50 ± 0.69	K	g kg−1	10.94 ± 1.07
Al3+		31.11 ± 1.97	Mg	g kg−1	10.88 ± 0.77
ECEC		52.92 ± 7.94	Mn	g kg−1	0.43 ± 0.02
			Na	mg kg−1	135.83 ± 3.54
Fe oxides	g kg−1	35.08 ± 4.17	Ni	mg kg−1	31.92 ± 1.26
Al oxides		5.83 ± 0.70	Pb	mg kg−1	9.50 ± 1.32
Mn oxides		0.15 ± 0.02	Zn	mg kg−1	107.67 ± 2.45

nd: not detected; ECEC: effective cation exchange capacity.

.

The sandy-loam texture of the soil indicates a high sand content with relatively low amounts of silt and clay, which limits its capacity to retain heavy metals. Clay particles are key for metal retention due to their high surface area, cation exchange capacity, and the ability to form stable complexes with metal ions [40]. However, the low clay content in sandy-loam soils diminishes these effects, potentially increasing metal mobility and environmental risk, making it suitable for evaluating the effects of HANPs on Cd immobilization without interference from clay–mineral interactions. The soil showed a moderately acidic pH (5.75 in water and 4.85 in KCl), which is an essential factor in metal solubility and bioavailability. Acidic soils increase the solubility of heavy metals due to the abundance of H⁺, which compete for sorption sites on soil particles [40]. The lower pH in KCl shows the exchangeable acidity, further highlighting the presence of H⁺ and Al³⁺ in the exchange complex. This acidic environment suggests higher Cd mobility and bioavailability as a result of the lack of HANP treatments. Organic matter plays an essential role in the biogeochemical cycling of heavy metals through mechanisms such as complexation, adsorption, and chelation [41]. However, the organic matter content in this soil was low (2%), limiting its capacity to retain heavy metals and interact with HANPs. The total N content was 0.13%, which is considered low for agricultural soils, showing limited N availability for plant growth, while the available phosphorus content was 28.60 mg kg⁻¹, which is within the medium range for agricultural soils [42]. The ECEC of the soil was 52.92 cmol₍₊₎kg⁻¹, mainly due to the high concentration of exchangeable Al³⁺. This is characteristic of acid soils, where Al³⁺ dominates the exchange complex over basic cations such as Ca²⁺, Mg²⁺, and Na⁺, which are present in much lower concentrations. The predominance of Al³⁺ contributes to the high ECEC but also influences the soil buffering capacity, promoting Cd mobility by reducing the availability of exchange sites for Cd retention [43]. Iron and manganese oxides are key components that influence heavy metal retention. These oxides form stable complexes with metals, reducing their bioavailability [44]. The soil contained relatively high levels of Fe oxides (35.08 g kg⁻¹), while Al and Mn oxides were less abundant (5.83 g kg⁻¹ and 0.15 g kg⁻¹, respectively). The high contents of Fe oxides suggest their significant role in heavy metal retention, particularly through chemisorption mechanisms [43]. The pseudototal metal analysis showed that Cd was below the detection limit (<0.01 mg kg⁻¹), confirming the absence of Cd in the soil. This ensures that Cd detected in the soil originates from the experimental treatments. In addition to Cd, pseudototal concentrations of other metals were analyzed to provide a detailed characterization of the agricultural soil (Table 2). These metals can have potential competitive interactions with HANPs during the immobilization of Cd. In particular, Cu and Zn may have a strong affinity for adsorption onto mineral surfaces, including H [45]. High concentrations of these metals may compete with Cd for sorption sites on the HANPs, potentially reducing their effectiveness in immobilizing Cd. In this manner, Cr and Ni, although less abundant in agricultural soils, also show potential mobility under acidic soil conditions and their ability to form stable complexes with soil organic matter and mineral oxides, which may influence the dynamics of metal immobilization [46]. In any case, only the Cu levels exceeded the reference values for non-industrial soils in Galicia [34]. This elevated Cu concentration may be attributed to agricultural practices, such as the use of Cu-based fungicides, which are common in the region. Other metals, such as Zn, Cr, and Ni, were within typical background levels for soils in similar environments. Thus, all these characteristics make the soil suitable for studying the effects of HANPs on Cd immobilization, providing a controlled environment to evaluate the interactions between Cd, treatment with HANPs, soil components, and quinoa.

3.2. Assessment of Cd Distribution in Different Soil Fractions

The Cd content in each fraction obtained by sequential chemical extraction is shown in

Table 3. Fractions F1 to F6 of sequential Cd extraction (mg kg⁻¹).

Soil + HANPs	Cd	F1	F2	F3	F1 + F2 + F3
-	0	nd	nd	nd	nd
1%	0	nd	nd	nd	nd
-	5	nd	1.04 ± 0.3 e	0.79 ± 0.34 d	1.83 ± 0.64 d (36.60%)
1%	5	nd	1.41 ± 0.26 d	0.50 ± 0.24 d	1.91 ± 0.50 d (38.20%)
-	10	0.11 ± 0.03 c	3.49 ± 1.28 c	2.45 ± 0.85 c	6.05 ± 2.16 c (60.50%)
1%	10	0.04 ± 0.02 d	3.73 ± 0.02 c	2.13 ± 0.49 c	3.77 ± 0.53 c (37.70%)
-	25	0.28 ± 0.07 b	8.72 ± 3.2 b	6.11 ± 2.13 b	15.11 ± 5.40 b (60.44%)
1%	25	0.11 ± 0.04 c	9.31 ± 1.23 ab	5.33 ± 0.39 b	14.75 ± 1.66 b (59.00%)
-	50	0.73 ± 0.54 a	13.56 ± 3.84 a	11.3 ± 1.65 a	25.59 ± 6.03 a (51.18%)
1%	50	0.61 ± 0.11 a	14.77 ± 2.8 a	11.03 ± 0.84 a	26.41 ± 3.75 a (51.82%)
Soil + HANPs	Cd	F4	F5	F6	% Recovery
-	0	nd	nd	nd	-
1%	0	nd	nd	nd	-
-	5	1.12 ± 0.18 d	0.13 ± 0.03 d	0.66 ± 0.0.8 d	74.80
1%	5	0.82 ± 0.25 d	nd	1.14 ± 0.21 d	77.30
-	10	1.87 ± 0.63 c	0.79 ± 0.32 c	0.57 ± 0.13 e	92.80
1%	10	1.36 ± 0.16 c	nd	2.44 ± 0.62 c	75.65
-	25	4.68 ± 1.57 a	1.97 ± 0.81 b	1.43 ± 0.43 d	92.76
1%	25	3.40 ± 0.26 b	nd	3.42 ± 0.76 c	86.28
-	50	7.80 ± 1.33 a	4.38 ± 0.78 a	5.86 ± 1.05 b	87.25
1%	50	5.58 ± 0.55 a	0.08 ± 0.03 d	8.96 ± 0.79 a	82.05

nd: not detected. F1 + F2 + F3 in parentheses: percentage (%) of total Cd associated with these fractions over total Cd. Different letters next to the values indicate significant differences among treatments for each fraction at a significance level of p < 0.05.

. The results indicate that Cd recovery after sequential extraction exceeded 75% for all treatments with or without HANPs, indicating the effectiveness of the method in assessing the distribution of Cd among geochemical fractions.

The distribution of Cd among the geochemical fractions depends on soil characteristics (Table 1). The sandy-loam texture, characterized by a low clay content, reduces the soil capacity to retain Cd. In addition, the low organic matter content limits the formation of organic–metallic complexes, while the acidic pH increases Cd solubility by enhancing competition between H⁺ and heavy metals for sorption sites on soil particles. These characteristics explain the relatively high proportion of Cd associated with bioavailable fractions (F1–F3), particularly in untreated soils. The high ECEC, predominantly influenced by exchangeable Al³⁺, contributes to the mobility of Cd. The Al³⁺ occupies exchange sites that might otherwise bind Cd, facilitating the presence of the metal in soil solution. This is relevant to the inhibition of quinoa germination and growth observed during phytotoxicity tests and pot experiments. The efficacy of HANPs in mitigating Cd toxicity seems to depend on their ability to immobilize the metal in less bioavailable forms, as evidenced by the increased F6 content and reduced bioavailable fractions observed in the sequential extraction results.

The non-presence of Cd in the fractions from the control soils, both untreated and treated with HANPs, confirms the lack of natural Cd in the soil samples, as indicated in the pseudototal metal analysis (Table 1). This confirms that Cd comes from the experimental treatments. In the Cd-treated soils, the proportion associated with the most bioavailable fractions (F1, F2, and F3) ranged from 36.6% (Soil + Cd 5 mg kg⁻¹) to 60.4% (Soil + Cd 25 mg kg⁻¹). These fractions, representing exchangeable and weakly bound Cd, are responsible for the risks of quinoa exposure to Cd and its potential mobility in soil.

There was no significant difference in the distribution of Cd for the sum of F1–F3 between soils treated with HANPs and untreated soils. This suggests that HANPs did not significantly reduce the proportion of Cd in bioavailable forms under the conditions tested. The persistence of bioavailable Cd in HANP-treated soils is consistent with the results of phytotoxicity tests and pot experiments, where reductions in germination rates and root growth were observed at Cd concentrations ≥ 10 mg kg⁻¹. This indicates the limited effectiveness of HANPs in immobilizing Cd in these highly available fractions at the highest Cd contamination levels.

In the less bioavailable fractions (F4–F6), the influence of HANPs is more significant. Specifically, the Cd content complexed with organic matter (F5) decreased significantly in soils treated with HANPs compared to untreated soils. This indicates that HANPs may modify the Cd distribution from organic matter-bound forms to more stable and less bioavailable forms. In this line, an increase in the formation of stable metal–phosphate complexes in soils amended with HANPs was observed in a previous study [47]. Additionally, a significant increase in irreversibly adsorbed Cd (F6) was also observed in soils treated with HANPs, particularly at higher Cd concentrations (10, 25 and 50 mg kg⁻¹). This suggests that HANPs promote Cd immobilization through mechanisms such as chemisorption or precipitation, leading to the formation of highly stable Cd–phosphate complexes. In fact, a previous study demonstrated that hydroxyapatite application reduced bioavailable Cd concentrations by transforming exchangeable and soluble fractions into more stable forms [22]. Similarly, it was found that combining HA with humic acids decreased Cd availability in maize while enhancing crop performance, highlighting the potential synergistic benefits of using multiple amendments [48]. These findings support that HANPs alone may effectively transform Cd into less bioavailable forms (F6), thereby reducing its translocation to plant tissues, as observed in the pot experiments with quinoa. However, the limited impact of HANPs on highly bioavailable Cd fractions (F1–F3) at elevated contamination levels highlights the need for complementary remediation strategies. Combining HANPs with organic amendments may further enhance their effectiveness by stabilizing Cd in soil and improving quinoa growth.

3.3. Phytotoxicity Assays

Phytotoxicity assays with quinoa seeds were conducted on untreated soils and soils treated with Cd and/or HANPs to assess Cd toxicity and the effects of HANPs on seed germination and early plant growth. Germination percentages (G%), shoot and root lengths, as well as the calculated G_Index and Ap_Index values, are presented in

Table 4. Germination percentage G (%); the lengths of aerial parts and roots (cm); the index of influence on the aerial part (Ap_Index); and the germination index (G_Index) of quinoa.

Soil + HANPs	Cd (mg kg−1)	G (%)	Shoot Length (cm)	ApIndex (%)	Root Length (cm)	GIndex (%)
-	0	83 ± 15 a	4.28 ± 0.12 a	100 ± 17 a	2.85 ± 0.75 a	100 ± 31 a
1%	0	80 ± 17 a	5.06 ± 0.93 a	112 ± 22 a	2.87 ± 0.80 a	93 ± 13 a
-	5	80 ± 10 a	4.83 ± 1.26 ab	106 ± 21 a	3.35 ± 0.59 a	111 ± 11 a
1%	5	77 ± 12 a	4.45 ± 0.99 ab	97 ± 32 ab	2.58 ± 0.53 ab	85 ± 30 a
-	10	73 ± 15 a	4.11 ± 0.18 b	84 ± 14 b	2.08 ± 0.36 b	64 ± 16 b
1%	10	83 ± 6 a	5.03 ± 0.78 a	119 ± 20 a	2.11 ± 0.79 ab	74 ± 25 ab
-	25	77 ± 6 a	4.15 ± 0.08 b	87 ± 7 b	2.14 ± 0.95 ab	70 ± 34 ab
1%	25	87 ± 15 a	4.84 ± 0.74 a	119 ± 25 a	2.55 ± 0.21 a	94 ± 22 a
-	50	77 ± 15 a	3.28 ± 0.22 c	70 ± 10 b	2.06 ± 0.43 b	67 ± 32 b
1%	50	90 ± 10 a	3.16 ± 0.85 bc	78 ± 16 b	2.08 ± 0.77 ab	77 ± 25 ab

Different letters next to the values in each column indicate significant differences among treatments for each measured parameter at a significance level of p < 0.05.

To assess the potential toxicity of Cd and the effect of HANPs on germination, root and shoot growth were measured for each seedling, and the G_Index and Ap_Index values were calculated.

The observed germination percentages for untreated soils without Cd were in accordance with the typical values reported for quinoa under non-stress conditions [39]. A reduction in germination rates was not evident as Cd concentrations increased, highlighting the lack of impact of Cd on seed germination. The results for shoot length and Ap_Index showed no significant differences in quinoa shoot growth between soils without Cd (0 mg kg⁻¹) and those treated with 5 mg kg⁻¹ of Cd, regardless of the presence of HANPs. However, there was a notable reduction in shoot development and Ap_Index values in seedlings grown in soils with Cd concentrations of 10, 25, and 50 mg kg⁻¹ without HANPs. For soils treated with HANPs, the results showed that the HANPs mitigated the toxic effects of Cd at concentrations of 10 and 25 mg kg⁻¹, as shown by the higher shoot lengths and Ap_Index values compared to untreated soils. With the application of 50 mg kg⁻¹ of Cd, the HANPs did not have a significant effect on shoot growth or Ap_Index values, indicating a limitation in their effectiveness at the highest Cd concentration applied. The improvement in shoot growth and P_Index values in HANP-treated soils can be attributed to the ability of HANPs to reduce Cd toxicity. HANPs likely immobilize Cd through mechanisms such as adsorption, ion exchange, or precipitation, which decreases Cd availability in the soil and water solution [49,50]. Additionally, HANPs may provide calcium and phosphate ions to the plants, which could support cellular functions and partially mitigate the stress induced by Cd [51]. However, the lack of significant improvement at 50 mg kg⁻¹ Cd suggests that the mitigation capacity of 1% HANPs is limited under high metal stress, which highlights the importance of optimizing nanoparticle dosage based on the level of contamination.

The root length and G_Index results indicated that adding HANPs to soils without Cd generally did not promote early root growth compared to the control. The G_Index values below 90% indicate an inhibitory effect on early root growth when HANPs were added to soils, with Cd concentrations between 10 and 50 mg kg⁻¹. The potential phytotoxicity of HANPs may be due to several factors, including their interaction with trace elements and soil components, which may limit the availability of nutrients to the plant, or cause structural alterations in the plants [52]. Root length and G_Index are susceptible indicators of heavy metal toxicity, as root tissues are directly exposed to contaminated soil. The observed inhibition of root growth in untreated soils at Cd concentrations ≥10 mg kg⁻¹ is consistent with the known toxic effects of Cd, such as alterations in cell division and elongation [28]. The improvement in G_Index with HANP treatments at moderate Cd concentrations (≤10 mg kg⁻¹) indicates the partial mitigation of these effects. However, the inhibitory trend at higher Cd concentrations (>10 mg kg⁻¹) suggests that HANPs may not completely shield root systems under extreme stress. It is noteworthy that, despite the variability of results in soils treated with HANPs, there were no significant differences in root growth, regardless of Cd concentration, compared to the control soil without Cd. This suggests an improvement in root development for quinoa seedlings in Cd-contaminated soils when HANPs were applied. On the other hand, the observed improvement in shoot growth and partial mitigation of root inhibition at moderate Cd concentrations (≤10 mg kg⁻¹) may have significant implications for crop productivity. By limiting Cd toxicity, HANPs may enable the cultivation of crops in moderately contaminated soils, potentially reducing the risk of Cd entering the food chain. However, their limited effectiveness at high Cd concentrations (>10 mg kg⁻¹) underscores the need for complementary remediation strategies in highly contaminated areas. This may include using other phytoremediation plants or soil amendments with greater immobilization capacities.

3.4. Pot Assays

Pot experiments were conducted to evaluate the toxicity of Cd and the effects of HANPs on the germination, growth, and medium-term development of quinoa (Figure 1). The main results obtained are summarized in

Table 5. The germination and physiological characteristics of quinoa for different treatments with Cd and HANPs in pot experiments at the end of the assays.

Soil + HANPs	Cd (mg kg−1)	Germination (%)	Fb (g)	Db (g)	h (cm)
-	0	92 ± 8 a	16.4 ± 0.5 a	1.6 ± 0.4 a	48.9 ± 8.2 a
1%	0	95 ± 3 a	15.3 ± 1.2 a	2.4 ± 0.4 a	48.3 ± 2.5 a
-	5	95 ± 5 a	12.5 ± 2.5 b	1.1 ± 0.6 b	39.2 ± 13.2 ab
1%	5	95 ± 2 a	14.0 ± 2.4 ab	2.0 ± 0.5 b	47.0 ± 1.3 a
-	10	75 ± 14 b	10.4 ± 2.8 bc	1.1 ± 0.4 bc	33.5 ± 7.1 bc
1%	10	75 ± 12 b	12.9 ± 2.4 b	0.9 ± 0.3 c	47.3 ± 4.2 a
-	25	58 ± 10 bc	6.0 ± 2.9 c	0.6 ± 0.1 cd	26.5 ± 4.7 bc
1%	25	55 ± 9 c	7.2 ± 2.0 c	0.8 ± 0.2 cd	33.1 ± 9.1 bc
-	50	42 ± 16 d	7.0 ± 0.8 c	0.7 ±0.3 cd	25.1 ± 3.8 c
1%	50	60 ± 4 c	3.1 ± 1.4 d	0.5 ± 0.2 d	20.1 ± 4.7 c
Soil + HANPs	Cd (mg kg−1)	La (cm2)	N° leaves	Root-Cd (mg kg−1)	A.p.-Cd (mg kg−1)
-	0	11.3 ± 2.6 a	31 ± 5 b	nd	nd
1%	0	8.0 ± 2.2 ab	37 ± 3 a	nd	nd
-	5	6.6 ± 4.3 ab	39 ± 4 ab	nd	55.1 ± 17.0 c
1%	5	7.8 ± 5.1 ab	42 ± 3 a	nd	16.4 ± 3.6 d
-	10	8.5 ± 3.6 ab	30 ± 2 b	nd	89.5 ± 36.3 bc
1%	10	6.4 ± 5.0 ab	31 ± 5 b	nd	27.6 ± 8.8 d
-	25	5.4 ± 2.1 b	24 ± 1 d	nd	153.8 ± 45.2 ab
1%	25	5.8 ± 4.0 ab	35 ± 3 b	nd	56.0 ± 3.1 c
-	50	4.8 ± 3.2 b	28 ± 2 c	nd	215.7 ± 43.3 a
1%	50	3.7 ± 3.0 b	26 ± 2 c	nd	91.9 ± 10.2 b

Fb: fresh biomass; Db: dry biomass; h: height. La: leaf area; N° leaves: average number of leaves per individual; Root-Cd: Cd content in root; A.p.-Cd: Cd content in aerial part; nd: not detected. Different letters next to the values in each column represent significant differences among treatments for each measured variable at a significance level of p < 0.05.

.

Germination rates in pot experiments differed from those observed in phytotoxicity assays. In pots, quinoa seed germination decreased with increasing Cd concentration, regardless of HANP application, from 10 mg kg⁻¹ Cd. This difference might be due to direct contact between seeds and soil components in pot experiments. In contrast, in the phytotoxicity assays, the black filter acted as a barrier, isolating the seeds from direct interaction with the soil. Consequently, in phytotoxicity assays, only the soil solution and its ions influenced germination rates.

Regarding fresh and dry biomass (stems and leaves), no significant differences were observed between individuals grown in soils treated with and without HANPs at the same Cd concentrations. However, biomass decreased as Cd concentration increased, regardless of HANP application. This indicates that the presence of HANPs did not substantially influence quinoa plant biomass. For plant height, individuals from control soils (0 mg kg⁻¹ Cd) showed no significant differences between treatments with or without HANPs. However, for 5 and 10 mg kg⁻¹ Cd with HANPs, plant height was similar to the control height, whereas, for treatments with 25 and 50 mg kg⁻¹ without HANPs, plant height significantly decreased. In this line, at Cd 25 and 50 mg kg⁻¹ concentrations with HANPs, plant height was also significantly reduced, underscoring their limited effectiveness under high Cd contamination levels. This suggests that HANPs may mitigate Cd toxicity at moderate contamination concentrations (≤10 mg kg⁻¹) but are less effective at higher Cd concentrations (>10 mg kg⁻¹).

The decrease in leaf area and leaf number at higher Cd concentrations shows the negative effects of Cd on the photosynthetic activity and health of quinoa. Cadmium toxicity is known to affect chlorophyll synthesis, affect water transport, and alter enzymatic activities involved in carbon assimilation [53]. The presence of HANPs mitigated these effects at Cd concentrations between 5 and 25 mg kg⁻¹, possibly by providing essential calcium and phosphorus ions that ameliorate Cd-induced nutrient deficiencies. However, at 50 mg kg⁻¹ Cd, the mitigation capacity of HANPs was limited, showing the need for more robust remediation strategies at high contamination levels.

Root Cd concentrations were below the detection limit for all treatments, while individuals from control soils with or without HANPs showed no Cd accumulation in shoots. Cd content in quinoa shoots was lower in soils with HANPs than soils without HANPs at the same soil Cd concentration. The reduction in Cd content in shoots observed in this study is consistent with the ability of HANPs to immobilize Cd by forming stable phosphate complexes, which reduce its bioavailability. This mechanism aligns with findings indicating that HA effectively reduces the availability of exchangeable and soluble fractions of Cd in contaminated soils [49].

Recent studies have demonstrated the efficacy of HANP to immobilize Cd in contaminated soils, favoring the growth of different plants and improving the yield of several crops. For example, the application of HANPs significantly reduced Cd availability (≈90%) in soils, enhancing the germination and early growth of Sinapis alba L. [13]. Similarly, HA combined with a hydrated lime and an organic fertilizer were effective in decreasing Cd uptake in rice and wheat under a rice–wheat rotation system, leading to improved crop yields [54]. Likewise, HA and humic acid application on alkaline soils contaminated with Cd also led to significant reductions in Cd concentrations in grains, shoots, and roots of wheat [55]. Other studies also showed that the co-application of HANPs with other (nano)materials improved Cd fixation in contaminated soils. For example, the combined application of zinc oxide and HANPs enhanced Cd immobilization and wheat growth, indicating that synergistic effects between different nanoparticles may be beneficial [56]. Also, combining a porous biochar composite with HANPs effectively immobilized Cd in soils [57], and combining HANPs with Fe₃O₄ nanoparticles improved the stress caused by high Cd concentrations in rice seedlings, promoting growth and reducing metal accumulation [58].

However, our results indicate that Cd content in quinoa increases with the concentration of Cd applied to the soil, suggesting that while HANPs can reduce Cd bioavailability, their efficacy may vary depending on plant species and specific soil conditions. Consistent with these results, a previous study showed that the effectiveness of biochar-enriched soils in immobilizing potentially toxic elements, including Cd, depends on the soil properties and the nature of the contaminants [59]. The selection of application rates of HANs as phosphorus fertilizers and Cd immobilizers is essential to avoid their potential phytotoxicity to HNAPs [60].

3.5. Results of Statistical Analysis

Statistical data analysis provides a detailed understanding of the significant effects of hydroxyapatite nanoparticles (HANP) and added Cd concentrations on soil properties, metal immobilization, and quinoa response. The soil characterization results indicated that the sandy-loam-textured soil had a low organic matter content and moderately acidic pH, which significantly influenced the mobility of Cd in the soil. The low ECEC, where exchangeable Al³⁺ predominates, limits the availability of sorption sites for Cd retention.

Sequential chemical extraction showed that Cd distribution among geochemical fractions varied significantly between treatments. Soils treated with Ca but not with HANPs showed a high proportion of Cd in the most bioavailable fractions (F1–F3), ranging from 36.6% to 60.4% of total Cd, depending on the Cd concentration applied. In contrast, soils treated with HANPs showed a redistribution of Cd towards less bioavailable fractions, particularly the irreversibly adsorbed fraction (F6). At 50 mg kg⁻¹ Cd, the F6 fraction increased significantly from 5.86 mg kg⁻¹ in untreated soils to 8.96 mg kg⁻¹ in soils treated with HANPs. This change shows the ability of HANPs to promote the formation of stable Cd–phosphate complexes, reducing metal mobility and environmental risks of contamination.

In phytotoxicity tests, statistical comparisons revealed significant differences in germination percentage (G%), root and shoot length, and G_Index and Ap_Index between treatments. In untreated soils, Cd concentrations above 10 mg kg⁻¹ significantly reduced shoot length and Ap_Index values compared to the control. For example, Ap_Index decreased from 100% in control soils to 70% at 50 mg kg⁻¹ Cd. However, the application of HANPs mitigated these effects at moderate contamination levels. At 10 mg kg⁻¹ Cd, Ap_Index increased significantly from 84% in untreated soils to 119% in HANP-treated soils. Root growth was particularly sensitive to Cd toxicity, with G_Index values below 90% at 10 mg kg⁻¹ Cd or higher. HANPs partially improved root growth inhibition, as evidenced by G_Index values, which remained significantly lower than those of the control.

Pot assays confirmed the mitigating effects of HANPs on Cd toxicity under controlled conditions. Germination rates in pots decreased significantly with increasing Cd concentrations, particularly above 10 mg kg⁻¹. Fresh and dry shoot and root biomass also showed significant reductions at higher Cd levels, regardless of HANP application. However, HANP-treated soils showed a statistically significant improvement in plant height and Cd content in shoots compared to untreated soils at equivalent Cd concentrations. For example, at 25 mg kg⁻¹ Cd, HANP reduced Cd accumulation in shoots from 153.8 mg kg⁻¹ in untreated soils to 56.0 mg kg⁻¹ in treated soils, highlighting its potential to limit Cd translocation to aboveground plant tissues.

The statistical analysis indicates the efficacy of HANPs in mitigating Cd toxicity through its immobilization and redistribution to less bioavailable soil fractions. Significant differences between treatments demonstrate that nanoparticles effectively reduce Cd bioavailability, improve plant growth parameters, and limit Cd translocation under moderate levels of contamination. However, their lower effectiveness under high levels of Cd concentrations suggests the need for complementary strategies, particularly for soils with elevated Cd concentrations.

4. Conclusions

Phytotoxicity tests demonstrated that HANPs do not enhance quinoa seed germination or early root growth and, in some cases, exhibit an inhibitory effect. However, at Cd concentrations of 10 mg kg⁻¹ or higher, HANPs significantly contributed to early shoot development, improving the Ap_Index.

Sequential chemical extraction revealed that HANPs do not significantly affect the most bioavailable soil fractions (F1–F3). However, the Cd content strongly complexed with organic matter (F5) decreased in soils treated with HANPs, while irreversibly adsorbed Cd (F6) increased, suggesting the formation of stable, insoluble Cd–phosphate complexes.

Germination rates in pot experiments differed from those in phytotoxicity tests, likely due to the direct interaction of seeds with the soil in the former. No significant differences in the physiological characteristics of quinoa plants were observed between HANP-treated and untreated soils. However, Cd content in the aerial parts of quinoa plants significantly decreased in HANP-treated soils, indicating reduced Cd translocation, likely due to its immobilization associated with HANPs.

The results confirm the potential of HANPs to reduce Cd bioavailability in contaminated soils and limit its accumulation in quinoa tissues. However, their limited efficacy in reducing highly bioavailable Cd fractions at elevated contamination levels highlights the need to combine their use with complementary remediation strategies, such as organic amendments.

This study establishes a basis for understanding the potential of HANPs in agricultural remediation. Future investigations may focus on field assays to assess the long-term impact of HANPs on quinoa development, including their effects on seed safety and quality for human consumption, as well as the assessment of their combined use with other remediation strategies to optimize effectiveness under varying Cd contamination levels.