Nanocellulose enhances the dispersion and toxicity of ZnO NPs to green algae Eremosphaera viridis

Abstract

The widespread cellulose nanomaterials from industrial production and natural plant degradation inevitably lead to the accumulation of nanocellulose in the aquatic environment. However, the effect of nanocellulose on the fate, transport and biotoxicity of zinc oxide nanoparticles (ZnO NPs) remains largely unexplored. The present study investigated the interactions between nanocellulose and ZnO NPs. The addition of naturally-derived cellulose nanocrystals (CNCs) significantly reduced the aggregation of ZnO NPs, resulting in enhanced bioavailability and toxicity to the algae Eremosphaera viridis. The ZnO–CNC association enhanced the envelopment of the algal cells and exerted strong oxidative stress as compared to bare ZnO NPs. The excessive reactive oxygen species generation could result in the breakdown of membrane lipid and disruption of antioxidant enzyme activity, where lipid synthesis was inhibited and protein folding might occur. The presence of CNCs also improved the concentration of Zn ions inside the algal cell through intracellular transportation. This can affect the flow of substances between algae cells and the environment, and further influence the metabolism of microalgae. This work is crucial for improving our insight into the mechanism for combined effects from multiple nanomaterials, such that the composite risks of such combined effects to aquatic organisms can be identified.

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Environmental significance

Nanocellulose is an emerging nanomaterial that can be generated from various natural resources. It has been applied in the fields of textile, paper manufacturing, and food packaging. Nanocellulose may come into contact with the environment during production, with the waste streams or even the degradation of aquatic plants. Natural celluloses are ubiquitous in surface water. They exhibit a strong adsorption affinity for various engineered nanomaterials. Widespread ZnO NPs will inevitably enter the freshwater ecosystem and interact with nanocellulose, altering the fate, transport, and toxicity of the nanomaterial mixtures. Therefore, it is important to evaluate their combined toxicity to aquatic organisms, such that the environmental risk of the nanomaterials can be better identified.

1. Introduction

With the mass production and application of metal oxide nanoparticles in various fields, their release into the aquatic environment has provoked increasing concern. Among the diverse nanomaterials, zinc oxide nanoparticles (ZnO NPs) have broad applications in industrial and daily supplies.^1,2 They have been widely used in consumer products such as paint, coatings, cosmetics, pharmaceuticals, and electronics, with an estimated output of 30 000 tons per year.³ The extensive use of ZnO NPs elevates the probability of their exposure in the natural environment, leading to adverse effects on various organisms.⁴ Given the ever-increasing environmental risks posed by ZnO NPs, a large number of studies have been conducted to evaluate their toxicity toward aquatic organisms at different trophic levels.^5–8 However, the vast majority of these studies have focused on the toxic effect of individual ZnO NPs, neglecting the interactions between ZnO NPs and coexisting materials in complicated aquatic systems.⁹ Therefore, further investigations regarding the combined toxicity of multiple contaminants are indispensable to obtain a more complete picture of their ecological risks.

ZnO NPs integrated with diverse environmental contaminants can undergo complex physicochemical transformations, including adsorption, coagulation, and dispersion, which may exert synergistic or antagonistic toxic effects on organisms.^10,68 For example, Wang et al.¹¹ indicated that surfactants of different ion types presented antagonistic effects on ZnO toxicity to bacteria P. phosphoreum. This joint effect was attributed to the interactions between the surfactants and dissolved Zn ions. Tong et al.¹² found that the combined toxicity of ZnO and TiO₂ NPs towards bacteria E. coli was more attenuated than that of individual nanoparticles. This could be explained by the reduced bacteria/nanoparticle contact and adsorption between TiO₂ NPs and Zn ions. Kteeba et al.¹³ reported that the presence of dissolved organic matter could alleviate the toxicity of ZnO NPs to zebrafish. This mitigation effect was primarily derived from ZnO NPs rather than the dissolved Zn ions. In a recent study, Ye et al.¹⁴ tested the toxicity of graphene oxide nanoplatelets and ZnO NPs to three aquatic species (alga, cladoceran, and freshwater fish larva). The results revealed that the joint effects were antagonistic to D. rerio but additive to S. obliquus and D. magna. Despite these studies, the behaviors and toxicity of binary nanoparticle mixtures remain largely unexplored.

Cellulose nanocrystals (CNCs) are an emerging type of nanomaterial that can be generated from various natural processes (e.g., hydrolysis of wood, grass, and agricultural waste).^15,16 This nanomaterial has attracted significant interest due to its broad availability and outstanding properties, such as a large surface area, good mechanical properties, biodegradability, and negligible toxic potential in regard to the environment.¹⁷ Nanocellulose materials have been applied in the field of textile, paper manufacturing, food packaging, and pharmaceutical industries.^18,19 The annual production of cellulose and its derivatives was reported to be one trillion tons in 2020.²⁰ This will unavoidably lead to the accumulation of nanocellulose in the natural environment. Cellulose nanomaterials may come into contact with the environment during production, with the waste streams or as accidental release.²¹ Apart from the industrial discharge of cellulose-containing wastewater, natural biodegradation of aquatic plants can also cleave cellulosic biomass into smaller fragments.²² Suaria et al.²³ compiled a global dataset from 916 seawater samples in six ocean basins and revealed that most of the oceanic fibers were natural cellulose (79.5%). However, the large number of natural cellulosic fibers in water systems is greatly underestimated in the environmental pollution literature.²⁴ In addition, cellulose nanomaterials are not inert and may act as vectors for other hazardous chemicals into the environment.^25,67 Therefore, it is important to raise the safety concern over CNCs since various organisms are under exposure to such abundant natural polymers.

To date, most attention has been dedicated to the toxicity of single ZnO NPs to freshwater algae. Research on the combined toxicity of ZnO NPs mediated by natural nanocellulose materials is still very limited. This is the first study to investigate the biotoxicity of ZnO NPs as affected by CNCs. In addition, previous studies mainly explain the mechanisms from community and cellular levels. This study provides a new perspective to analyze the toxic effect in a single cell through synchrotron-based FTIR and XRF mapping techniques, such that the toxicity mechanism at the molecular level can be revealed. The objective of this study is to conduct a systematic assessment of ZnO NP toxicity in the presence of CNCs and explore the mechanisms responsible for their interactive toxic effects. Various physicochemical properties are characterized to analyze the interactions between ZnO NPs and CNCs. Multiple biomarkers are investigated to evaluate the cytotoxic effects of the ZnO–CNC nanohybrid on algal biomass. Synchrotron-based FTIR and XRF mappings are implemented to analyze the biochemical alteration and elemental distribution in a single living cell. This work is crucial for improving our insight into the mechanism for combined effects from multiple nanomaterials, such that the composite risks of such combined effects to aquatic organisms can be identified.

2. Experimental

2.1. Materials and characterization

The CNCs were purchased from CelluForce Inc. (Montreal, Canada). Their detailed structure and characterization are shown in Fig. S1.† ZnO NPs were purchased from Innochem, China. The CNC (1.5 g L⁻¹) and ZnO stock solutions (200 mg L⁻¹) were prepared in pure water and subjected to water bath sonication for 30 min at 40 kHz (Branson M5800H ultrasonic cleaner bath, USA). The size and morphology of ZnO NPs were determined using scanning electron microscopy (FEI Quanta 400, USA). The hydrodynamic diameter and zeta potential of the NPs, either alone or as a mixture, were further analyzed with a Litesizer™ 500 analyzer (Anton Paar, Austria). To investigate the interactions between ZnO and CNCs, various techniques, including XRD, FTIR, XPS, UV-vis spectroscopy, and SEM, were used to characterize the ZnO, CNCs, and ZnO/CNCs. The details are provided in ESI, Text S1.† All other chemicals were of reagent grade or higher.

2.2. Algae culture and toxicity tests

Microalgae Eremosphaera viridis (Fig. S2†) were used as an ecological indicator, based on their characteristics in terms of large size, toxicant sensitivity, and primary role in the food chain.²⁶ The freshwater algae were purchased from the Canadian Phycological Culture Center at the University of Waterloo, Canada. They were cultured in an Erlenmeyer flask with 100 mL Bold's basal medium (BBM). The temperature of the culture medium was maintained at 23 °C with a photoperiod of 12/12 h light/dark cycle. The algae were collected at the exponential growth phase with an initial concentration of 10⁴ to 10⁵ cells per mL. ZnO was added to the culture medium with concentrations of 1, 5, and 10 mg L⁻¹, in the absence or presence of 100 mg L⁻¹ CNCs. Although the predicted environmental concentrations of these two nanoparticles were rather low, they were expected to increase continuously in the future.^27,28 The dissolved Zn²⁺ concentration was quantified using inductively coupled plasma mass spectrometry (PerkinElmer DRC II, USA). The detailed Zn adsorption experiment is shown in ESI,† Text S2. After 72 h of exposure, the algal dry weight was determined to evaluate the algal growth inhibition, as shown in ESI,† Text S3.

2.3. Biochemical indicators and cellular ultrastructure/physical damage

After 72 h of exposure to a ZnO–CNC mixture or individual NPs, the chlorophyll a/b concentration in algae was measured with a UV-vis spectrophotometer (Agilent Technologies, USA);²⁹ the method is specified in ESI,† Text S3. The generation of intracellular reactive oxygen species (ROS) was detected by the H₂DCFDA staining method (ESI,† Text S4), and the corresponding fluorescence images of algal cells were recorded with a fluorescence microscope (Zeiss Axio Observer Z1, Denmark). The CAT activity levels and MDA content were also determined by commercial assay kits from the Nanjing Jiancheng Institute of Biotechnology (ESI,† Text S5 and S6), according to the manufacturer's protocols. The algal morphology and cellular ultrastructure change after exposure to 5 mg L⁻¹ ZnO or ZnO–CNCs for 72 h were observed by SEM (Hitachi SU8010, Japan) and TEM-EDS (FEI Tecnai Spirit, USA), respectively. The details are presented in ESI,† Text S7.

2.4. Synchrotron-based FTIR spectromicroscopy and X-ray fluorescence imaging

The synchrotron-based FTIR spectromicroscopy (SR-FTIR) and X-ray fluorescence (SR-XRF) mapping analysis were implemented at the Canadian Light Source in Saskatoon, Canada (Fig. S2†). The composition and spatial distribution of macromolecules in a single living algal cell were determined under the mid-IR beamline (01B1-1). Imaging live cells is crucial to unravel how their chemical composition changes under toxic exposure. SR-FTIR gives a higher brightness and better signal/noise ratio than conventional infrared light, satisfying the need for a single cell resolution and higher spectra quality without damaging any biological samples.³⁰ At least 10 spectra for each sample were collected to generate the average spectra. The detailed sample preparation is shown in Text S8.† Resolving the subcellular distribution of Zn and other trace metal elements is essential to understand the mechanisms involved in their accumulation and detoxification in the microalgae. SR-XRF mapping was further carried out at the VESPERS beamline (07B2-1) to identify multi-element distribution and co-localization in algal cells. SR-XRF is much better than conventional X-ray sources because of its small source size, high spatial resolution (few μm to tens of nm), and reasonable acquisition times (≤1 s per pixel).²⁹ All measurements were performed in air, and the fluorescence images were recorded by a four-element silicon drift Vortex detector.³¹ The algal samples after 72 h of exposure were washed with distilled water three times, and the obtained algal cells were freeze-dried for 24 h. Dried samples were finally fixed on double-sided Kapton tape for SR-XRF imaging. The element distributions of K, Ca, Fe, Zn, Cu, and Mn in a single algal cell were recorded within a 120 μm × 120 μm scanned area using a step-size of 5 μm.

2.5. Statistical analysis

All experiments were conducted in three replicates, and the data were presented as the mean ± standard deviation. One-way analysis of variance (ANOVA) followed by Fischer's least significant difference (LSD) test was used to determine the significant difference among treatments (p < 0.05). All data analyses were performed using SPSS Statistics 19.0 (IBM, USA). The FTIR spectra were analyzed using OPUS 7.2 software (Bruker Optics Inc., USA), and the FTIR images were processed with CytoSpec 2.0 software (Cytospec, USA). The XRF images were generated using SigmaPlot (Systat Software Inc., USA). Principal component analysis (PCA) was performed by SPSS 19.0 to visualize the correlation of relevant responses. All other experimental data were processed using OriginPro 8.0 software (OriginLab Corp., USA).

3. Results and discussion

3.1. Dissolved Zn concentration

Measurements of dissolved Zn present in different ZnO NP suspensions (1 mg L⁻¹, 5 mg L⁻¹ and 10 mg L⁻¹) were quantified by ICP-MS (Fig. S3†). Results showed that 31%, 26% and 17% of Zn were already dissolved when the initial exposure concentration was 1 mg L⁻¹, 5 mg L⁻¹, and 10 mg L⁻¹. After 72 h of incubation, the percentage of dissolved Zn in the ZnO NP suspensions shifted to 82%, 36%, and 22%, respectively. This indicated that the dissolution of Zn²⁺ from ZnO NPs was inhibited at higher initial particle concentration, which could be explained by the reaction-induced proton depletion, oxygen depletion, or inhibition of surface reactions.³² Meanwhile, the negatively charged CNC has good adsorption capacity towards Zn ions. The addition of 100 mg L⁻¹ CNC dramatically increases the dissolution of Zn ions from ZnO NPs. After 1 h, the actual dissolved Zn in the 1 mg L⁻¹, 5 mg L⁻¹, and 10 mg L⁻¹ ZnO suspensions reached up to 71%, 68%, and 71%, respectively. The unique template structure of CNC is favorable to control the physical deposition of nanoparticles, greatly inhibiting the aggregation and improving the dispersion uniformity and stability of nanoparticles.³³ This meant that the presence of CNC could inhibit the sedimentation of ZnO NPs, and subsequently enhance the dissolution of Zn ions from ZnO NPs. The molecular interactions, morphological alterations, and hydrodynamic diameter changes of ZnO NPs after CNC adsorption are reported in the following section.

3.2. Binding interaction of ZnO with CNC

The FTIR spectra of ZnO, CNC, and ZnO–CNC are displayed in Fig. 1a. The spectrum of ZnO shows a characteristic absorption peak at 533 cm⁻¹, which was ascribed to the transverse optical stretching modes of ZnO.³⁴ For CNC, the peaks at around 3333 cm⁻¹, 2893 cm⁻¹, 1636 cm⁻¹, 1158 cm⁻¹, 1053 cm⁻¹, and 898 cm⁻¹ can be attributed to the stretching vibration of O–H, asymmetric stretching of C–H, O–H bending of absorbed water, C–C ring stretching, C–O–C stretching of the pyranose ring in the skeletal vibration, and C–H rocking, respectively.³⁵ Compared with CNC, a new peak at 477 cm⁻¹ was observed in ZnO–CNC, which was assigned to Zn–O stretching vibration. Besides, the introduction of ZnO weakened the intensity of the O–H stretching vibration and shifted the wavenumber to a lower value, confirming the interactions between hydroxyl groups of CNC and ZnO NPs. Fig. 1b displays the deconvolution of the XPS spectrum (O 1s) for the ZnO–CNC nanocomposite, which can be well fitted with two symmetrical peaks located at 530.65 and 532.50 eV, respectively. These two peaks corresponded to two different kinds of O species in ZnO–CNC. The low energy peak belonged to the lattice oxygen of ZnO, while the high energy peak was associated with the chemisorbed oxygen which arose from the surface hydroxyl group.^36,37 The evidence from the XPS results also indicated an interaction between CNC and ZnO NPs. The UV-vis absorption spectra of single nanoparticles and their nanohybrids are presented in Fig. 1c. A typical absorption peak at 363 nm was assigned to the basic bandgap absorption of ZnO due to electron transitions from the valence to the conduction band (O 2p to Zn 3d).³⁸ Interestingly, a blue-shift (about 8 nm) in the peak of the ZnO–CNC nanocomposite was observed, which was ascribed to the decreased size of ZnO NPs.³⁹ The XRD pattern of the nanoparticles is shown in Fig. S4.† According to the Scherrer equation, the average crystallite size of ZnO (11 nm) and ZnO–CNC (10 nm) was calculated from the most intense peak at 2θ = 36.4°. This suggests that a smaller crystallite size could be obtained through the adsorption of CNC molecular chains on the surface of ZnO NPs.³⁴ The nanoparticle morphology and size distribution are further supported by the SEM observations and dynamic light scattering (DLS) technique presented below.

Fig. 1 Characterization of ZnO, CNC, and ZnO–CNC. (a) FTIR spectra; (b) XPS spectra (O 1s); (c) UV-vis absorption spectra; (d–f) SEM images; (g) hydrodynamic diameter; (h) zeta potential. The “*” symbols denote p-values <0.05 compared to the treatment at 1 h.

The SEM image of ZnO NP powder (Fig. 1d) demonstrates irregular spherical-like structures which are intensely aggregated over the size range between 2.0 and 5.0 μm. CNC NP powder is displayed as rod-like structures adhering to each other (Fig. 1e), and the size of the nanocrystals (∼100 nm) was confirmed by AFM, as shown in Fig. S1.† Upon the precipitation reaction between ZnO and CNC, homogeneous dispersed ZnO NPs with an average diameter of 1 μm were anchored on the surface of CNC (Fig. 1f). Compared with the original ZnO NPs, the ZnO–CNC nanocomposite hindered the aggregation process and reduced the size distribution of ZnO NPs. Also, the morphology of CNC changed to flat elongated ribbon-like membranes after the hydrolysis reactions, which might have enhanced the dispersion of ZnO NPs through the repulsive interaction of a negatively charged end group on CNC.⁴⁰ Fig. S5† shows the elemental distribution in the ZnO–CNC nanocomposite. A good correlation between Zn, O, and C was observed, indicating the homogeneous binding of ZnO and CNC in the binary system.

Fig. 1g shows the hydrodynamic diameter of ZnO NPs, CNC, and the ZnO–CNC nanohybrid in the culture medium at 1 h and 72 h. The size of 1 mg L⁻¹ ZnO NP agglomerates was 2005 nm at 1 h. However, due to the homoagglomeration favored by nanoparticle–nanoparticle interactions,⁴¹ a small increase in the size (2900 nm) of ZnO NPs was observed at 72 h. The change in hydrodynamic diameter was dose-dependent, as an obvious increase was observed with the rising ZnO NP concentration. It was interesting to note that the introduction of CNC into the ZnO NP suspension dramatically reduced the hydrodynamic diameter of the ZnO–CNC nanocomposite (300 nm for 1 mg L⁻¹ ZnO–CNC at 1 h), and the size of this association remained stable up to 72 h without significant aggregation.

Both ZnO NP and CNC suspensions were negatively charged, as determined by the zeta potential in Fig. 1h. The zeta potential decreased with increasing ZnO NP concentration, and slightly increased after 72 h due to the positively charged ions binding on the surface of the nanoparticles. The 1 mg L⁻¹ ZnO–CNC nanohybrid (−30.9 ± 0.7 mV) exhibited a higher absolute zeta potential than ZnO (−26.2 ± 0.9 mV) after 72 h. Nanofluids with an absolute zeta potential value of more than 30 mV are considered as stable colloidal dispersions,⁴² indicating that the ZnO–CNC nanofluid was more stable than the ZnO suspension in the BBM medium. This phenomenon was also confirmed by direct observation of the nanoparticle suspensions (Fig. S6†). These findings are consistent with previous studies indicating that CNC could promote the dispersion stability of nanoparticles.⁴³ In order to eliminate the influence of ionic strength induced by the culture medium, the samples were also prepared in MilliQ water for size and zeta potential measurement (Fig. S7†). A significant decrease in hydrodynamic diameter and an increase in absolute zeta potential were observed after CNC was introduced into the ZnO suspension.

3.3. CNC enhanced the algal toxicity of ZnO

After 72 h of exposure, the addition of CNC had negligible effects on the cell dry weight due to its non-toxic nature (Fig. 2a). The biomass weight decreased with increasing ZnO exposure concentration. Meanwhile, the dry weight of cells co-exposed to ZnO and CNC was significantly lower than that of cells exposed to individual ZnO or CNC, indicating an enhanced toxic effect on algal biomass. This might be attributed to the interaction between ZnO and CNC, leading to a reduced hydrodynamic diameter of ZnO in the CNC matrix and an enhanced dissolution of Zn ions from ZnO NPs. Since the ZnO toxicity may be attributed to dissolved Zn ions, insoluble NPs, or both particles and released ions, we conducted another comparison study using Zn ions as controls (Text S9†). The findings indicated that the released Zn ions had the most contribution to the toxic effects. As shown in Fig. 2b, there was a decreasing trend in the chlorophyll a content of the treatment groups compared with the control, presumable because Zn ions inhibited the main pigment synthesis in the photosynthesis process. The decrease of the chlorophyll a concentration was more obvious under the coexistence of ZnO and CNC, which was consistent with the results of the dry weight. However, the reduction of chlorophyll a was not as significant as the cell density. This may be due to the fact that chlorophyll a plays an important role in the inhibition of lipid peroxidation. This defense mechanism made them more active under toxic stress. Chlorophyll b was the accessory pigment involved in photosynthesis, passing the trapped energy into chlorophyll a.⁴⁴ A similar decreasing trend was found between chlorophyll b and a (Fig. 2c), which implied that the accessory pigment synthesis was affected in the algal cells.

Fig. 2 Algal toxicity of ZnO, CNC, and ZnO–CNC at 72 h. (a) Dry weight; (b) chlorophyll a content; (c) chlorophyll b content; (d) relative ROS level; (e) CAT activity; (f) MDA content; (g–i) SEM images of algal cells. The “*” symbols denote p-values <0.05 compared to the control.

The generation and clearance of ROS are in a dynamic equilibrium state in the algae. However, adverse conditions such as contaminant exposure can induce the excessive production of ROS, resulting in oxidative damage and even apoptosis of algal cells.⁴⁵ As shown in Fig. 2d, the ROS production exhibited a dose–response relationship. All the treatment groups induced significantly higher ROS levels than that of the control, except for the exposure to 1 mg L⁻¹ ZnO. In addition, ZnO–CNC induced a higher ROS level than bare CNC and ZnO. The generation of ROS was further examined under fluorescence microscopy (Fig. S8†). The algal cell exhibited intense green fluorescence upon exposure to 5 mg L⁻¹ ZnO and ZnO–CNC, as compared to the control group. The ROS production mainly occurred in the chloroplast, nucleus, and plasma membrane, which would lead to the disruption of enzyme and lipid function.

It has been reported that algae can activate antioxidant enzymes under environmental stress, removing excess ROS and protecting the cell from damage.⁴⁶ Catalase (CAT) is directly involved in cellular antioxidant defense mechanisms, responsible for the decomposition of intracellular hydrogen peroxide into water and oxygen.⁴⁷ As shown in Fig. 2e, the group treated with CNC and 1 mg L⁻¹ ZnO showed higher CAT activity than the control, due to the production of excess ROS in algal cells. However, the CAT activity exhibited a significant decline when the concentration of ZnO exceeded 5 mg L⁻¹. Moreover, ZnO–CNC co-exposure had lower enzyme activity than bare ZnO. The rapid decrease in CAT could be attributed to the death of algae upon constant exposure to a large amount of ROS rather than recovery from or tolerance to them.⁴⁸ This trend corresponded with previous studies in which antioxidant enzyme activities in algae increased at low concentrations of xenobiotics but decreased at exposure to high concentrations.^47,49 MDA content as a biomarker of membrane damage has been widely used to quantify the degree of lipid peroxidation (Fig. 2f). After 72 h of exposure, both CNC and ZnO groups triggered membrane damage relative to the untreated control. The MDA content increased in response to the increasing concentration of ZnO, which was in agreement with the upregulation of ROS in the algal cells. This implies that the accumulation of free radicals causes membrane lipid peroxidation and elevates the content of MDA. However, an opposite trend was observed in the co-exposure group of ZnO–CNC. The level of MDA in the 10 mg L⁻¹ ZnO–CNC treatment group was significantly lower than that of the control. This suggests that the structure of algal cells might be severely damaged by the overgeneration of ROS, leading to the leakage of intracellular matter and decrease of MDA, CAT, and chlorophyll.

The morphology and ultrastructure changes of the algal cells after exposure to ZnO and ZnO–CNC are visualized in the SEM (Fig. 2g–i) and TEM (Fig. S9†) images. The SEM images showed that the algal cells in the control group had an intact shape with irregular ruffles on the cell surface. However, following exposure to 5 mg L⁻¹ ZnO, the algae became relatively smaller, with blighted and wrinkled cell walls. Furthermore, the attachment of ZnO nanoparticles to the algal cell was observed (indicated in the yellow pane), which could be responsible for membrane damage. The interaction of ZnO and CNC led to severe damage to the cell surface, where most algae were shrunk. The yellow pane denotes the envelopment of algal cells by the ZnO–CNC hybrid. The presence of CNC enhanced the dispersion of ZnO and promoted the adsorption of ZnO on the algae surface. This would affect the flow of substances and energy between algae cells and the environment, reduce the fluidity of cell membranes, and influence the metabolism of the microalgae, finally leading to the growth inhibition of algae. Fig. S9a and b† show the TEM image of an untreated algal cell with chloroplasts, starch granules, pyrenoid, thylakoid, and other cytoplasmic organelles. No ultrastructure damage was found in the control group, and the organized thylakoids with lamellar structure were a typical indicator of a healthy and functional chloroplast. In contrast, after being exposed to 10 mg L⁻¹ ZnO NPs for 72 h, chloroplast shrinkage was observed, as shown in Fig. S9c and d.† In addition, a large number of black globular lipoproteins were detected at the curved regions of the thylakoid membrane, named plastoglobules (PG). The study conducted by Austin et al.⁵⁰ showed that the number of PG increased considerably during the upregulation of plastid lipid metabolism in response to oxidative stress. Therefore, it was confirmed that ZnO toxicity induced oxidative stress, which was consistent with the ROS levels in the algal cells. In Fig. S9e and f,† ZnO–CNC co-exposure leads to a greater loss of chloroplasts compared to the control and ZnO treatment groups. The thylakoids showed blurry boundaries and lysis in conjunction with the disappearance of starch granules as well as a significant increase in the number of PG. These changes indicate that both energy storage and lipid metabolism are affected by environmental stress. TEM-EDS is a useful tool to qualitatively monitor the uptake of NPs without the need to label. As shown in Fig. S9g,† Zn was identified in the PG of three different groups. The control microalgae contained low amounts of Zn, which was derived from the culture medium. More Zn was determined in the PG with the coexistence of ZnO–CNC than in the ZnO groups. Thus, it was inferred that the combined toxicity of ZnO–CNC would exert severe damage on algal metabolism.

3.4. Zn distribution and colocalization of multiple endogenous elements

Synchrotron-based XRF was a useful tool to investigate the distribution of Zn and multiple endogenous elements in a single algal cell, with or without the presence of CNC. Six elements – K, Ca, Fe, Zn, Cu, and Mn – were analyzed, as shown in Fig. 3. The elements appeared to have specific distribution patterns, some being partially co-localized or segregated. K plays a major role in the transportation of water and nutrients throughout plant cells and is required for the activity of several enzymes.⁵¹ Ca can be involved in regulating lipid synthesis, which is usually complexed with polyphosphate in algae cells.⁵² Fe, as a redox-active metal, has a typical function in the cellular respiration of the photosynthesis process. Up to 80% of the cellular Fe is found in the chloroplasts; therefore, its distribution is a good indicator of the chloroplast's location.⁵³ Zn is expected to bind and transport with more than 1200 proteins, forming the largest group of essential micronutrients–proteins, such as phytochelatins. Zn enzymes located outside the chloroplast have many functions, including protein synthesis, energy production, DNA-transcription, and RNA translation.⁵⁴ Cu also plays an important role in photosynthesis, oxidative stress protection, and mitochondrial respiration.⁵⁴ Mn is essential for plant metabolism, and Mn-containing superoxide dismutase can protect the cell from the damaging effects of free radicals.⁵⁵ To assess the correlations among the distribution in each cell, PCA was performed on the XRF imaging data set of a single cell after exposure to ZnO. As shown in Fig. S10,† the score plot was defined by the first and second principal components (PC1 and PC2). Altogether, PC1 and PC2 corresponded to 84.0% of the variance, ensuring the relevance of this two-dimensional projection space.⁵⁶ The six elements were grouped into three main clusters: K, Ca, Fe, Zn; Mn; and Cu. These three clusters matched the visual observation shown in Fig. 3 and could be interpreted as below.

Fig. 3 Elemental distribution in a single algal cell. (a) The untreated control group; (b) cell exposure to 5 mg L⁻¹ ZnO; (c) cell exposure to 5 mg L⁻¹ ZnO in the presence of 100 mg L⁻¹ CNC. Six cells from each treatment were analyzed to ensure the reproducibility of the collected data.

As shown in Fig. 3a, K was almost evenly distributed throughout the whole cell, with almost the same localization as Zn. They appeared to be concentrated in the cytosol and bind with several proteins to perform diverse activities. The Fe distribution was a good indicator of the chloroplast location, which occupies the major part of the cell. There was some co-localization of Fe with Zn, probably due to the small number of Zn proteins existing in the chloroplast. Some Fe and Zn hotspots were present in the algal cell, which was likely related to the efflux mechanisms as a means of detoxification. Several electron-dense spherical structures were presented in the XRF maps of Ca, presumably representing polyphosphate bodies that contain polyphosphate complexed with Ca.⁵⁷ Cu and Mn had patterned distributions, co-localizing to some extent, especially under the metal pressure of ZnO and ZnO–CNC. This was probably due to their detoxification roles in binding with superoxide dismutases (SODs), capable of scavenging damaging ROS.⁵⁸ After exposure to 5 mg L⁻¹ ZnO (Fig. 3b), the concentration of Zn significantly increased throughout the cells, and the associated Fe and Cu also increased, implying effects on the photosynthesis system and antioxidant behaviors. When 100 mg L⁻¹ CNC was introduced (Fig. 3c), more Zn was accumulated around the cell walls, indicating that the addition of CNC could enhance the bioavailability of Zn to algae cells. Small nanoparticles (5–20 nm) can pass the cell walls and enter into the algae through endocytosis, passive diffusion, and transport carrier proteins.⁵⁹ The interaction of ZnO and CNC might damage the cell walls, increase the ROS level, and disrupt enzyme functions, resulting in more negative effects on algae cells.

3.5. Biochemical change in a single living cell

FTIR microspectroscopy and imaging can provide unique fingerprints of the macromolecular composition and reveal biochemical changes in the biomass even without obvious physiological and morphological damage,⁶⁰ as well as allowing the detection of changes in the relative abundance of macromolecular pools such as lipids and proteins. The average FTIR spectra of these microalgae are shown in Fig. 4. The amide I band (around 1650 cm⁻¹) was due to the C=O stretching vibrations in the secondary structure (alpha helices and beta sheets) of the protein. The amide II band (around 1570 cm⁻¹) arose from a combination of C–N stretching and N–H bending vibrations of the protein backbone.⁶¹ The band at 1740 cm⁻¹ belonged to the vibrations of C=O in lipids. The bands at 3000–2800 cm⁻¹ were associated with the symmetric and antisymmetric C–H stretch from CH₂ groups in lipids. When the algae were exposed to different concentrations of ZnO NPs, they exhibited a dose-dependent response on specific biomass components. Fig. S11† displays the change of FTIR spectra induced by ZnO NPs with and without the presence of CNC. It is known that the intensity of the amine I and amid II bands is sensitive to the peptide–ion interaction.⁶² Under Zn stress, we observed a significant decrease in the amine I and amid II regions as the concentration of ZnO was elevated, indicating the alteration of protein conformation. The addition of CNC increased the bioavailability of Zn to algae, leading to a more obvious declining trend in protein. It is speculated that protein folding and aggregation may occur upon interaction with ZnO NPs.⁶³ This speculation was further supported in the following FTIR mapping analysis. The lipid synthesis was also inhibited by an increasing concentration of ZnO. The adsorption of methylene and ester carbonyl bands showed a slight decrease in all treatments, which might be due to the cell membrane damage caused by lipid peroxidation.⁶⁴

Fig. 4 SR-FTIR absorption spectra of blank algae E. viridis in four target regions.

Fig. 5 shows the FTIR mapping of proteins and lipids in a single algal cell under exposure to 5 mg L⁻¹ ZnO. It was noted that both amid I and amid II exhibited a diffused distribution in the untreated cells, whereas they uniformly accumulated at the center of the cytosol upon ZnO exposure. This might be related to the protein aggregation and folding with nanoparticle interaction. When cells were treated with both ZnO and CNC, the concentration of amid I and amid II became lower, due to the impact on the protein secondary structure, which is in accordance with the results of FTIR spectra. Lipid C=O and lipid C–H were mainly distributed at the periphery of the blank algae. However, a heterogeneous accumulation trend was found when the algae interacted with ZnO. It was noted that the distribution of overall lipids showed a smaller, more scattered pattern than the protein mapping, indicating that a lipid-rich organelle was present in the cell.⁶⁵ This result was consistent with the increasing trend of PG observed in the TEM analysis (Fig. S9†). The regions verified as having a high lipid concentration were also related to the dense packing of thylakoid membranes. Besides, there was some overlap between the profiles of chemical concentration of lipid and protein, which could be explained by the simultaneous occurrence of lipids and proteins in the chloroplasts. Conversely, regions which coincide with the nucleus were rich in proteins (histones) but poor in lipids.⁶⁶ This could explain the different distributions of proteins and lipids in the algal cell. From Fig. S12,† it was concluded that both protein and lipid levels were reduced with increasing dose of ZnO. The introduction of CNC could promote the oxidative stress and intensify the toxicity of ZnO to algal cells, resulting in the change of protein structure and damage to lipid synthesis.

Fig. 5 FTIR mapping of proteins and lipids in an algal cell without treatment, under exposure to 5 mg L⁻¹ ZnO, and co-exposure to 5 mg L⁻¹ ZnO and CNC. Six cells from each treatment were analyzed to ensure the reproducibility of the collected data.

3.6. Intracellular toxicity mechanisms

The interaction of ZnO with CNC enhanced the stability and dispersion of the suspension. It reduced the hydrodynamic diameter of ZnO NPs and increased the Zn²⁺ bioavailability and toxicity to the algae Eremosphaera viridis. The H–O functional group on the surface of CNC was a favorable binding site for ZnO NPs through electrostatic interactions. The role of ZnO–CNC association enhanced the envelopment of the algal cells and exerted strong oxidative stress as compared to bare ZnO NPs. The excessive ROS generation could result in the breakdown of membrane lipid and disruption of antioxidant enzyme activity, ultimately contributing to the inhibition of cell proliferation and the process of cell death. The presence of CNC also promoted the dissolution of ZnO NPs on the cell wall, resulting in a shift from the particle-related toxic effect to the ion-related toxic effect. The released Zn ions may transport inside the algae. This may affect the flow of substances and energy between algae cells and the environment, which consequently influence the metabolism of microalgae. From the molecular perspective, the released Zn ions could be immobilized inside the algae through a variety of metal binding proteins in the chloroplast and nucleus. The enzyme capable of scavenging damaging ROS could be altered upon interaction with Zn ions. Protein folding and aggregation might occur and thus cause the downregulated enzyme activity. The lipid synthesis was also inhibited, possibly due to the damage to the cell membrane and chloroplasts. Fig. 6 shows the schematic illustration of detailed intracellular toxicity mechanisms.

Fig. 6 Possible intracellular toxicity mechanisms to algal cells.

4. Conclusions

Natural cellulose is ubiquitous in surface water. It exhibits a strong adsorption affinity for various engineered nanomaterials. To the best of our knowledge, this is the first attempt to explore the combined toxicity of engineered NPs and nature-derived cellulose to green algae. Our study revealed that negatively-charged CNC could interact with ZnO NPs through electrostatic repulsion, reduce the homoaggregation of NPs, and promote the probability of contact with algal cells. The presence of CNC enhanced the bio-uptake of Zn and the joint toxicity to algal cells. The mechanisms involved in the toxic effects were clearly manifested in the upgraded level of oxidative stress, changed pattern of elemental distribution, disruption of enzyme activity, and damage to lipid synthesis. The findings of this work imply that current laboratory toxicological studies may underestimate the risk of engineered NPs in the aquatic environment. Similarly, the toxicity of ZnO NPs is largely governed by the behavior of other natural or engineered materials (e.g., clay, microplastics, and extracellular polymeric substances released by algae) in the exposure medium. The role of their associations in the toxicity assay deserves further investigation. Furthermore, there are many other factors contributing to the combined effects, such as the environmental pH, ionic strength, natural organic matter addition, and size and shape of the nanomaterials. These complex components will help us better understand the fate and biotoxicity of various pollutants under relevant environmental conditions.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

This research was supported by Canada Research Chair Program, Natural Science and Engineering Research Council of Canada, Western Economic Diversification (15269), Natural Science Foundation (U2040212), and MITACS. The synchrotron-based techniques were performed using beamline Mid-IR and VESPERS at the Canadian Light Source, a national research facility of the University of Saskatchewan, which is supported by the Canada Foundation for Innovation (CFI), the Natural Sciences and Engineering Research Council (NSERC), the National Research Council (NRC), the Canadian Institutes of Health Research (CIHR), the Government of Saskatchewan, and the University of Saskatchewan. We are also very grateful for the helpful inputs from the Editor and anonymous reviewers.

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Footnote

† Electronic supplementary information (ESI) available: Methods for preparation and characterization of the ZnO/CNC hybrid, dissolved Zn concentration, algal dry weight and chlorophyll a/b measurement, reactive oxygen species (ROS) assay, catalase (CAT) activity level, MDA content, electron microscopy observations, and SR-FTIR imaging are provided in Texts S1–S9. Additional figures and tables related to nanoparticle characterization, algal toxicity analysis, biochemical imaging, and elemental distribution are also provided. See DOI: 10.1039/d1en00881a

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