Jindi Zhao,Yongqiang Ruan,Zhe Zheng,Yunhan Li,Muhammad Sohail,Fanghui Hu,Jiahuan Ling,Lihui Zhang
Gold nanoparticles-anchored peptides enable precise colorimetric estimation of microplastics
Environmental toxicology,Hazard identification,Microenvironment
Summary Microplastics (MPs, particle size < 5 mm) are an emerging contaminant in aquatic environment, which have attracted increasing attention worldwide. In this study, a colorimetric method for MPs detection was developed based on gold nanoparticles (AuNPs)-anchored peptides (LCI or TA2), which are able to specifically recognize and adhere to polypropylene (PP) or polystyrene (PS). The AuNPs-anchored peptides accumulated on the surface of MPs, rendering a color change from red to gray-blue and transforming the surface plasmon absorption intensity and wavelength. The designed method presented high selectivity, stability, and reproducibility, with a detection range of 2.5–15 μg/mL. The results demonstrated that the developed approach will be valuable in the precise, facile, and cost-effective estimation of MPs in different matrices, regulating the control over MPs pollution and its hazardous impact on health and ecosystems.
Gold nanoparticles-anchored peptides enable precise colorimetric estimation of microplastics

Microplastics (MPs, particle size < 5 mm) are an emerging contaminant in aquatic environment, which have attracted increasing attention worldwide. In this study, a colorimetric method for MPs detection was developed based on gold nanoparticles (AuNPs)-anchored peptides (LCI or TA2), which are able to specifically recognize and adhere to polypropylene (PP) or polystyrene (PS). The AuNPs-anchored peptides accumulated on the surface of MPs, rendering a color change from red to gray-blue and transforming the surface plasmon absorption intensity and wavelength. The designed method presented high selectivity, stability, and reproducibility, with a detection range of 2.5–15 μg/mL. The results demonstrated that the developed approach will be valuable in the precise, facile, and cost-effective estimation of MPs in different matrices, regulating the control over MPs pollution and its hazardous impact on health and ecosystems.

The term "microplastics" (MPs) was first coined in 2004 to refer mainly to smaller pieces of debris,, but includes a much broader range at present. Polymers in the form of fibers, films, fragments, pellets, or particles with diameters ranging from 1 nm to 5 mm are considered MPs by the national oceanic and atmospheric administration., Even if the world embarked immediately on a globally coordinated effort to reduce plastic consumption, there would still be 710 million metric tons of plastic by 2040. Large plastic waste can be transformed into MPs by natural weathering, seawater erosion, ultraviolet irradiation, and biological effects. MPs, a type of persistent pollutant, are widely distributed in the environment and are difficult to degrade, with half-lives of hundreds of years. In the second session of the United Nations Environment Assembly in 2015, MPs pollution was included as the second major scientific problem in the field of environmental and ecological sciences. MPs were detected in many places, such as the Yellow Sea, the Three Gorges of the Yangtze River, and Taihu Lake. Kosuth et al. studied the content of MPs in tap water and estimated that the average annual intake of MPs in the human body was more than 5,800, of which about 88% came from tap water. Since MPs are widespread in the water environment and are able to cause serious harm to human health through the food chain, monitoring the content of MPs in water (drinking water, oceans, lakes, etc.) is crucial for environmental analysis and water safety. In order to detect the MPs in water, a series of detection methods have been developed. Conventional methods for the determination of MPs in water samples include pyrolysis gas chromatography, Fourier transform infrared spectroscopy, Raman spectroscopy, etc. Although these methods exhibit high selectivity and sensitivity, they are costly, labor-intensive, and time-consuming. Gold nanoparticles (AuNPs) are ideal nanomaterials for optical biosensing because of their high molar extinction coefficient, large specific surface area, and good biocompatibility, and their optical properties are related to shape, size, and distance distribution. AuNPs typically bind to biomolecules (such as nucleic acids, proteins, peptides, etc.) to form complexes with different properties. When these complexes are combined with the test objects, detection results with high sensitivity and accuracy can be obtained by using simple instruments, such as an ultraviolet (UV) spectrophotometer. The composite not only possesses the unique features of biomolecules but also owns the optical, electrical, magnetic and catalytic properties of AuNPs. The diversity of the polypeptide structure and its high biological activity make it more suitable to synthesize the composite with AuNPs for the analysis and detection of the target analytes. Parnsubsakul et al. designed the polypeptide EKEKEKPPPPC and constructed a nickel ion colorimetric sensor based on the EKEKEKPPPPC-AuNPs complex with the help of the high affinity between the polypeptide and nickel ions. Hiroshi and colleagues reported a lectin sensing strategy based on a polypeptide-gold nanoparticle carbohydrate complex. In recent years, polypeptide-AuNPs complexes have shown promising applications in the field of biosensing. However, it is rarely used in the detection of MPs. The anchored peptide liquid chromatography peak I (LCI) has β Folded structure, stable in an aqueous solution. Polypropylene (PP) surface is hydrophobic, and the interaction with peptide LCI is mainly controlled by hydrophobic interaction. Tachystatin A2 (TA2) was reported to be enriched in aromatic amino acids and the side group of the polystyrene (PS) macromolecular chain is the benzene ring. The binding mechanism of the aromatic amino acids of TA2 and the aromatic moieties of PS is primarily based upon the stacking of aromatic ring systems.,, This means that the anchoring peptides LCI and TA2 can bind to PP and PS, respectively. To circumvent the drawbacks of the available MPs detection approaches, such as a high detection limit, low sensitivity, high cost, labor-intensive, and time-consuming, we fabricated a colorimetric method for the estimation of MPs using AuNPs-anchored peptides (LCI or TA2) with specific recognition and adhesion to MPs. In principle, the AuNPs-anchored peptides accumulated on the surface of MPs, rendering a color change from red to gray-blue and transforming the surface plasmon resonance absorption intensity and wavelength. At present, AuNPs combined with peptides have not been studied in the detection of MPs. Therefore, our project aimed to explore the specific recognition and detection of MPs using AuNPs-anchored peptide, facilitating the commercial detection and control over MPs pollution.

The amino groups of the N-terminals of the anchored peptides LCI and TA2 (the sequences are shown in Table S1) were conjugated to the carboxyl group on the surface-modified AuNPs via an amide linkage. Theoretically, in an alkaline environment, AuNPs-LCI (pI = 5.43) and AuNPs-TA2 (pI = 5.13) are more stable than AuNPs because the anchored peptides are negatively charged and form a uniform layer, increasing the electrostatic repulsion between AuNPs. In addition, LC1 and TA2 can attach specifically to the surfaces of MPs, PP, and PS, through hydrophobic interactions, further inducing the aggregation of AuNPs, Scheme 1.

To justify the specific binding of anchored peptides and MPs, the EGFP was introduced into the anchored peptide structure and was combined with EGFP anchored peptide and interval sequence (10 alanine and tobacco etch virus [TEV] protease cleavage sites) to form a fusion protein, as shown in Figure S1A. The sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) verified the normal expression of the fusion protein, as shown in Figure S1B. Fluorescence microscope images showed that EGFP, as a positive control, did not affect the binding of anchored peptides and plastic. The LCI and TA2 specifically interacted with PP and PS, respectively, as shown in Figure 1.

The absorption peak of AuNPs is generally in the range of 500–600 nm, and the UV-PC1800 spectra showed that the absorption peak of AuNPs synthesized in this experiment was 535 nm. The stability of AuNPs was extremely important in the testing process; hence, AuNPs were placed at 4°C for one month and tested for stability. The results showed that the absorption peak of the AuNPs did not change significantly after being placed at 4°C for one month, as shown in Figure 2A, indicating that AuNPs had good stability and were suitable for the detection of MPs. In addition, the shape of the absorption peak was symmetrical, demonstrating that the AuNPs were of uniform particle size. Furthermore, TEM analysis was performed to verify the correlation between UV-PC1800 spectral analysis and AuNPs morphology. As shown in Figure 2A, AuNPs were spherical with a uniform size distribution and good dispersibility. In addition, as shown in Figure S2, the particle size distribution of AuNPs is mainly around 30 nm. Peptides have become a core material for the surface modification of nanomaterials due to their good biocompatibility and facile modifications. Peptides can be modified on the surface of materials in both covalent and non-covalent ways, and the modified materials can be used for the detection of various substances., In this study, amino groups were present in the anchored peptide (as shown in Figure 2B), and the modification process was realized by forming an amide bond between amino groups in the anchored peptide and carboxyl groups in the modified AuNPs. To modify AuNPs with anchored peptides, thiol-containing undecanoic acid was assembled on the AuNPs surface by AU-S bond to form a monolayer so that carboxyl groups were obtained on the surface of AuNPs. In this case, the anchored peptides were modified on the surface of AuNPs by covalent bonding, as shown in Figures 2C and 2D. It can be seen that the FTIR spectra of AuNPs modified with anchored peptides were consistent with those of pure anchored peptides. The characteristic bands of anchored peptides, such as the amide I band at 1600 cm−1, the amide II band at 1450 cm−1, and the amide III band at 1250 cm−1, were completely retained in the infrared spectra of AuNPs. Therefore, it can be inferred that the backbone and functional groups of the anchored peptide chain remained intact after binding of the anchored peptide to the AuNPs. Based on the FTIR results, we speculated that the anchored peptide binding and AuNPs remained bioactive. In addition, as shown in Figure S3, The zeta potential of AuNPs (−44 mV) has no significant difference from that of AuNPs-LCI (−40 mV) and AuNPs-TA2 (−42 mV), indicating that the LCI or TA2 modification has no significant effect on the stability of AuNPs.

To demonstrate the binding effect of AuNPs-anchored peptide and MPs, UV spectra of AuNPs, anchored peptide, MPs, AuNPs-anchored peptide, AuNPs-MPs, and AuNPs-anchored peptide-MPs were obtained, as shown in Figures 3A and 3B. The AuNPs presented an absorption peak at about 535 nm, while the AuNPs modified with anchoring peptides generated symmetrical absorption peaks at 570 nm and 567 nm. The results showed that the anchored peptides were successfully conjugated on the surface of AuNPs, and the AuNPs-anchored peptide complex offered good dispersibility in solution. The presence of MPs in the reaction matrix decreased the value of the absorption peak of the AuNPs-anchored peptide system to 0.42 and 0.49. There was no obvious absorption peak in the UV absorption spectra of the anchored peptide, MPs, and anchored peptide-MPs, and the absorption peak of AuNPs-MPs was consistent with that of AuNPs. In addition, the absorption peaks of AuNPs-LCI-PP + PS and AuNPs-LCI-PP, AuNPs-TA2-PP+PS, and AuNPs-TA2-PS did not show obvious changes, as shown in Figure S4. In conclusion, the MPs interacted with AuNPs-anchored peptide molecules and changed the absorption traits of the AuNPs-anchored peptide system, enabling the detection of MPs. Moreover, transmission electron microscope (TEM) analysis was performed to further elaborate the interaction between the AuNPs-anchored peptide and MPs. The TEM images in Figures 3C and 3D showed that the AuNPs-anchored peptide had good dispersibility. Besides, AuNPs-anchored peptide molecules aggregated in the presence of MPs, as shown in Figures 3E and 3F.

The maximum output of a reaction system is directly related to the principle of parameter optimization; hence, various parameters, including the anchored peptide concentration, reaction pH, and mixing time, were optimized. The size of the AuNPs and the concentration of the anchored peptides were optimized, and the ratios of A750/A570 and A800/A567 reached maximum when the average size of AuNPs was 30 nm and the anchored peptide concentration was 2.5 mg/mL in the presence of MPs, as shown in Figure 4. In addition, the pH value of the solution also played a critical role. When the pH of the solution was below the pI of the anchored peptide, the anchored peptide had a positive charge and caused the aggregation of AuNPs. However, in an alkaline environment, the negatively charged anchored peptide formed a uniform charged layer, which increased the electrostatic repulsion between AuNPs. It can be seen from Figures 4C and 4G that the most appropriate pH was 7.4. Under the above-mentioned optimal conditions, the influence of reaction time on the system was also studied. It can be observed from Figures 4D and 4H that the values of A750/A570 and A800/A567 increased gradually with time and stabilized after 15 min, indicating that the reaction had been at equilibrium state.

We prepared 2.5–15 μg/mL PP and PS suspensions in water. The color of the AuNPs gradually changed from red to purple and finally to blue-gray as the MPs concentration was increased, Figures S5A and S5B, and the intensities of absorption peaks at 570 nm and 567 nm decreased gradually, Figures 5A and 5B. These results indicated that the aggregation of AuNPs-anchored peptides depended on the concentration of MPs. We used the ratio of absorptions (A750/A570 and A800/A567) to express the degree of aggregation as mentioned above. Figure 5C showed that the concentration of PP relied on the colorimetric response (A750/A570 values) of the assay. The A750/A570 values increased linearly with the PP concentration in the range of 2.5–15 μg/mL, and the linear regression correlation coefficient was 0.9918. Figure 5D showed that the response of the A800/A567 assay was linear, with a linear regression correlation coefficient of 0.9931, at the PS concentration range of 2.5–15 μg/mL. The values of regression correlation coefficients were close to 1, indicating that our new method was sensitive to the detection of MPs.

Other MPs may interfere with the detection of PP and PS, so we selected some common MPs, such as PS, PVC, PS, PET, PE, and PP. Under optimized conditions, the AuNPs-LCI system presented a much higher value of A750/A570 in the presence of PP as compared to the other MPs, Figure 6A. In addition, the A800/A567 ratio of MPs in the presence of PS in the AuNPs-TA2 system was much higher than that of other MPs, as shown in Figure 6B. Besides, the colorimetric method demonstrated significant selectivity for PP and PS, as shown in Figures S5C and S5D. The above results indicated that the two detection systems had good selectivity for the monitoring of PP and PS.

To validate the accuracy and sensitivity of this method, we applied the fabricated method to the real environmental samples. MPs may exist in some environmental samples, such as lake water or tap water, so the samples were collected from the lake water of Nanjing Normal University. The samples were tested using the traditional Raman method, and no MPs were detected, as shown in Figure S6. Therefore, we added different concentrations of PP and PS (3.75, 7.5, and 12.5 μg/mL) to the lake water. The recovery results of MPs in the spiked lake water samples were shown in Table 1 and Table 2. The PP recovery of this method was within the range of 92.50–99.47%, and the relative standard deviation was between 1.52 and 2.07%. The PS recovery of this method was within the range of 96.57–110%, and the relative standard deviation was between 1.52 and 2.39%. The results showed that the designed colorimetric method could be used to detect MPs in real water samples. In addition, Table S2 gives several commonly used methods for the detection of microplastics at present. It can be seen that this experimental method has the advantages of time saving, labor-saving, and low detection limit. Therefore, it is worthwhile to develop anchoring peptides that specifically recognize other MPs in the future.

In conclusion, we presented a rapid and simple method for the detection of MPs using MPs as aggregation promoters for AuNPs-anchored peptides. To the best of our knowledge, there are few reports on the detection of MPs using the interaction between anchored peptides and MPs. The detection results were highly linear, and the reaction was completed within 15 min. Besides, anchored peptides (LCI or TA2) rendered the detection system specific for the precise detection of MPs. Briefly, the results demonstrated that the developed approach will be valuable in the precise, facile, and cost-effective estimation of MPs in different matrices, regulating the control over MP pollution and its hazardous impact on health and ecosystems.

This proposed methodology has still many limitations to apply for other plastics, various sizes, and media types. Nevertheless, we hope that this suggestion will give rise to a new measurement method that can be used to measure various sizes and types of microplastics in the future.



Further information and requests for resources should be directed to and will be fulfilled by the lead contact, Prof. Lihui Zhang (zhanglihui@njnu.edu.cn).

This study did not generate new unique reagents. All chemicals were obtained from commercial resources and used as received.

Firstly, 20 mL of 0.01% chloroauric acid was added to a 50-mL round-bottom flask with a magneton, fitted with a spherical condensing tube, and heated to boiling. After that, 2 mL of the 1% sodium citrate solution was added at once to the boiling mixture, and vigorous stirring was continued during reflux for 10 min. Heat source was turned off mixture was stirred for another 15 min and cooled to room temperature. The suspension of AuNPs (0.05 mg/mL, 25 mL) was mixed with the solution of 11-mercaptoundecanoic acid (10 mM, 25 mL) in a 50-mL centrifuge tube and placed at 4°C for 12 h. Then, the suspension was centrifuged at 8000 rpm for 1 h, and the supernatant, which contained unreacted 11-mercaptoundecanoic acid, was removed to get the surface-functionalized AuNPs.

The plasmid pET28a (+): anchoring peptide was synthesized as the vector skeleton of the fusion protein. The vector backbone pET28a (+): anchoring peptide was amplified using F-vector and R-vector, and the primers F-EGFP and R-EGFP were used to amplify the EGFP fragment. Linearized vectors and inserted fragments were cloned and converted to DH5α by thermal shock. The positive clones were screened by PCR and sequenced by Nanjing Tsingke. The plasmids extracted from the positive clones and verified by sequencing were transformed into E. coli BL21 (DE3). Single colonies were selected and cultured in Luria-Bertani (LB) media containing Kana. Isopropyl β-D-Thiogalactoside (IPTG) with a final concentration of 0.1 mM was added to induce protein overexpression at 20°C. Cells were disintegrated by ultrasound and the supernatant was collected. The EGFP-anchored peptide fusion protein was purified by His gravity column (Takara, Beijing, China), dialysis, and ultrafiltration. The concentration of the protein was determined by the BCA kit and UV-Vis spectroscopy, and the expression of the recombinant protein was detected by SDS-PAGE using 5% concentrated glue and 12% separated glue.

The PP fibers and PS particles were added to 2 mL solution of EGFP-fusion protein and EGFP protein, and they were allowed to react for 10 min at room temperature. Then the suspension was washed thrice with 10 mL portions of Tris/HCl buffer. Finally, they were observed under a fluorescence microscope (Olympus, Beijing, China).

The EGFP-fusion protein was cleaved by the tobacco etch virus (TEV) protease, which specifically recognized the amino acid sequences of Glu-Asn-Leu-Tyr-Phe-Gln-Gly and precisely cleaved the fusion protein into EGFP and the anchor peptide. Then the lysate was transferred to His gravity column (Takara, Beijing, China), and the anchored peptide was collected in circulation. The anchored peptide was dissolved in PB solution (pH 7.4) to a concentration of 10 mM for further use. The 11-mercaptoundecanoic acid-modified AuNPs (3 mL) were mixed with 50 μL of anchor peptide and 50 μL of EDC-NHS mixture and allowed to react at 4°C for 24 h. The unreacted anchor peptide was removed by centrifugation (9500 rpm, 25 min) and washing thrice. AuNPs and AuNPs-anchored peptides were analyzed by Fourier transform infrared spectroscopy (FTIR) (Vertex 70, Bruck, Germany). The absorbance in the mid-infrared region of 4000-400 cm−1 was measured with a scanning resolution of 4 cm-1. The background absorbance was measured and subtracted from the tested samples. The surface zeta potential of AuNPs and AuNPs-anchored peptides in solution was tested using a dynamic light scattering particle size distribution instrument (Zeta PALS, BIC, America).

PP/PS solutions with different concentrations (2.5-15 μg/mL) were prepared. The AuNPs-anchored peptide complex (3 mL) was evenly mixed with MPs solutions of different gradient quantities and incubated at 4°C for 15 min. After incubation, color changes were observed, and the UV-Vis spectra of the reaction mixture were recorded. In the AuNPs-LCI system, the absorption values at 570 nm and 750 nm corresponded to the number of dispersed and aggregated AuNPs, respectively. Therefore, the relationship of dispersed to aggregated AuNPs could be described by the ratio of the absorption value at 750 nm to the absorption value at 570 nm (A750/A570). In the AuNPs-TA2 system, the absorption values at 567 nm and 800 nm belonged to the number of dispersed and aggregated AuNPs, respectively. Therefore, the ratio of dispersed to aggregated AuNPs could be represented by the ratio of absorption values at 800 nm to those at 567 nm (A800/A567), respectively.

The same method was used in the selectivity experiments. We prepared suspensions of common MPs (such as Polyethylene (PE), Polyvinylchloride (PVC), Polyethylene terephthalate (PET), PS and PP) with equivalent concentrations and added them to the solution of AuNPs-anchored peptide. The reaction solution was characterized by the UV-Vis spectrophotometer (1800PC, Shanghai, China). The recovery experiments were accomplished using MPs-spiked Lake water. Compared with the standard curve of MPs, we confirmed the PP and PS concentration in the samples by using the responses (A750/A570 and A800/A567) of the colorimetric assay against MPs-spiked water samples. Raman's method was used for the analysis of lake water without MPs-spiking. A portable Raman spectrometer (Prosp-Micro-S1, Hangzhou, China) with 785 nm excitation light was used to collect SERS signals. The laser power was about 105 mW and the exposure time for each SERS amount was 3 s.

All the experiments were performed in triplicates with the results presented as a mean value with standard deviation (Mean ± SD). The data were analyzed using analysis of variance, and the significant difference between the means was compared using t-test at a significance level at p < 0.050. Statistical analysis was done using Statistical Processor System Support (SPSS) Statistics 27.0 software.