Real-time detection of label-free submicron-sized plastics using flow-channeled differential interference contrast microscopy

doi:10.21203/rs.3.rs-2610338/v1

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Real-time detection of label-free submicron-sized plastics using flow-channeled differential interference contrast microscopy

https://doi.org/10.21203/rs.3.rs-2610338/v1

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Owing to the surge in plastic waste generated during the COVID-19 pandemic, concern regarding microplastic pollution in aqueous environments is increasing. Since sub-micron plastics can accumulate in the human body, their real-time detection in water is necessary. However, the decrease in the scattering cross-section of small polystyrene (PS) particles in aqueous media precludes detection by bright-field microscopy. To address this problem, we propose and demonstrate a differential interference contrast (DIC) system that incorporates a magnification-enhancing system to detect PS beads in aqueous samples. To detect micron-scale and sub-micron-scale PS beads in the stationary and mobile phases, a microfluidic chip was designed considering the imaging depth of focus and flow resistance. PS beads of various sizes flowing in deionized and tap water at varying speeds were quantitatively observed under Static and Flow conditions. The real-time detection and quantification of PS beads of size ≥ 200 nm at a constant flow rate in water was successful. It was also possible to classify two different sizes, 2 µm and 750 nm, in a mixed solution. Thus, the proposed novel method can significantly reduce the analysis time and improve the size-detection limit. The proposed DIC microscopy system can be coupled with Raman or IR spectroscopy in future study for chemical composition analysis.

Earth and environmental sciences/Environmental sciences

Earth and environmental sciences/Environmental sciences/Environmental chemistry/Environmental monitoring

The increasing amount of plastic waste in the aqueous environment has led to increasing accumulation of microplastics (MPs, < 1 µm) in the human body ¹. The COVID-19 pandemic has increased the quantity of plastic waste, owing to the use of personal protective equipment, disposable food packaging, cutlery products, etc. According to the World Health Organization, 89 million facemasks and 76 million gloves were consumed every month during the COVID-19 pandemic ². These changes have caused an enormous increase in plastic waste discharge into the aquatic environment. Therefore, there have been many reports on the role of decomposed MPs as carriers of pollutants ³ and the spread to other organisms through the food chain ^4,5. For example, a recent study verified that nanoscale MPs could be sequentially transferred to higher trophic levels in marine organisms, potentially leading to human exposure via the food chain ⁶.

Since MPs have been detected in drinking water ^7,8, concerns are increasing because of their potential accumulation in the human body. It has been stated that MPs smaller than 1.5 µm are likely to pass through biological membranes ^9,10. Leslie et al. ¹ reported that MPs smaller than 0.7 µm were detected in the blood of 17 out of 22 adults. It was demonstrated that MPs smaller than 1 µm are found in the blood because they can penetrate through the cell membrane, thereby traveling throughout the body. One of the main pathways by which the human body is exposed to MPs is ingestion ^10,11. Drinking water is the main source of MPs in the human body. Because large MPs can be broken down into smaller sizes by physical and chemical reactions, such as weathering and sunlight exposure, it is important to develop a real-time analysis method that can detect sub-micron and nanoscale MPs in water samples before ingestion.

In general, the common process flow of MP detection and analysis in environmental samples follows four major steps: extraction, separation, identification, and quantification ¹². Prior to analysis, the preparation of dispersion and desiccation of particles on substrates, e.g, glass, are usually employed as pretreatment methods. Raman spectroscopy and Fourier-transform infrared (FT-IR) spectroscopy are the most widely used techniques for instrumental analysis. However, these analysis methods have two common limitations: expensive instruments and high labor and time costs. Long detection time and image resolution are other bottlenecks for measuring small MPs, which have a high impact on human health.

Recently, to address the limitations of the detectable size mentioned above, there have been a few attempts at the nanoscale level, including the application of scanning probe techniques, such as atomic force microscopy-based infrared spectroscopy (AFM-IR) ^13,14. The AFM-IR technique has been successfully used to obtain images and IR spectra of 100-nm polystyrene beads (PS beads) ¹⁵. However, it cannot be used for precise measurement when the surface roughness is non-uniform. In general, samples with height variation of 4–6 µm are possible to analyze, depending on the specific topography ¹⁶. AFM-IR, owing to the detection mechanism depending on the mechanical movement of a cantilever, is also a time-consuming method because only a small area of the sample with a limited number of particles can be analyzed locally through scanning. Thus, to shorten the analysis time, a new technique for detecting the flowing plastic in aqueous media is required instead of time-consuming scanning. Gillibert et al. ¹⁷ proposed a new technology combining the use of an optical tweezer with Raman analysis and succeeded in detecting MPs of varying sizes (ranging from 20 µm to 50 nm) using an optical force in aqueous media. However, Raman spectroscopic detection of MPs collected in a microchamber by tweezing requires several tens of seconds, which is not suitable for high-speed imaging.

Differential interference contrast (DIC) microscopy can be a powerful tool for observing any existing particles, including MPs, in flowing water at an early stage without complicated sample pretreatment, because imaging can be performed in real time. If any sample can be filtered while it is detected by DIC microscopy, the total volume that needs monitoring can be dramatically reduced. DIC microscopy allows easy discrimination of the three-dimensional morphology of MPs, regardless of their transparency in aqueous media ¹⁸. Because the transmittance of PS MPs in water is approximately 90–95% in visible light, monitoring PS MPs in a flowing fluid under a bright-field (BF) microscope is extremely difficult ¹⁹. In addition, Buzzaccaro et al. ²⁰ applied the term ‘ghost particles’ to 50–200-nm PS particles in aqueous media while using a velocimetry technique, because their speckle pattern and not their shape is observed under the BF microscope. Owing to the advantage that it can be used to detect transparent materials, DIC has been effectively utilized to confirm MP-cell interactions. Recently, Ramsperger et al. ²¹ successfully distinguished internalized PS MPs in murine macrophages with a diameter of 3 µm.

A novel flow-channeled DIC system was proposed herein for real-time monitoring PS beads. In particular, a microfluidic chip was fabricated considering the imaging field of view and depth of focus of the DIC system, while a high-precision pressure pump was utilized to deliver PS beads with sizes ranging from 2 µm to 200 nm in water suspension. The use of a high-magnification DIC microscope was optimized by controlling the DIC components and introducing an additional optic system for the observation of the sub-micron-sized PS beads (up to 200 nm). To validate the resolution of the proposed system and the reliability of its quantitative data obtained by the DIC system, scanning electron microscopy (SEM) and Raman spectroscopy were also performed and the results were compared. In addition, this real-time image-based system was experimentally investigated using PS bead-spiked deionized (DI) and tap water samples without pretreatment. Finally, DIC images captured by real-time video recording using a charge-coupled device (CCD) camera were analyzed to determine the size, morphology, and number of PS beads in the aqueous suspension.

Imaging the PS beads using high-magnification DIC microscope

We developed a self-designed reflective DIC microscope, as shown in Fig. 1a. The degree of optical path difference between ordinary and extraordinary beams in the DIC prism causes the amplitude in the DIC image to appear darker (or brighter) ^22,23. The bias retardation between the two beams can be adjusted by shifting the DIC prism or introducing a de Sénarmont compensator, which causes amplitude changes based on the degree of the bias retardation ²⁴. To simplify the DIC microscope, it was optimized by moving the DIC prism back and forth rather than introducing a de Sénarmont compensator. Supplementary Fig. 2a shows images of PS beads of different sizes according to the bias retardation values applied by DIC prism shifting. The smaller the size and the more sensitive are the changes in the appearance of PS beads according to the value. The contrast of the PS beads was maximized when bias retardation between one-twelfth and one-twentieth wavelength was introduced. In addition, the contrast was increased owing to the minimized chromatic aberration by inserting the bandpass filter (Supplementary Fig. 2b) ²⁵. As a result, the DIC image was optimized, as shown in Fig. 2. We compared the images obtained by DIC with SEM and confirmed that all PS beads, including 200-nm-sized PS beads, were clearly observed. Although Hamilton et al. ²⁶ reported that there was an intensity difference in image contrast due to geometric differences derived from size, similar shapes were clearly observed. Furthermore, the PS beads were observed as a three-dimensional sphere by DIC microscopy, but as a plane by SEM, because the brightness of the sample for the DIC microscope varied along the shear axis (indicated by the double-headed arrow in Fig. 2) caused by the image-forming mechanism of DIC microscopy.

Cross-validation and confirmation: comparison of DIC microscope and Raman spectroscopy

To confirm the accuracy of the quantitative evaluation, DIC microscopic images were compared with Raman mapped images, as shown in Fig. 3a,b. Raman spectroscopy is a chemical method most commonly used to identify PS beads smaller than 20 µm because of its high spatial resolution (< 1 µm) and lack of spectroscopic interference from water ^27–29. Raman images of 30 particles in dried PS beads droplets were obtained in a drilled polydimethylsiloxane (PDMS) mold laid over a cover glass. It appears that most of the particles in the DIC image are matched with the particles in the Raman image in the figure.

When PS beads were distributed in the center, both DIC and Raman images showed spherical beads, as shown in the left column of Fig. 3c. In this case, the particles were placed exactly at the depth of focus of the Raman excitation laser. However, the PS beads on the rim of the mold were located at higher positions because of the attractive force to the PDMS wall. As a result, the peak intensity was weaker in the Raman image. The right column in Fig. 3c shows a weaker Raman image of the PS beads alone or in clusters at the rim. Obviously, the Raman image of the beads faded around the rim of the PDMS mold because of positional deviation from the focus. It has been reported that the causes of the Raman signal intensity variation at 1000 cm^− 1 are heavily dependent on focal differences, imaging locations, and lumping conditions ^27,30. The difference in the Raman intensity was remarkable for each position, as shown in Fig. 3d.

Because the experiments were repeated five times, the DIC microscope could present an average of 28 beads, while 23 beads were counted by Raman spectroscopy (Fig. 3e), presumably because of the weak signals at the rim. We took the same Raman image five times and the measurement for an area of 1.5 mm × 1.5 mm took less than 15 min with DIC microscopy while it took 53 h 15 min with Raman microscopy.

Flow-channeled DIC system: Cross-validation and confirmation on PS bead detection in DI water

Figure 1b and Supplementary Fig. 1 show the setup of the flow-channeled DIC system in flowing water and a photograph of the flow-channeled DIC system, respectively. To investigate whether real-time quantification of the PS beads flowing through the microfluidic channel is possible, an experimental verification was conducted by comparing the Static and Flow conditions. PS beads (2-µm, 750-nm, 200-nm) were dispersed to 1.2 × 10⁶ particles/mL. The control experiment, wherein a microchannel with the dimensions of 50 mm × 50 µm × 10 µm (total volume of 0.025 µL) and no flow rate was scanned using the DIC microscope, was referred to as the Static condition. Under the Static condition, DIC imaging covering the field of view (FOV; 338 µm × 225 µm) was performed 200 times by traveling over the entire channel with a length of 50 mm. In the experimental group, referred to as the Flow condition, the fluids containing the same concentration of PS beads as the Static conditions were quantified by DIC imaging at the center point of the microchannel with a constant flow rate of 0.001 µL/min.

Figure 4 shows three types of PS beads with diameters of 2-µm, 750-nm, and 200-nm in the microfluidic channel, obtained under (a) Static and (b) Flow conditions, imaged under the flow-channeled DIC microscope. Three-times-enlarged PS bead images are shown in the corner. Since the DIC image is produced by an optical path difference, it can be noted that the DIC image of PS beads in aqueous media (Fig. 4) does not differ significantly from that of the dried PS beads (Fig. 2). Each PS bead image appears to have the same contrast style as the DIC image of the 20-µm PS microsphere floating in water taken by Cui et al. ³¹. In addition, 200-nm PS beads corresponding to the theoretical spatial resolution of the optical microscope were also prominently detected.

As Braslavsky et al. ³² mentioned, the observation of small-sized PS in aqueous media was difficult due to less cross-section scattering under the BF microscope, not because of its spatial resolution. Therefore, the PS beads observed under the BF microscope were blurred and indistinguishable (Supplementary Fig. 3). The BF image was captured in the same region and at the same magnification without DIC components (polarizer, analyzer, and DIC prism). This demonstrates that the DIC microscope is a more suitable method to observe PS beads in aqueous media, such as our microfluidic system, than the BF microscope.

Figure 4b shows the DIC image extracted from the DIC image recorded at a fixed microfluidic channel position using a video-editing program (VLC media player, VideoLAN, France). While imaging the PS beads under flow conditions, the exposure time of the CCD camera was maintained below 80 ms (15 fps) to minimize motion blurring ³³. This allowed the clear observation of PS beads flowing at velocities between 0.9 and 1.8 mm/min, including beads as small as 200-nm for Flow condition. As Culver et al. ³⁴ mentioned, the DIC image contrast of a sphere is formed independently of the direction because it interacts equally with both ordinary and extraordinary rays, and the PS beads were observed with the same contrast while flowing in the microfluidic channel.

Quantitative evaluations of detected PS beads in DI water

To confirm the reproducibility and accuracy of the established analytical method, quantitative evaluations were performed under both Static and Flow conditions in DI water. All experiments were repeated three times. It is expected that an average of 30 particles must be observed in the Static condition when the microfluidic channel (50 µm × 50 mm × 10 µm) is filled with 0.025 µL of a suspension. In the Static condition, the mean and standard deviation values of the 2-µm, 750-nm, and 200-nm size counts were 30 ± 0 particles/0.025 µL.

To evaluate performance in the Flow condition, DIC microscopy was used to observe the 0.001-µL/min flow of PS beads in water for 25 min, which would produce the same number of PS beads as encountered under the Static condition. After comparing the Static and Flow conditions for each size group (2-µm, 750-nm, and 200-nm), the means and standard deviation values of the particle counts, which were repeated three times, are shown in Table 1. For the Flow condition, the mean and standard error values of the 2-µm and 200-nm size counts were 30 ± 0 particles/0.025 µL, and the 750-nm size count showed a value of 29 ± 1 particles/0.025 µL. Given that the number of beads counted in the Static and Flow states was similar, it was confirmed that the detection accuracy was high not only for the PS beads in the stopped state (Static condition), but also in the motion state (Flow condition).

Detection of mixture of sizes of PS beads in DI water

Real-time detection videos for the Flow condition over a wide FOV (338 µm × 225 µm) were streamed at 25× speed in Supplementary Movie 1. The flowing PS beads were imaged with sufficient contrast to enable detection and measurement of size, and the bright intensity of the PS beads image was higher for the 2-µm beads than for the 750-nm beads. With this feature, it was possible to distinguish between two different sizes of PS beads in the mixture. The 2-µm and 750-nm PS beads are indicated with blue and orange squares, respectively (Fig. 4).

By scanning the microchannel and counting stationary PS beads under Static conditions, 30 particles were detected in all three experiments in a mixed solution of 2-µm and 750-nm beads. In the Flow condition for the same solution condition, as shown in Table 2, the detection rate of the 2-µm beads was 30 ± 1 particles/0.025 µL for 25 min, while that of the 750-nm PS beads was 30 ± 2 particles/0.025 µL of mean count (+ standard deviation).

Detection of PS beads in tap water

Figure 5 depicts the detection of PS beads in drinking tap water using a flow-channeled DIC system. The 2-µm, 750-nm, and 200-nm PS beads were dispersed to 1.2 × 10⁶ particles/mL and introduced into the microfluidic chip by the pressure pump at a constant flow rate of 0.001 µL/min, which is the same condition as DI water. As shown in Fig. 5, the PS beads in tap water under both (a) Static and (b) Flow conditions were clearly observable as a spherical shape without any pretreatment. In tap water, each of the single sizes of the 2-µm, 750-nm, and 200-nm PS beads were repeatedly quantified three times. When comparing the Static and Flow conditions for each size group, the standard deviations were 2 particles/0.025 µL, 3 particles/0.025 µL, and 3 particles/0.025 µL for the 2-µm, 750-nm, and 200-nm PS beads, respectively (Table 3, Supplementary Fig. 5). To determine whether the microfluidic chip is suitable for tap water, we compared the movement tendency of the PS beads in DI (Supplementary Fig. 4a) and tap water (Supplementary Fig. 4b) in the microfluidic channel. The detection time of each PS bead flowing at a flow rate of 0.001 µL/min was examined thrice for 25 min. The detection time of each PS bead and its average time are represented by a colored box and a circle in Fig. S4, respectively. The relatively large box indicates a slow passage of the PS beads because they moved at the microfluid channel boundary. The red and blue lines represent linear regressions of the average detection time of PS beads under flow conditions that were repeated three times. The R-square (> 0.99) of the slope indicates that each PS bead was observed at a relatively uniform interval of every 38–48 s in the microchannel, regardless of the bead size or water type. Collectively, the flow-channeled DIC system used in this study was evaluated to be suitable for real-time quantification of beads ranging from 200 nm to 2 µm in both DI and tap water.

Evaluation of the flow-channeled DIC system in natural water, and environmental significance

Overall, in this study, PS beads (≥ 200 nm) flowing at a constant flow rate in aqueous phase were successfully detected and quantified in real time by DIC microscopy. More importantly, this system was also successful in detecting PS beads flowing in water up to submicron-sized PS beads without any sample labeling such as fluorescent dyeing. As shown in Fig. 6, the flow-channeled DIC system can detect spiked PS beads, even in pond water. Collectively, our findings highlight new and efficient MP quantification approaches with sizes ranging from 200 nm to 2 µm, which could significantly reduce the scanning time required compared with other spectroscopic methods for MPs detection. However, flow-channeled DIC systems do not provide a complete solution for the detection and characterization of MPs. Further research is needed to ensure that the flow-channeled DIC system can detect MPs in real-water samples containing various small particles, such as microorganisms, organics, and inorganics. For example, sample pretreatment, such as concentration, extraction, and filtration, can be combined with the DIC system to deal with a large volume of water samples ³⁵. In addition, the integration of the flow-channeled DIC system with spectroscopic methods, such as Raman or absorption spectroscopy, can improve the total detection and characterization of MPs ³⁶.

Microplastics and preparation

Standard PS beads (2.5% w/v aqueous suspension, 10 mL, Polysciences Inc, Warrington, USA) with four primary nominal sizes of 200-nm, 750-nm, 2-µm, and 10-µm were used without any surface modifications. For SEM and Raman analyses, PS beads were dispersed in deionized water (MilliQ; ≥18 MΩ) and then sonicated for 30 min for monodispersion. Subsequently, a droplet (3 × 10⁴ particles/mL) was spread onto a glass slide and dried for 24 h at room temperature. For image mapping comparison in large areas between the Raman and DIC microscopes, a PDMS (Silgard 184, Dow Chemical, Midland, MI, USA)-based substrate (thickness: 1 mm) with a 1.5-mm-diameter circular hole was attached to the slide glass, and a 10-µm PS bead droplet was dropped. To test the constructed flow-channeled DIC system, the stock PS bead suspensions (200-nm, 750-nm, and 2-µm) were diluted to 1.2 × 10⁶ particles/mL in the reservoir. After dilution, sonication was performed for 30 min prior to application. The water matrices used in the flow-channeled DIC system were DI, tap, and pond water. Tap and pond water were sampled at the Korea Institute of Science and Technology (KIST, Republic of Korea). The pond water sample was filtered through Whatman 42 paper (diameter 42.5 mm, pore size 2.5 µm) using a vacuum pump.

High-magnification DIC microscope

A 250-W halogen lamp (Mighty Light Plus, SPECTROLIGHT Inc., USA) was used, and the light intensity incident on the sample was 0.3 mW. The transparent specimen was visualized using three optical components: a polarizer, DIC prism, and analyzer. The polarizer and analyzer in our system were installed in a direction that minimized the light loss in the sample and optics, including beam splitters. The polarizer and analyzer were set to perpendicular polarizations of 0° (east–west) and 90° (north–south), which corresponded to the S-pole and P-pole of the reflection plane, respectively. The DIC prism (D1, Leica-Microsystems, Germany) with a shear axis of 45° was located in the back focal plane. The DIC prism is a birefringent material that splits a linearly polarized beam into ordinary and extraordinary beams. Because the two beams oscillate orthogonally, they travel separately until they are recombined in the DIC prism after being reflected by the sample. The optical image was produced by the interference of two beams with different phase shifts. The optical path difference between the two beams, produced by the spatial difference in the sample, was visualized as an amplitude in the DIC microscopy. The DIC prism was adjusted for contrast optimization of the PS beads in aqueous media. In addition, an achromatic lens and detachable bandpass filter (525/15 nm) were used to improve the spatial resolution by minimizing chromatic aberration ²⁵. To observe and count the PS beads at high magnification, we used an objective lens (20×, NA 0.5, Leica HC PL FLUOTAR) and placed a pair of achromatic lenses (magnification-enhancing system) between the analyzer and tube lens. We acquired DIC images using a CCD camera located at an image plane (Lt1265R, Lumenera Inc, Kanada).

Flow-channeled DIC system

The setup consisted of an air compressor, a pressure controller (OB1 Base MkIII, Elveflow, Paris, France), a computer monitor, liquid reservoirs, a microfluidic chip for the transportation of PS beads, and a DIC microscope. The pressurized air in the air compressor was controlled by a pressure controller, and the PS beads in the fluid were transferred through the microfluidic chip. To distribute the PS beads evenly, they were stirred with a magnetic bar and stirrer in a reservoir filled with the PS bead suspension. The PS beads transported through the microfluidic chip were detected by the DIC microscopy system in real time and flowed into the reservoir. A Teflon (FEP) tube was inserted and secured into the 2-mm-diameter circular inlet of the microfluidic chip to apply air pressure to the fluid, enabling the flow of PS beads through the chip channel, and another with the same diameter of 2 mm to the outlet to the reservoir. The material of the input and output tubes through which the PS bead suspension flows was selected as the FEP with minimal PS beads attached ³⁷.

Fabrication of microfluidic chip

The microfluidic chips were manufactured using a normal soft-lithography process. An SU-8 photoresist (Amed Co., Seoul, Korea) was spin-coated on a 4-inch silicon wafer, and a straight-linear rectangular channel was patterned by dry etching to fabricate a mold. The PDMS base and curing agent were mixed in a ratio of 10:1 for 3 min and then placed in the vacuum desiccator for 1 h to remove air bubbles. The PDMS mixture was poured into the mold and then placed again in a desiccator for 1 h to remove air bubbles. After baking in an oven at 80°C for 2 h, the PDMS was peeled off from the silicon wafer. A thin cover glass substrate and PDMS slab were adhered to the oxygen plasma (Covance Plasma System; Femto Science, Korea). The plasma-treated PDMS slab and the cover glass substrate were combined and placed in an oven at 100°C for 5 min. According to the Hagen–Poiseuille law, the geometry of the channel is one of the factors that directly affect the pressure within the microchannel ³⁸. As a result, the channel height is optimized between the DIC microscope's depth of field (DOF) and a high resistance value for the pressure to control the fluid flow. The PDMS microfluidic channel was 50 µm in width, 50 mm in length, and 10 µm in height, with a total volume of 0.025 µL.

SEM and micro-Raman spectroscopy

To confirm the imaging resolution of the DIC and evaluate the morphology of the PS beads, the different sizes (200-nm, 750-nm, 2-µm, and 10-µm) were imaged using a DIC microscope and compared with ultrahigh-resolution scanning electron microscopy (Regulus8230, Hitachi, Japan; magnification, 3–4 K). Samples were dried for 24 h and coated with a 4-nm-thick platinum layer (E-1045 Ion sputter, Hitachi, Japan).

Raman images of the plastic samples were obtained using a confocal Raman system (inVia Raman microscope, Renishaw, UK). During our experiment, the second harmonic of 532 nm from the Nd:YAG laser was irradiated for Raman excitation while the power was maintained at 6 mW, and the Raman signal was collected via a 20× objective lens. A whole Raman image covering an area of 1.5 mm × 1.5 mm was a mosaic image of 5 × 3 domains, with a single domain corresponding to an area of 436.7 µm × 272.5 µm. Each domain was mapped by raster scanning at intervals of 1 µm, and the images were visualized as a function of the intensity of the ring-breathing mode of PS located at approximately 1000 cm^{-1 39 40}. At each point, full Raman spectra were observed from 100 cm^-1 to 4000 cm^-1 for 3 s and the data acquisition time for a domain was 3 h 33 min. During the measurement with the DIC microscope, the sample with randomly displaced PS beads was moved in a spatial interval of 150 µm along the x- and y-axes.

ACKNOWKEDGMENTS This study was supported by the Institutional Program of the Korea Institute of Science and Technology (KIST) [grant number 2E32442].

AUTHOR CONTRIBUTIONS The manuscript was written with the contributions of all authors. All authors have read and agreed to the published version of the manuscript. Each author’s contributions are as follows: Jiyun Han: Conceptualization, Methodology, Data analysis, Writing- Original draft. Subeen Park: Conceptualization, Methodology, Data analysis, Writing- Original draft. Mingizem Gashaw Seid: Data curation, Validation, Reviewing. Byeongho Park: Data curation, Validation, Reviewing. Soo Hyun Lee: Data curation, Validation. Hyung Min Kim: Data curation, Writing- Reviewing and Editing. Jaesang Lee: Data curation, Validation, Reviewing. Jae Hun Kim: Supervision, Funding acquisition, Writing- Reviewing and Editing. Seok Won Hong: Supervision, Resources, Funding acquisition, Writing- Reviewing and Editing.

COMPETING INTERESTS The authors declare no competing interests. DATA AVAILABITY Data generated and analyzed during this study are included in the published articles and supplementary materials.

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Table 1 Particle count of 2-μm-, 750-nm-, and 200-nm-sized PS beads in DI water using flow-channeled DIC system.

Particle count (particles/microchannel volume, 0.025 μL)
	Number of 2-μm PS beads		Number of 750-nm PS beads		Number of 200-nm PS beads
	Static condition	Flow condition	Static condition	Flow condition	Static condition	Flow condition
Trial 1	30	30	30	30	30	30
Trial 2	30	31	30	28	30	30
Trial 3	30	30	31	31	31	30
Mean count	30	30	30	29	30	30
Standard deviation	0	0	0	± 1	0	0

Table 2 Particle count of the mixture of 2-μm and 750-nm PS beads in DI water using flow-channeled DIC system.

Particle count (particles/microchannel volume, 0.025 μL)
	Number of 2-μm PS beads		Number of 750-nm PS beads
	Static condition	Flow condition	Static condition	Flow condition
Trial 1	30	32	30	33
Trial 2	30	30	30	31
Trial 3	30	30	30	28
Mean count	30	30	30	30
Standard deviation	0	± 1	0	± 2

Table 3 Particle count of 2-μm, 750-nm, and 200-nm PS beads in tap water using flow-channeled DIC system.

Particle count (particles/microchannel volume, 0.025 μL)
	Number of 2-μm PS beads		Number of 750-nm PS beads		Number of 200-nm PS beads
	Static condition	Flow condition	Static condition	Flow condition	Static condition	Flow condition
Trial 1	30	26	30	31	30	29
Trial 2	30	30	30	24	30	24
Trial 3	30	31	30	30	30	30
Mean count	30	29	30	28	30	27
Standard deviation	0	± 2	0	± 3	0	± 3

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Real-time detection of label-free submicron-sized plastics using flow-channeled differential interference contrast microscopy

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Abstract

Figures

Introduction

Results And Discussion

Imaging the PS beads using high-magnification DIC microscope

Cross-validation and confirmation: comparison of DIC microscope and Raman spectroscopy

Flow-channeled DIC system: Cross-validation and confirmation on PS bead detection in DI water

Quantitative evaluations of detected PS beads in DI water

Detection of mixture of sizes of PS beads in DI water

Detection of PS beads in tap water

Evaluation of the flow-channeled DIC system in natural water, and environmental significance

Methods

Microplastics and preparation

High-magnification DIC microscope

Flow-channeled DIC system

Fabrication of microfluidic chip

SEM and micro-Raman spectroscopy

Declarations

References

Tables

Additional Declarations

Supplementary Files

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