Even if ammonia gas production is reported in previous reports [24] and color change of phenol red indicates pH change in basic condition within droplets, it was not clearly identified that ammonia gas is the metabolite crosstalk between droplets. To validate this, the production of ammonia gas during bacterial growth, bacterial metabolite was determined with Nessler's reagents, which is the conventional indicator of ammonia gas (Figure 10). Phenol red turned red with *E. coli* K-12 culture in LB broth (Figure 10b,e), and Nessler's reagent also immediately reacted with *E. coli* K-12-containing LB broth (Figure 10c,f). These results suggest that pH change was raised due to the production of ammonia gas during bacterial growth and ammonia gas was the metabolite which was reacted with phenol red.

#### *3.3. Validation of Ammonia Gas Crosstalk between Droplets*

To identify the permeability of ammonia gas through the droplet barrier, sodium hydroxide and ammonium chloride were co-encapsulated within 80 μm droplets for producing ammonia gas (ammonia gas generating droplets) [24]. As shown in Figure 11, we hypothesized that ammonia gas production could be mimicked by co-encapsulating sodium hydroxide and ammonium chloride. If produced ammonia gas crosstalk into phenol red droplets, the pH indicator will turn red. To validate our hypothesis, droplets were separately generated with 2 M sodium hydroxide (Figure 12b) or 2 M ammonium chloride (Figure 12e). Each droplet was then mixed with phenol red droplets (Figure 12a,d,g), but a color change was not observed (Figure 12c,f). However, phenol red droplets turned red when they were mixed with droplets that co-encapsulated with sodium hydroxide and ammonium chloride to produce ammonia gas within droplets (Figure 12i). These results indicate that ammonia gas was produced from the chemical reaction, and also produced gas was transferred through the droplet barrier.

#### *3.4. E*ff*ects of Ammonia Gas Concentration and Surfactant Concentration on Crosstalk between Droplets*

To identify the effect of gas concentration on permeability between droplets, sodium hydroxide and ammonium chloride were co-encapsulated at a different concentration from 0.002 M to 2 M. Collected droplets were then mixed with indicator droplets and imaged immediately. More bright color changes were observed from indicator droplets due to a higher production of ammonia gas (Figure 13). This result indicates that gas crosstalk was increased in a higher concentration of gas.

In previous reports on DMF, PFPE–PEG–PFPE surfactants have been used at a concentration of 2% on average, which is why we have identified gas crosstalk with a 2% surfactant in this report. However, the effect of surfactant concentration also identified whether it affects gas permeability between droplets. To validate this hypothesis, 0.02 M sodium hydroxide and 0.02 M ammonium chloride were co-encapsulated with HFE-7500 that contains different concentrations of surfactant at 0.8, 2 and 5%. Collected ammonia gas generating droplets were then mixed with indicator droplets, and gas permeability was decreased with a higher concentration of surfactant (Figure 14). These results suggest that the droplet barrier became more densely when surfactant concentration was increased, and it decreased gas permeability.

#### *3.5. Time-Course Measurement of Ammonia Gas Crosstalk through Droplet Barrier*

To measure the time-course transfer of produced ammonia gas through the droplet barrier, 0.02 M sodium hydroxide and 0.02 M ammonium chloride were co-encapsulated with HFE-7500 containing 2% surfactant prior to mix with indicator droplets. Once the indicator droplet was contacted with ammonia gas generating droplets, it started to turn red in a time-dependent manner (Figure 15, Supplementary Video S1). When sodium hydroxide and ammonium chloride were encapsulated at a concentration of 0.2 M for both chemical compounds, the color change was immediately terminated turned red once the indicator droplet was contacted with ammonia gas generating droplets. We also observed that ammonia gas could be transferred through fluorinated oil even without contact between droplets. As shown in Figure 16 (Supplementary Video S2), phenol red-containing droplets turned red slowly due to gas diffusion through HFE-7500 from two white droplets (that contain 0.2 M sodium hydroxide and 0.2 M ammonium chloride). This result suggested that ammonia gas is soluble in HFE-7500 oil.

**Figure 16.** Time-course measurement of ammonia gas crosstalk between droplets. (**a**) Microscopic images of droplet mixture between indicator droplet and ammonia gas generating droplets, the video was recorded for 24 s under a DMi-8 microscope. (**b**) Mean values of color change in the indicator droplets.

#### **4. Discussion**

The PFPE–PEG–PFPE surfactant is one of the most commonly used surfactants in DMF, and it was mentioned that fluorinated fluids exhibit high gas solubility [23,35]. However, gas crosstalk between droplets has not been validated yet. In this study, we first demonstrated that bacterial metabolic gas penetrates through barriers between droplets, and it contaminates neighboring droplets. Interestingly phenol red dye cannot penetrate the barrier between droplets due to its relatively higher molar mass (354.38 g/mol), but ammonia gas can across the droplet barrier because of its lower molar mass (17.031 g/mol) in our hypothesis.

It is already known that the weak interactions of fluorinated compounds in relatively high compressible fluids result in high gas solubility, so the interfacial surfactant molecules may be the main barriers that affect gas crosstalk [22]. To validate this hypothesis, the relation of surfactant concentration and gas transport was identified. To provide a higher surface density, surfactant concentration was increased up to 5%, and it decreased gas permeability between droplets and this result indicates surfactant density is the potential reason for the gas crosstalk between droplets.

We also confirmed that ammonia gas was produced during cultivation within droplets, and these results suggest that live bacteria can be grown and monitored using DMF by analyzing the production of ammonia gas. Since bacterial contamination is one of the most serious issues in drinking water, bacterial growth can be monitored by the identification of ammonia gas production using DMF without complex and expensive approaches such as real-time PCR. Even droplets were analyzed with an expensive and large microscope in our study; the microscope can be replaced with inexpensive and portable imaging systems, including smartphones and portable CCD cameras.

Basically, leaking of the droplets is the potential issue on analytical approaches, including basic biological studies and diagnosis because some of the interested biomarkers can be penetrated into neighboring droplets if it is small enough. Since contamination can be a potential reason for false-positive droplets, this issue must be eliminated in case of various diagnosis purposes. Even large-sized, highly hydrophobic and highly hydrophilic components have less chance for crosstalk through the droplet barrier; there is still potential that tiny components can be a useful biomarker for diseases. It turns that a dense droplet barrier must be constructed, which means stronger surfactants must be required for DMF in the future. However, leaking also can be a useful tool that can provide internal components to neighboring droplets. For example, bacteria droplets can be a tiny microcapsule that can supply ammonia gas into the microsized reactor that requires ammonia gas for the reactions such as chemical synthesis, analytical procedures and catalytic reactions.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2079-6374/10/11/172/s1, Video S1: Gas crosstalk between droplets when droplets are contected. Video S2: Gas crosstalk through fluorinated oil without contact between droplets.

**Author Contributions:** Conceptualization, S.K. and D.-K.K.; methodology, S.K.; software, S.K.; validation, S.K.; formal analysis, S.K.; investigation, D.-K.K.; resources, D.-K.K.; data curation, D.-K.K.; writing—original draft preparation, S.K.; writing—review and editing, D.-K.K.; visualization, S.K.; supervision, D.-K.K.; project administration, D.-K.K.; funding acquisition, D.-K.K. Both authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the Incheon National University Research Grant in 2017 and the National Research Foundation of Korea (NRF) (NRF2016R1A5A1010148).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

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© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

## **The E**ff**ect of Optically Induced Dielectrophoresis (ODEP)-Based Cell Manipulation in a Microfluidic System on the Properties of Biological Cells**

### **Po-Yu Chu 1,**†**, Chia-Hsun Hsieh 2,3,4,**†**, Chien-Ru Lin 5 and Min-Hsien Wu 3,5,6,\***

Received: 11 May 2020; Accepted: 14 June 2020; Published: 16 June 2020

**Abstract:** Cell manipulation using optically induced dielectrophoresis (ODEP) in microfluidic systems has attracted the interest of scientists due to its simplicity. Although this technique has been successfully demonstrated for various applications, one fundamental issue has to be addressed—Whether, the ODEP field affects the native properties of cells. To address this issue, we explored the effect of ODEP electrical conditions on cellular properties. Within the experimental conditions tested, the ODEP-based cell manipulation with the largest velocity occurred at 10 Vpp and 1 MHz, for the two cancer cell types explored. Under this operating condition, however, the cell viability of cancer cells was significantly affected (e.g., 70.5 ± 10.0% and 50.6 ± 9.2% reduction for the PC-3 and SK-BR-3 cancer cells, respectively). Conversely, the exposure of cancer cells to the ODEP electrical conditions of 7–10 Vpp and 3–5 MHz did not significantly alter the cell viability, cell metabolic activity, and the *EpCAM*, *VIM*, and *ABCC1* gene expression of cancer cells. Overall, this study fundamentally investigated the effect of ODEP electrical conditions on the cellular properties of cancer cells. The information obtained is crucially important for the utilization of ODEP-based cell manipulation in a microscale system for various applications.

**Keywords:** optically induced dielectrophoresis; microfluidic systems; cell manipulation; cell property; cell physiology

### **1. Introduction**

The fine manipulation of biological cells has attracted considerable interests in a wide variety of biomedical studies (e.g., cell sorting [1–8], the isolation and purification of cells [1–9], cell patterning [10–14], tissue engineering [10,11,14], or various cell-based bioassays [12,13]). These attempts are technically challenging to realize conventionally, due to the inadequate scale of tools or equipment used for cell manipulation in general biological laboratories. Leveraging the appropriate dimensional feature in a microfluidic system, and various novel approaches for microparticle manipulations (e.g., acoustophoresis [1,13], magnetophoresis [2], thermophoresis [15], and dielectrophoresis (DEP) [3,4]), researchers could achieve the above-mentioned goals in a high performance manner [16,17].

Among the techniques for microparticle manipulation in a microfluidic system, microparticle manipulation using DEP mechanism has been widely adopted for various applications (e.g., microparticle enrichment [18,19], microparticle patterning [10,11], cell-type classification [20], or rare cell isolation [3,4]). DEP-based microparticle manipulation, first presented in the 1950s [21], is well-described elsewhere [22], and can be briefly described as follows. When dielectric microparticles (e.g., biological cells) are suspended in a solution in which an electric field is exerted, charges can be electrically polarized on the microparticles' surface. The interaction between the induced charges on the microparticles and the electric field specifically exerted on them can generate a DEP force on such microparticles [22]. In practice, therefore, the scientists can delicately control the electric field exerted on biological cells via a designed microelectrode array, to manipulate these cells in a manageable manner. Although the DEP technique is proven feasible for the fine manipulation of cells, this technique normally requires a costly, time-consuming, and technically demanding microfabrication process to create a unique metal microelectrode layout that is specific to the application [23,24]. In addition, cell manipulation using the DEP mechanism might be technically challenging for general scientists, due to its complexity. To tackle the technical issues, the optically induced dielectrophoresis (ODEP)-based technique could open up a new horizon for the manipulation of microparticles in a more efficient, flexible, and user-friendly manner [23,24].

Microparticle manipulation based on the ODEP mechanism was first proposed in 2005 [23]. Similar to the DEP mechanism described earlier, an electric voltage is applied between the top and bottom substrates of an ODEP system to generate a uniform electric field in the solution layer sandwiched between the two substrates. Under this circumstance, the dielectric microparticles suspended in the solution layer are electrically polarized. In contrast to the DEP mechanism, however, the key technical merit of the ODEP mechanism is that it can easily and quickly create or modify an electrode layout through the control of optical patterns, by acting as a virtual electrode [23,24]. When the photoconductive layer of an ODEP system is briefly projected with light, it can lead to a significant reduction in electrical impedance in the light-illuminated area. This phenomenon can cause the exerted voltage to decrease across the solution layer inside the light-projected region. This phenomenon, therefore, generates a locally nonuniform electric field within an ODEP system. For microparticle manipulation, scientists can simply utilize a commercial projector to project optical images on an ODEP system, by which the interaction between the generated nonuniform electric field and an electrically polarized microparticle is used to manipulate the microparticle. In terms of operation, one can control the light image projected onto the ODEP system and thus the nonuniform electric field created to manipulate dielectric microparticles within the ODEP system.

The use of ODEP mechanism in microfluidic systems has been successfully applied for the high-accuracy positioning and assembling of metallic beads [25], dynamic analysis of cancer-immune microenvironment [26], or the isolation and purification of rare cell species in clinical samples (e.g., Raji cells [7], circulating tumor cells (CTCs) [5,9], and CTC clusters [6]) in a higher performance manner than their conventional counterparts. Although the incorporation of the ODEP mechanism in a microscale system has provided a powerful tool for the biological cell-relevant studies or applications, one fundamental question has to be answered before its widespread applications—Whether the cell manipulation using the ODEP mechanism influences the cellular properties. If this is the case, the use of such a mechanism for cell manipulation could complicate the subsequent biological studies or applications. To the best of our knowledge, however, this fundamental issue has not yet been well explored.

To investigate the effect of ODEP-based cell manipulation (the operating conditions explored—(1) the magnitude of AC voltage: 5–10 V; (2) frequency of AC voltage: 1–5 MHz, (3) operating time: 3 min) on the properties (e.g., cell viability, metabolic activity of cells, or gene expression of cells) of biological cells (e.g., PC-3 and SK-BR-3 cancer cells), we performed various experiments. Within the experimental conditions investigated, the ODEP-based cell manipulation with the best performance (e.g., highest cell manipulation velocity) occurred at 10 Vpp and 1 MHz, for the two cancer cell types

explored. Under this ODEP electrical condition, however, the cell viability of the cancer cells tested was significantly affected (e.g., 70.5 ± 10.0% and 50.6 ± 9.2% reduction for the PC-3 and SK-BR-3 cancer cells, respectively), possibly due to the electrical lysis of cells. Conversely, the exposure of cancer cells to the ODEP field with particular conditions (e.g., 7–10 Vpp, 3–5 MHz, and 3 min exposure time) might not significantly alter the cell viability, cell metabolic activity, and gene expression of cancer cells. Within this ODEP operating condition, the highest maximum velocity (e.g., 106.7 ± 30.6 μm s−1 and 100.0 ± 20.0 μm s−1 for the PC-3 and SK-BR-3 cancer cells, respectively) of a light image that can manipulate cells occurred at the voltage magnitude and frequency of 10 Vpp and 3 MHz, respectively. The above-mentioned cell manipulation velocities are technically sufficient for various applications. As a whole, this study fundamentally investigated the effect of ODEP electrical conditions on the cellular properties of cancer cells. The information obtained is crucially important for the utilization of ODEP-based cell manipulation in a micro-scale system for various applications.

#### **2. Materials and Methods**

#### *2.1. Microfluidic Chip and Experimental Setup*

To explore the effect of ODEP on the properties of biological cells, we designed a simple microfluidic chip. Figure 1a schematically presents the top-view layout of the microfluidic chip, mainly encompassing a microchamber (L = 6.0 mm, W = 4.0 mm, and H = 50.0 μm) connecting two microchannels (L = 2.0 mm, W = 1.0 mm, and H = 50.0 μm). In this work, an ODEP field was applied to the microchamber (Figure 1a). The structure of the microfluidic chip is illustrated in Figure 1b. Briefly, the microfluidic chip consists of two custom-made polydimethylsiloxane (PDMS) (Sylgard® 184, Dow Corning, Midland, MI, USA) adapters for tubing connection (A), an indium-tin oxide (ITO) glass (10 Ω, 0.7 mm; Innolux Corp., Miaoli County, Taiwan) (B), a processed adhesive tape (L298, Sun-yieh, New Taipei City, Taiwan; thickness: 50 μm) containing hollow microchamber and microchannel structures (C), and a bottom ITO glass (D) with a coating layer of photoconductive material (a 1.0 μm-thick hydrogenated amorphous silicon (a-Si:H) layer). The fabrication processes (e.g., PDMS replica mold, metal mold-punching fabrication, plasma-enhanced chemical vapor deposition (PECVD)-based thin-film technology, and plasma oxidation-aided bonding) were well-described previously [8,27,28]. After each substrate was fabricated (Figure 1b), substrate A was bonded with substrate B through the surface treatment of plasma oxidation. This step was followed by the assembly with the substrate D with the aid of the processed double-sided adhesive tape (substrate C) (Figure 1b). In operation, the prepared cell suspension was manually loaded into the microchamber, using a pipette and a tip. For cell manipulation using the ODEP mechanism, the commercially available ODEP-based cell isolation equipment (Celnostics, Ace Medical Technology Co., Ltd., Taipei City, Taiwan) was used to achieve the ODEP-based cell manipulation in the proposed microfluidic chip. Within this equipment, briefly, a function generator (AFG-2125, Good Will Instrument Co., Ltd., New Taipei City, Taiwan) was utilized to apply an AC bias between the two ITO glasses. A computer-interfaced digital projector (EB-X05, Epson, Nagano, Japan) was used to illuminate light images onto the photoconductive material (i.e., the Substrate D) of microfluidic chip to generate the ODEP force on the cells. An illustration of the important operating modules of the equipment is schematically presented in Figure 1c (a photograph of the overall experimental setup is provided in a Supplementary Figure S1).

**Figure 1.** Schematic presentation of (**a**) the design of the microfluidic chip (top view), (**b**) the structure of the microfluidic chip ((A) polydimethylsiloxane (PDMS) adapters for tubing connection, (B) indium-tin oxide (ITO) glass, (C) processed adhesive tape, and (D) ITO glass with a coating layer of photoconductive material), and (**c**) the overall experimental setup.

#### *2.2. The Assessment of the ODEP Manipulation Force Generated on Cells*

The working principle of ODEP for cell manipulation is described in the Introduction. The ODEP force generated on a cell can theoretically be described by Equation (1) [24,27,28]:

$$F\_{DEP} = 2\pi r^3 \varepsilon\_0 \varepsilon\_m \text{Re} [\mathbf{f}\_{\text{CM}} \| \mathbf{V} | E]^2 \tag{1}$$

In Equation (1), *r* (cellular radius), ε*0* (vacuum permittivity), ε*m* (relative permittivity of the surrounding solution), ∇|*E*| *2* (gradient of electric field squared), and Re[fCM] (real part of the Clausius–Mossotti factor (fCM)) are the important parameters [24,28]. The fCM is described by Equation (2) [29–31]:

$$\mathbf{f\_{CM}} = \frac{\boldsymbol{\varepsilon^\*\_{cell}} - \boldsymbol{\varepsilon^\*\_{m}}}{\boldsymbol{\varepsilon^\*\_{cell}} + 2\boldsymbol{\varepsilon^\*\_{m}}} \tag{2}$$

where ε∗ *cell* and ε∗ *m* represent the complex permittivity of the cell and the surrounding solution, respectively. For a single-cell model, the complex permittivity of the cell and the surrounding solution can be further described by Equations (3) and (4):

$$\varepsilon^{\*}\_{\rm ccell} = \mathcal{C}^{\*}\_{\rm mcm} \frac{3\text{Re}^{\*}\_{\rm int}}{3\varepsilon^{\*}\_{\rm int} + 3\mathcal{C}^{\*}\_{\rm mcm}\mathcal{R}}, \mathcal{C}^{\*}\_{\rm mcm} = \frac{\varepsilon\_{\rm mcm}}{d} - \frac{\text{j}\sigma\_{\rm mcm}}{d}, \varepsilon^{\*}\_{\rm int} = \varepsilon\_{0}\varepsilon\_{\rm int} - \frac{\text{j}\sigma\_{\rm int}}{\alpha} \tag{3}$$

$$
\varepsilon\_m^\* = \varepsilon\_0 \varepsilon\_m - \frac{\text{j}\sigma\_m}{\alpha} \tag{4}
$$

In Equations (3) and (4), *C*∗ *mem* represents the complex cell membrane capacitance, ε∗ *int* represents the complex permittivity of the cellular interior (i.e., cell cytoplasm), *R* represents the radius of the cellular interior, d represents the thickness of cell membrane, ε represents the relative permittivity of the cell membrane, cellular interior, or surrounding solution (denoted by the subscript *mem*, *int*, or *m*, respectively), σ represents the conductivity of the cell membrane, cell interior, or surrounding solution (denoted by the subscript *mem*, *int*, or *m*, respectively), j represents the imaginary vector ( √ −1), and ω represents the angular frequency (i.e., ω = 2π*f*) of the applied AC field, respectively [29–31]. Under a given solution condition, overall, the magnitude and frequency of the electric voltage applied could play important roles in the ODEP force generated on a particular cell [24,28]. In this work, the effect of electric conditions (i.e., the magnitude and frequency of the electric voltage exerted) on the ODEP force generated on cells was experimentally evaluated. In the evaluation, the ODEP manipulation force, a net force between the ODEP and the friction force, acting on the manipulated cells was then experimentally assessed, based on the method described previously [8,27,28]. In a steady state, the ODEP manipulation force acting on a cell was balanced by the viscous drag of fluid acting on such a cell under a continuous flow condition. As a result, the hydrodynamic drag force of a moving cell was used to evaluate the net ODEP manipulation force of a cell according to Stokes' law (Equation (4)):

$$F = 6\pi r \eta v \tag{5}$$

In Equation (5), *r* (cellular radius), η (fluidic viscosity), and *v* (the velocity of a moving cell) are the important parameters. Under the given solution and cellular size conditions, overall, the ODEP manipulation force of the manipulated cell could then be experimentally assessed through the measurement of the maximum velocity of a moving optical image that can manipulate such a cell [8,27,28]. In practice, briefly, a light bar image with different moving velocities (e.g., from low to high velocities) was used to manipulate a cell (e.g., attracted and pulled a cell). Through this process, the maximum velocity of a moving optical image that can manipulate such a cell was then determined. In this work, therefore, the above-mentioned velocity was utilized as an index for the evaluation of the ODEP manipulation force generated on a specific cell under a particular electric condition. Based on this, the effect of electric conditions (e.g., magnitude of AC electric voltage: 7–10 Vpp and frequency of AC electric voltage: 1–5 MHz) on the ODEP manipulation of the cells tested (e.g., PC-3 and SK-BR-3 cancer cells) was evaluated. Briefly, the cell sample tested was prepared in a cell suspension (cell density: 106 cells mL−1), followed by loading into the microchamber of the microfluidic chip (Figure 1a). The maximum velocity of a moving light bar (L: 1.3 mm W: 100.0 μm) that could manipulate these cells was then assessed [27,28].

#### *2.3. Evaluation of the Properties of Cancer Cells Treated with Varied ODEP Operating Conditions*

For the analysis of the ODEP effect on the cellular properties, the cancer cells tested (e.g., PC-3 and SK-BR-3 cancer cell lines, two of the commonly-used cancer cell lines in cancer-related studies [32,33]) were first treated with the ODEP fields under different conditions for 3 min, followed by assaying their cellular properties, including cellular viability, cellular metabolism activity, and gene expression. In this study, the biological assays were carried out at 1.5 ± 0.2 h after the ODEP exposure treatment. In brief, the background medium of the prepared cancer cell suspension (cell density: 5 × 106 cells mL−1 for PC-3 cancer cells, and 3 × 106 cells mL−1 for SK-BR-3 cancer cells) was first replaced by a 9.5% (*w*/*v*) sucrose buffer solution (relative permittivity: ~76.19 [34], fluid viscosity: ~1.0389 × 10−2 g s−1 cm−1 [35], osmolality: 270–290 mOsmol kg−1, and conductivity: 1–5 μS cm−1), the commonly used low conductivity buffer solution for ODEP-based cell manipulation [5,8,9,27,28]. The processed cell suspension sample was then loaded into the microchamber of a microfluidic chip (Figure 1a). In this process, however, a small portion of cells might retain in the microchannel area. In this work, the cell manipulation using ODEP was first performed to quickly (operation time—within 5 s) transport these cells to the microchamber area before ODEP treatment. At the microchamber zone, 40 rectangular light bar images (L: 4.0 mm, W: 100.0 μm, the interval between light bars: 50.0 μm) were then illuminated to provide an ODEP field on the cancer cells tested. In this study, ODEP fields with different electric conditions (magnitude of AC electric voltage: 7–10 Vpp and frequency of AC electric voltage: 1–5 MHz) were used to treat the cancer cells tested. After exposure to the ODEP field for 3 min, the treated cells were then harvested for the subsequent bioassays. In the process, briefly, a suction-type syringe pump was utilized to collect the treated cancer cells through the hole for harvesting cells (Figure 1a). Before the following bioassays, the collected cells were kept in the form of cell suspension in the original sucrose buffer solution under the thermal condition of 25 ◦C.

In this work, the commonly-used Cell Counting Kit-8 (CCK-8) (CK04–05, Dojindo, Kumamoto, Japan) and ATP Colorimetric/Fluorometric assays (K354-100, BioVision Inc., Milpitas, CA, USA) were utilized to assay the cell viability and cell metabolic activity of cancer cells, respectively. The bioassays were carried out according to the manufacturer's instructions. For further analysis of whether cell manipulation using ODEP could affect cellular gene expression, the expression levels of genes in cancer cells treated with different ODEP fields were experimentally evaluated. As the ODEP-based cell manipulation was commonly used for the isolation and purification of circulating tumor cells (CTCs) [5,9], the important CTC-related gene expressions were thus explored in this work. For the cancer cells (PC-3 and SK-BR-3 cancer cells) tested, the mRNA levels of epithelial-to-mesenchymal transition (EMT)-associated genes [*EpCAM* (Hs00158980\_m1) and *VIM* (Hs00958111\_m1)], the multidrug resistance-associated protein 1 (MRP1) gene [*ABCC1* (Hs01561502\_m1)], and the housekeeping gene [*GAPDH* (Hs02758991\_g1)] were experimentally quantified. The bioassay was based on a method described previously [8,9,27]. In brief, RNA was extracted from the cancer cells tested using a bromochloropropane (BCP)-based TRI Reagent procedure (Thermo Fisher Scientific, San Jose, CA, USA [36]). This process was followed by the reverse transcription using a SuperScript® IV Reverse Transcriptase Kit (Thermo Fisher Scientific, San Jose, CA, USA). The mRNA level was subsequently quantified using a StepOne™ Real-Time PCR System (Thermo Fisher Scientific, San Jose, CA, USA).

#### *2.4. Statistical Analysis*

In this study, data were obtained from three separate experiments, and are presented as the mean ± standard deviation (n = 9). To compare the results from different operating conditions, we used one-way ANOVA and Tukey's honestly significant difference (HSD) post-hoc test for the statistical analysis.

#### **3. Results and Discussion**

#### *3.1. E*ff*ect of the Electric Conditions on ODEP-Based Cell Manipulation*

In this study, the effect of the ODEP electrical conditions (e.g., magnitude of AC electric voltage: 7–10 Vpp, and frequency of AC electric voltage: 1–5 MHz) on the cell manipulation of the cancer cells (e.g., PC-3 and SK-BR-3 cancer cells) tested were experimentally evaluated. Figure 2 reveals the quantitative relationship between the maximum velocity of a moving light bar that can manipulate the cancer cells tested (thus, the ODEP manipulation force of the cancer cells; Equation (5)) and the frequency of electric voltage applied under different voltage magnitude conditions. The results (Figure 2a,b) showed that the highest values of the maximum velocity of a light image that can manipulate the cancer cells, both occurred at 10 Vpp and 1 MHz for the two cancer cell types explored (i.e., 475.0 ± 25.0 μm s−1 and 458.3 ± 38.2 μm s−1 for the PC-3 and SK-BR-3 cancer cells, respectively). When the voltage frequency was lower than 1 MHz, some undesirable phenomena such as cell adhesion (10 Vpp and 500 kHz) (Figure 2c), cell aggregation (10 Vpp and 750 kHz) (Figure 2d), and cell lysis (10 Vpp and 100 kHz) (Figure 2e), would occur that could significantly affect the cellular properties [28]. Within the experimental conditions explored, moreover, the appearance rate of the above-mentioned phenomena was near 100% when the operating conditions were set at the particular electric conditions as indicated. These resulting phenomena might be due to the enhancement of electrically induced cell deformability, mutual DEP, and electrical lysis of cells at a voltage frequency near or below 1 MHz [26]. When the electric condition was set at 7 Vpp and 5 MHz (Figure 2a,b), conversely, we

observed that the maximum velocities of a light image that can manipulate the cancer cells tested both reached their lowest levels for the two cancer cell types tested (i.e., 3.3 ± 5.8 μm s−1 and 10.0 ± 10.0 μm s−1 for the PC-3 and SK-BR-3 cancer cells, respectively) within the experimental conditions investigated. When the voltage magnitude and frequency were lower and higher than 7 Vpp and 5 MHz, respectively, the above-mentioned velocities were further decreased, which might significantly affect cell manipulation using the ODEP mechanism. Based on the facts described above, the optimal window of electric conditions for effective cell manipulation using ODEP was set at 7–10 Vpp and 1–5 MHz for the voltage magnitude and frequency, respectively. The ODEP operating conditions within such ranges were also reported for the ODEP-based cell manipulation for different applications [7,8,27,28]. Within this electric condition range, moreover, the maximum velocity of a light image that could manipulate the cancer cells tested decreased significantly with an increase of voltage frequency (Figure 2a,b), which was also consistent with previous findings [28]. Furthermore, under a given voltage frequency condition, the increase in voltage magnitude was found to increase the maximum velocity of a light image that could manipulate the cancer cells tested. This phenomenon could be explained by Equation (1), in which the ODEP force generated on a cell is proportional to the electric field squared. In addition to the values of the maximum velocity of a light image (and thus the ODEP force) investigated in the above-mentioned studies, it might also be valuable to explore the values of electric-field distribution in ODEP-based cell manipulation.

**Figure 2.** The quantitative relationship between the maximum velocity of a moving light bar that could manipulate the (**a**) PC-3 and (**b**) SK-BR-3 cancer cells tested and the frequency (1, 2, 3, 4, and 5 MHz) of the electric voltage applied under different voltage magnitudes (7, 8.5, and 10 Vpp) conditions and bright-field microscopic images showing the phenomena of (**c**) cell adhesion (10 Vpp and 500 kHz), (**d**) cell aggregation (10 Vpp and 750 kHz), and (**e**) cell lysis (10 Vpp and 100 kHz), during the exposure of cancer cells to the ODEP field with a voltage magnitude and frequency, as indicated.

#### *3.2. E*ff*ect of the ODEP Field on the Cellular Viability of Cancer Cells*

In this study, PC-3 and SK-BR-3 cancer cells, two of the commonly used cancer cell lines in the cancer-related studies, were used as model cells to investigate the effect of the ODEP-based cell manipulation on cellular properties. In the operations, the prepared cancer cell suspension was loaded into the microchamber of the ODEP microfluidic chip (Figure 1a), followed by exposure to the ODEP field with varied electric conditions (i.e., 7–10 Vpp and 1–5 MHz for the voltage magnitude and frequency, respectively), as determined previously (Figure 2). In our preliminary tests, the impact of ODEP exposure time on the cell viability of PC-3 cancer cells was investigated. Results (Supplementary Figure S2) showed that the cell viability of such cells had no significant difference (*p* > 0.05) compared to the control (i.e., the cancer cells without exposure to the ODEP field) within the 5 min exposure time, when the voltage magnitude and frequency were set at 10 Vpp and 5 MHz, respectively. When the ODEP exposure time was as long as 10 and 15 min, conversely, the cell viability of the PC-3 cancer cells might be significantly affected (*p* < 0.05) (e.g., 43.7 ± 5.5% and 51.3 ± 10.3% reduction for the ODEP exposure time of 10 and 15 min, respectively) in comparison to the control. Based on the preliminary results, therefore, the ODEP exposure time was set at 3 min, and within the time period, general cell manipulation using the ODEP mechanism was normally carried out (e.g., the rapid separation of Raji cells [7] or isolation and purification of CTCs [5,9]). In practice, 40 rectangular light bar images were illuminated on the microchamber zone. During the period of ODEP exposure, the cancer cells tested were attracted within the light bars, as shown in Figure 3(aII,aV). After the exposure to the ODEP field, we observed microscopically that the treated cancer cells within the microchamber were aligned in accordance with the original light bar images, as exhibited in Figure 3 (aIII,aVI). Then, the ODEP-treated cancer cells were collected for the following bioassays.

**Figure 3.** (**a**) Microscopic observation (close-up view) of the cancer cells in the microchamber of the microfluidic chip before (the upper row), during (the middle row), and after (the lower row) treatment with ODEP exposure (the left column—PC-3 cancer cells; the right column—SK-BR-3 cancer cells), (**b**) the comparison of relative cell viability (%) of (I) the PC-3 cancer cells and (II) the SK-BR-3 cancer cells treated with the ODEP field with varied electric conditions (voltage magnitude and frequency: 7, 8.5, and 10 Vpp and 1, 3, and 5 MHz, respectively) (NS—No significant difference (*p* > 0.05), \* significant difference (*p* < 0.05)).

In this study, the Cell Counting Kit-8 (CCK-8) was utilized to investigate the viability of cancer cells treated with various ODEP fields. The results (Figure 3(bI)) revealed that the cell viability of PC-3 cancer cells had no significant difference (*p* > 0.05) compared to that of the control cells (i.e., the cancer cells without exposure to the ODEP field), when the voltage magnitude and frequency were set at 7–10 Vpp and 3–5 MHz, respectively. A similar result was also found for the SK-BR-3 cancer cells tested (Figure 3(bII)). However, once the voltage frequency was set at 1 MHz, the cell viability of the PC-3 and SK-BR-3 cancer cells decreased significantly (*p* < 0.05) compared to that of the control, irrespective of a voltage magnitude of 7, 8.5, or 10 Vpp. This finding could be explained as follows. If the equivalent circuit of a biological cell is assumed to be a lumped circuit, the electrical property of the cell membrane will be similar to the electric capacitance [28]. When the voltage frequency is altered from high to low frequency (e.g., from 10 MHz to 100 kHz), the electrical impedance of a cell membrane might increase. This phenomenon could, therefore, result in the shift of the exerted electric voltage from the cell cytoplasm to the cell membrane of a cell [28]. This fact might in turn lead to an increase in the transmembrane potential in a cell membrane when the voltage frequency is changed from high to low frequency. The above-mentioned phenomena could explain the low cell viability caused by cell electrical lysis (Figure 2e) when the voltage frequency is set at low conditions (e.g., 1 MHz) (Figure 3b).

Under the low voltage frequency condition of 1 MHz (Figure 3(bI)), moreover, the cell viability of the PC-3 cancer cells significantly decreased (*p* < 0.05) when the magnitude of voltage was higher than 8.5 Vpp (observed cell viability: 40.4 ± 3.6%). A similar result was also found for the SK-BR-3 cancer cells tested (Figure 3(bII)), in which the cell viability of the SK-BR-3 cancer cells was significantly downregulated (*p* < 0.05) (the observed cell viability: 49.4 ± 9.5%) when the magnitude of the voltage was higher than 10 Vpp. Overall, the lower cell viability of cancer cells found at the higher voltage condition could be explained by the fact that the electrical lysis of cells is prone to occur when a cell is exposed to a high-voltage magnitude [37,38]. Taken together, the increase of voltage magnitude or decrease of voltage frequency, respectively, could raise the maximum velocity of a light image that can manipulate cells (i.e., the ODEP manipulation force of cells; Equation (5), Figure 2). Nevertheless, the increase in voltage magnitude or decrease in voltage frequency, respectively, could accordingly increase the tendency of electrical lysis of cells, which could affect cellular viability. Within the experimental conditions explored (i.e., 7–10 Vpp and 1–5 MHz), the determinant electrical condition that could significantly affect cell viability was a low-voltage frequency of 1 MHz, when the voltage magnitude range was 7–10 Vpp.

#### *3.3. E*ff*ect of the ODEP Field on the Cellular Metabolic Activity and Gene Expression of the Cancer Cells Tested*

As suggested by the previous results (Figure 3b), cell manipulation using the ODEP mechanism at a low voltage frequency of 1 MHz could be lethal to the cancer cells tested. Such an ODEP electrical condition was ruled out in subsequent studies. In the following evaluations, the effect of the ODEP fields with varied electrical conditions (i.e., voltage magnitude and frequency: 7–10 Vpp and 3–5 MHz, respectively) on the metabolic activity and gene expression of cancer cells were explored. For the former, the ATP synthesis of cells was used as an indicator based on the previous studies [39]. The results (Figure 4a) showed that the ATP level of the ODEP-treated PC-3 cancer cells showed no significant difference (*p* > 0.05) compared to that of the control (i.e., the cancer cells without exposure to the ODEP field), within the experimental conditions explored. A similar result was also observed for the SK-BR-3 cancer cells (Figure 4b). The findings in this study could indicate that the ODEP field could not affect the ATP synthesis of a cell within the electric conditions tested. This result was, to some extent, similar to that found in a previous DEP-based study [40].

**Figure 4.** Comparison of the relative ATP levels (%) of (**a**) the PC-3 cancer cells and (**b**) the SK-BR-3 cancer cells treated with the optically induced dielectrophoresis (ODEP) field under varied electric conditions (voltage magnitude and frequency—7 and 10 Vpp and 3 and 5 MHz, respectively) [NS—No significant difference (*p* > 0.05)].

In addition to the effect on cellular metabolic activity, the ODEP effect on the gene expression of cancer cells was investigated. In this study, the gene expression of EMT-associated genes (e.g., *EpCAM* and *VIM*) [8] and the MRP1 gene (e.g., *ABCC1*) [8] in the PC-3 and SK-BR-3 cancer cells was quantified and then normalized to the gene expression of *GAPDH*. The results (Figure 5a) showed that the gene expression of the above-mentioned genes in the PC-3 cancer cells was not significantly different (*p* > 0.05) from that of the control cells (i.e., the cancer cells without exposure to the ODEP field) within the ODEP operating conditions (i.e., 7–10 Vpp, 3–5 MHz, and 3 min exposure time) studied. A similar outcome (Figure 5b) was also found for the SK-BR-3 cancer cells.

**Figure 5.** The comparison of relative gene expression ((I) *EpCAM*, (II) *VIM*, and (III) *ABCC1* genes) of (**a**) the PC-3 cancer cells and (**b**) the SK-BR-3 cancer cells treated with the ODEP field with varied electrical conditions (voltage magnitude and frequency: 7 and 10 Vpp and 3 and 5 MHz, respectively) (NS—No significant difference (*p* > 0.05).

Previous studies similar to this work revealed that the exposure to a DEP field (e.g., 20 Vpp, 250 kHz, and 1 h exposure time) could downregulate the expression of cell differentiation-related genes (e.g., *ADGRE5*/*CD97*, *RUNX2*, and *NES*) in mesenchymal stem cells (e.g., UE7T-13 cells) [41]. It was also reported that the exposure to a DEP field (e.g., 21 Vpp, 5 MHz, and 15 min exposure time) might not affect the cell morphology, cell oxidative respiration rate, and cell cycle dynamics of fibroblast-like cells (e.g., BHK-21/C13 cells) [40]. After exposure to the DEP field, however, the gene expression of the c-fos protein (commonly activated by environmental stress [42]) was observed to be upregulated [40]. The reasons behind the discrepancies found in the previous and current studies

might be complicated and could be related to the cell species, electrical conditions, and the genes explored. Further experiments will be required to systematically investigate this issue. In addition to the gene expressions investigated in this study, it might also be valuable to explore the expression profiles of the genes regulating the signal pathways downstream of voltage-gated ion channels.

#### **4. Conclusions**

The ODEP-based microsystem provides a powerful tool for biological cell-relevant studies or applications. Before its widespread application, one fundamental issue has to be addressed—Whether cell manipulation using the ODEP mechanism influences the native properties of cells. To address this issue, we carried out various experiments. In this study, the impact of the ODEP electrical conditions ((1) the magnitude of AC voltage: 5–10 V; (2) frequency of AC voltage: 1–5 MHz), (3) operating time: 3 min) on the properties (e.g., cell viability, metabolic activity of cells, or gene expression of cells) of biological cells (e.g., PC-3 and SK-BR-3 cancer cells) was explored. Within the experimental conditions studied, the ODEP-based cell manipulation with the highest performance (e.g., highest cell manipulation velocity) occurred at 10 Vpp and 1 MHz for the two cancer cell types explored. Under this ODEP electrical condition, however, the cell viability of the cancer cells tested was significantly affected (e.g., 70.5 ± 10.0% and 50.6 ± 9.2% reduction for the PC-3 and SK-BR-3 cancer cells, respectively) possibly due to cell electrical lysis. Conversely, the exposure of cancer cells to the ODEP field with a particular condition range (i.e., 7–10 Vpp, 3–5 MHz, and 3 min exposure time) might not significantly alter the cell viability, cell metabolic activity, and the gene expression of cancer cells. Within this ODEP operating condition, the highest maximum velocity (i.e., 106.7 ± 30.6 μm s−1 and 100.0 ± 20.0 μm s−1, for the PC-3 and SK-BR-3 cancer cells, respectively) of a light image that could manipulate cells occurred at the voltage magnitude and frequency of 10 Vpp and 3 MHz, respectively. The above-mentioned cell manipulation velocity ranges were technically sufficient for various applications. Overall, this study fundamentally investigated the effect of ODEP electrical conditions on the cellular properties of cancer cells. The information obtained is crucially important for the utilization of ODEP-based cell manipulation in a microscale system for various applications.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2079-6374/10/6/65/s1, Figure S1: Photograph of the overall experimental setup, Figure S2: Comparison of relative cell viability (%) of the PC-3 cancer cells treated with varied ODEP exposed times as indicated (electrical condition: 10 Vpp and 5 MHz; ODEP exposed time: 1~15 minutes) [NS: No significant difference (*p* > 0.05), \* significant difference (*p* < 0.05)].

**Author Contributions:** Conceptualization, P.-Y.C., C.-H.H., and M.-H.W.; methodology, P.-Y.C., C.-H.H., and C.-R.L.; validation, P.-Y.C., C.-H.H., and C.-R.L.; writing—original draft preparation, P.-Y.C.; writing—review and editing, P.-Y.C., C.-H.H., and M.-H.W. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was funded by the Ministry of Science and Technology, R.O.C. (MOST 107–2221-E-182–033-MY3) and Chang Gung Memorial Hospital (CMRPD2G0061–62, CMRPD2H0121–23, and CMRPD2J0031–32).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Role of Paper-Based Sensors in Fight Against Cancer for the Developing World**

**Amey Dukle 1, Arputharaj Joseph Nathanael 1,\*, Balaji Panchapakesan 2 and Tae-Hwan Oh 3**

**Abstract:** Cancer is one of the major killers across the globe. According to the WHO, more than 10 million people succumbed to cancer in the year 2020 alone. The early detection of cancer is key to reducing the mortality rate. In low- and medium-income countries, the screening facilities are limited due to a scarcity of resources and equipment. Paper-based microfluidics provide a platform for a low-cost, biodegradable micro-total analysis system (μTAS) that can be used for the detection of critical biomarkers for cancer screening. This work aims to review and provide a perspective on various available paper-based methods for cancer screening. The work includes an overview of paper-based sensors, the analytes that can be detected and the detection, and readout methods used.

**Keywords:** paper-based sensors; cancer screening; disposable sensors; sensors; paper fluidics; microfluidics

#### **1. Introduction**

Cancer is a major cause of death worldwide [1,2]. It is estimated to be the cause of every 1 in 6 deaths [3,4]. According to the World Health Organisation (WHO), in the year 2020 more than 10 million people lost their life to cancer [5]. Worldwide, an estimated 19.3 million new cancer cases and almost 10.0 million cancer deaths occurred in 2020 [6]. Although the causes of cancer may vary depending on the type, it has been observed that the incidence rate of disease is on the rise [7,8].Worldwide, by 2040, 28.7 million new cases of cancer are projected to occur, a 47% rise compared to 19.3 million in 2020. An increase in the global cancer burden in the next fifty years will come from low- and middle-income countries (400% in low-income countries, 168% in middle-income countries, and 53% in high-income countries) [9,10]. Though there are different treatment strategies that have been developed and the disease is no longer 'incurable' [11–14], the success rate of the treatment depends on the stage of disease progression [15–17]. An individual undergoing treatment in the early stages of cancer has a many times higher chance of survival than in the later stages [18,19]. Estimates suggest that approximately 30–50% of cancer deaths can be prevented by early detection and treatment [20–22].

Although the incidence rate of cancer is higher in wealthy nations compared to lowand middle-income countries, low- and middle-income countries have a lower survival rate, partly due to the late presentation of the disease [23]. The barriers of cancer care in developing countries are due to late-stage presentation, quality of care, affordability, and a lack of access to advanced clinical resources. Late-stage presentation puts tremendous burden on clinicians [10]. The premature death and loss of productive life in the working population results in a significant economic impact on these countries.

In low- and middle-income countries, access to health care facilities is not readily available [24–28]. The number of physicians in low- to middle-income countries can be as

**Citation:** Dukle, A.; Nathanael, A.J.; Panchapakesan, B.; Oh, T.-H. Role of Paper-Based Sensors in Fight Against Cancer for the Developing World. *Biosensors* **2022**, *12*, 737. https:// doi.org/10.3390/bios12090737

Received: 22 July 2022 Accepted: 31 August 2022 Published: 7 September 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

low as 0.1 to 2 per 1000 people. In many cases, they are heavily burdened, leading to long waiting times [29]. In many low- and middle-income countries, cancer screening is not covered under insurance, thus discouraging patients from undergoing cancer screening.

To address the issue of cancer screening in these countries, various strategies have been developed [30]. These screening devices need to be easy to manufacture, low-cost, portable, and should not require any special training. Paper-based sensors are gaining significance in this field as they possess all the features of an ideal screening device [31–50].

Paper was invented by the Egyptians in the fourth century BC. It is one material that has existed continuously since the beginning of Egyptian civilization. Paper-based products are the most sold products in the world. For example, the Bible is the most sold book in the world and is printed on paper. The question then becomes: can we print cancer screening devices on paper to enable their use as low-cost sensors for cancer screening? Paper-based sensors for cancer screening are akin to an at-home pregnancy test kit. These kits will indicate whether a specific cancer biomarker is present in the person's body. In many cases, the result will be qualitative, i.e., yes/no type. However, novel sensors have been developed that provide a quantitative output [51].

In case the test returns a positive result, the patient can consult medical professionals so that exhaustive testing can be performed and the stage of the disease identified.

The most common sensing principle used in paper-based sensors for cancer screening is ELISA, wherein the analyte is labelled and detected using a sandwich assay [52]. The results are then readout using a plate reader to specify analyte concentration against a standard curve [53–57]. Various other methods are also being explored for the development of sensors.

This work reviews the various paper-based sensors that have been developed for cancer screening. In the next sections, a brief introduction about paper-based sensors, the analytes used for cancer screening, and the detection and read-out techniques used are discussed.

#### **2. Paper-Based Sensors: Low-Cost Screening Devices for the Developing World**

Advancements in microfluidics have led to their widespread usage in sensing applications. Microfluidics, as the name suggests, uses small fluidic channels where liquids, proteins and cells can be manipulated through flow control [58]. The channels are designed in such a way that various operations such as separation of phases, biological cells, the size-based separation of particles, to name a few, can be performed. However, fabrication of microfluidic devices may not be possible in low-income countries due to the lack of availability of materials. Even poly dimethyl siloxance (PDMS)-based soft lithography techniques require a master that can be expensive to make in low-income countries and set-ups such as mask makers and lithography may not be readily available. Paper is a low-cost alternative for sensors in the fight against cancer that may not need lithographic processing. Droplet-based paper devices could be highly useful for biomarker detection. Even microfluidic devices using screen-printing techniques can be printed on paper.

Paper-based sensors are devices that are fabricated on a paper substrate. These devices are printed on a cellulose-based paper substrate using readily available printers, making them easily accessible.

A typical paper-based sensor is based on exploiting the capability of paper to wick liquid, leading to capillary action in the paper. A typical paper-based sensor is developed on an hydrophilic paper substrate with wicking capabilities. Using surface treatment methods, a hydrophobic barrier is created in order to guide the flow of liquid through the specific path or channels.

The most commonly used paper in the fabrication of paper-based sensors is filter paper. Due to its pores, it has sufficient wicking capabilities, providing a moderate flow rate. Whatmann-branded filter paper manufactured by General Electric Health Care is the most widely used filter paper due to its uniform pore size and distribution. In applications where filter paper is not suitable, nitro-cellulose paper is used as a substrate. The main advantage of nitrocellulose substrate is its easy and efficient binding of proteins [59].

For example, nitrocellulose film is used for protein immobilization and filter paper is used for its water absorption. In one particular study for detection of bladder cancer, a glass-cellulose film was used for sampling, a nitro-cellulose film was used for protein immobilization, and filter paper was used for sample transfer due to its adsorption capabilities [60].

Table 1 highlights the different types of paper substrates that have been developed for paper-based sensing applications.

**Table 1.** Paper substrates of interest for paper-based microfluidics.

For patterning of the microfluidic channels on paper substrate, various printing techniques such as wax-printing, ink-jet printing, screen printing, lithography, plasma processing, and manual pattern drawing have been explored. In a typical printing-based fabrication process, a CAD model of the microfluidic channels is printed using wax/inkjet–printer on the substrate. Since the model is printed on only one side of the substrate, the substrate is heated to cause the reflow of the hydrophobic ink/wax barrier over the

complete cross-section of the substrate. These barriers act as microfluidic channels guiding the flow of the fluid (Figure 1).

**Figure 1.** Workflow of the fabrication process for printing microfluidic channels on a paper substrate.

Although paper-based microfluidic channels can be used for most of applications as conventional microfluidic chips, there may be concerns regarding the utility of paper substrate for use in microfluidic applications requiring multiple layers of fluidic channels for phase separation applications. For such applications Japanese paper, folding techniques such as origami and Kirigami are being explored [87].

The low cost of substrate, the ability to print continuously, the ability to make arrays of devices, and minimal capital requirements for the fabrication setup make paper-based sensors an interesting candidate for use in mass screening process. Paper being a naturally derived substance is biodegradable in nature. This will also reduce the environmental impact due mass production and usage of screening kits.

#### **3. Design and Working of a Typical Paper-Based Sensor**

The paper-based sensor used for screening must be fast, accurate, reliable, and must have a low limit of detection. A low limit of detection will allow for successful detection even during the early onset of cancer. A typical sensing setup comprises the following elements:

The paper-based microfluidic platform is divided into different zones, with each zone having a specific functionality, as shown in Figure 2. In the sampling zone, the sample is placed on the paper substrate. While passing through the microfluidic channels, labeling elements get attached to the analyte. In the detection zone, the analyte is detected and a signal is generated for the readout. In the case of colorimetry-based devices, there is a special zone termed as 'control zone'. In case a test yields a positive or negative result, there is a color change observed in the control zone.

**Figure 2.** Typical layout of a paper-based sensor with the different zones. Each zone performs a specific function.

#### **4. Analytes for Cancer Screening**

For cancer detection, biomarkers play an important role. According to the National Cancer Institute (NCI), biomarkers are defined as "a biological molecule found in blood, other body fluids, or tissues that is a sign of a normal or abnormal process, or of a condition or disease" [88]. Though biomarkers may be generated due to various factors such as somatic mutations, transcriptional changes, or post-translational modifications, they an important differentiators for an affected individual compared to a healthy individual [88]. A handful of biomarkers have been approved for cancer detection. For example, high levels of carcino embryogenic antigen may mean the presence of cancer. Similarly, CA 125 is a protein that is detected in blood for ovarian cancer. Lysophosphatidic acid, leptin, osteopotin, and insulin-like growth factor receptor 2 are used as a biomarker for ovarian carcinoma. Early prostate cancer antigen 2 is used as a novel biomarker for prostate cancer [89]. Biomarkers are not only useful for the detection of screening of cancer, they are also used in monitoring the effectiveness on any treatment or therapy. Various biomarkers are present in blood, saliva, urine, stools, etc., making it possible to obtain samples for analysis in a non-invasive or minimally invasive manner. There are various biomarkers that are used for cancer screening, including antigens, micro-RNAs, proteins, antibodies, and tumor cells. For example, the antibody–antigen interaction creates a signal that is measured qualitatively or quantitatively on a paper substrate.

#### **5. Recognition Element**

Recognition elements are responsible for the recognition of target analytes (ex: receptors) and their conversion into a signal, which can be qualitative, semi-quantitative, or quantitative [90]. An ideal recognition element has a highly specific binding affinity towards the analyte of interest [90]. Recognition elements can either be natural or artificial bio-molecules that are synthetically obtained [90–94]. When the target analyte molecule attaches to the recognition element, it undergoes a biochemical reaction producing a signal [90]. In some cases, for the detection of analyte, labeling agents such as nanoparticles are attached to the analyte molecule [90].

#### *5.1. Antibodies*

Antibodies are a type of natural bio-receptor that can be derived from living organisms [90]. Monoclonal antibodies are widely preferred in cancer screening applications due to their high specificity to the target antigen [95,96]. Through the development of hybridoma technology, it is possible to obtain a reliable and uniform supply of monoclonal antibodies [96]. This has also led to better reliability and accuracy of sensors using monoclonal antibodies. Covalent binding of antibodies to cellulose paper discs has been developed for colorimetric immunoassays. The antibodies were coated on the amine-functionalized cellulose paper discs. Through a glutaraldehyde cross-linking agent, the antibodies showed enhanced binding activity to the target when compared to the periodate oxidation method [97]. Other methods for antibody immobilization on a paper with shelf-life up to 12 months have been described [98,99]. Polyclonal antibodies have multiple binding sites, each specific to a particular antigen. Although they are cheaper to produce compared to monoclonal antibodies, they are not suitable for sensing applications due to the multiple binding sites [100].

For the detection of analytes, the principle of enzyme linked immunosorbent assay (ELISA) is applied in sensor design. Nanoparticles such as gold are used as antibody carriers and signal enhancers for ELISA. In the range between 0 and 60 U/mL, the ELISA assay adopting gold nanoparticles as an optical signal enhancer resulted in higher sensitivity and shorter assay time when compared to classical ELISA procedures. This was used to detect breast cancer biomarkers [101].

A sensor for the detection of prostate-specific antigens (PSA) used multiwalled carbon nanotubes (MWCNT) activated with anti-PSA antibody for the detection [102] . Due to the site-selective interaction between the antigen and the antibody there was a change in the resistance. This change in resistance could be measured using a benchtop multi-meter [102]. This method was found to be cheaper and faster than the ELISA method used in cancer diagnosis [102]. Similarly, single-wall carbon nanotube based biosensors have been used in the identification of cancer antibody–antigen interactions in blood samples using electrical conductance measurements. Following the measurements, a classification algorithm was implemented to differentiate between cancer and controls with 90% accuracy [103–105].

In a colorimetric sensor for the detection of pancreatic cancer biomarker (PEAK1), gold nanoparticles that were used as a labeling element acted as a color dye catalyst to produce colorimetric signals [106]. A photothermal-effect-based sensor used a graphene oxide (GO)-gold–anti-EpCAM antibody composite as the recognition element for the detection of MCF-7 cancer cells specific antigen [107]. After laser irradiation at the test zone, the temperature contrast was recorded for the detection of cell numbers (Figure 3) [107].

**Figure 3.** Working of photothermal effect based sensor for MCF-7 cancer cell detection. (Reproduced with permission from [107] Copyright 2016, Elsevier).

#### *5.2. Aptamers*

Aptamers are single-stranded nucleic acids that are folded into a specific architecture [108,109]. Due to their specific binding of the target proteins, they are used for sensing applications [108,110]. Their size and chemical stability make them widely preferred for the detection of proteins and small molecules [110,111]. Their low cost makes them a preferred alternative to antibodies in sensing applications [111]. For example, carbon-nanotube-based RNA apatamer sensors were developed for detecting IL6 in blood samples. Apatamer sensors based on field effect transistor arrays suggested a shift in drain current versus gate voltage for 1 pg and 1 ng of IL-6 exposure. The concentration of 1 pg falls below the diagnostic gray zone for cancer (2.3 pg–4 ng/mL), which is an indicator of early-stage cancer [112].

For the design of aptamer-based sensors, various strategies such as sandwich, targetinduced structure switching, or competitive replacement modes have been used for biosensor design [111]. Electrochemical sensing is the most preferred sensing method with aptamer as the recognition element; however, other methods such as optical sensing have also been explored [110].

In a fluorescence-based paper-based sensor designed for the detection of multiple types of cancer cells, graphene oxide-coated with mesoporus-silica-labelled high-specificity aptamers was used as a labeling element [113]. Using the excitation wavelength of 350 nm, a color change was produced that could be observed through naked eye [113].

#### **6. Sensing and Readout Methods**

For achieving higher utility of paper-based sensors, the readout method used should be cost-effective and portable. The method provides fast and accurate results without a requirement for extensive handling by experts.

In most of the cases, qualitative readout methods should suffice. However, with focus on providing health professionals with important data at the point-of-care, qualitative readout methods are also gaining significance. Although there are various readout methods for sensing applications, electrochemical and optical are the most widely used readout methods for paper-based sensing applications. With advancements in smartphone and machine learning technologies, there have been works that use smartphones for signal interpretation and readout.

#### *6.1. Modified Electrodes*

For electrochemical sensing, the potential difference between the electrodes is proportional to the concentration of the analyte. In paper-based sensors, the working electrode is modified such that the binding of the analyte produces an electrical signal through a change in resistance, current, capacitance, or impedance.

Various nanocomposites have been used for the fabrication of modified electrodes. These nanocomposites perform a dual function: recognition and amplification. Amino functional graphene (NH2-G)/thionine (Thi)/gold nanoparticles (AuNPs) nanocomposites are coated with recognition elements such as üimmobilized anti-CEA [114] and anti-NSE [115] for the detection of specific analytes. The sensor could provide fast results with a low limit of detection of 10 pg/mL [114]. In a more recent work, an aptasensor with two working electrodes capable of the simultaneous detection of CEA and NSE has been developed. Along with the NH2-G/Thi/Au nanocomposite, Prussian blue (PB)- poly (3,4- ethylenedioxythiophene) (PEDOT)- AuNPs nanocomposite was used for the fabrication of the second electrode, which was coated with immobilized CEA and NSE aptamers [116]. The device worked on the principle of electrochemluminescnce and could achieve fast and accurate detection of CEA and NSE with a limit of detection of 2 pg/mL and 10 pg/mL, respectively (Figure 4) [116].

**Figure 4.** Mechanics of modified electrode paper-based apta sensor. (Reproduced with permission from [116] Copyright 2019, Elsevier).

#### *6.2. Electrochemical*

In electrochemical sensing method, the analyte generates an electrical signal proportional to its concentration [90]. The signals may be generated through a biorecognition event, modified electrodes, or enzyme mediated electrodes [90,117]. For electrochemical detection, the sensor should have three electrode systems with reference, working, and counter electrodes. For measuring the signals, electrochemical devices such as electrochemical workstations or bench-top multimeters are used.

Various routes such as the use of labeling agents or modified electrodes may be used for generating electrical output from biological signals. In a sensor developed for the detection of cancer antigens, a marker for ovarian cancer, a reduced graphene oxide/gold nanoparticle/thionine nanocomposite was used as working electrode [118].

For the reliable detection of signals, signal amplification techniques are used. In paper-based sensors for the detection of CEA using horseradish peroxidase (HRP)–Ophenylenediamine–H2O2 as a detection element, graphene was coated on the substrate for accelerating the electron transfer and amplifying the signals [119].

#### *6.3. Optical*

For optical sensing, signals are generated through a recognition process by the formation of an antigen–antibody complex [90]. The optical signals could be fluorosence, chemiluminescence, or color change [90]. Other than the signals that display a direct color change, a photo-detector is used for measuring the signals [90].

Surface-enhanced Raman scattering is a popularly used method for signal detection in paper-based sensors. Gold nanostar@Raman reporter@silica-sandwiched nanoparticles have been developed as surface-enhanced Raman scattering (SERS) probes for the paperbased lateral flow strip (PLFS) assay [120]. A sensor for the detection of CEA used a portable raman sensor for measurement (Figure 5) [120]. Using a paper-based lateral flow strip capable of plasma separation and using silica nanoparticles for labeling the sensor displayed a limit of detection of 1 ng/mL [120].

**Figure 5.** Working of a paper-based sensor for CEA concentration detection using surface-enhanced Raman scattering (SERS). (Reproduced with permission from [120] Copyright 2021, American Chemical Society).

For naked eye detection, luminiscent reporters are used as labeling elements These can be nanoparticles [51], conjugated polyelectrolytes [121], or multi walled carbon nanotubes [102].

#### *6.4. Smartphone/Machine-Learning-Based*

Smartphones are devices that are readily available, even in low- and middle-income countries. Mobile health is becoming increasingly popular in developing countries [122]. It is widely explored as a tool for the efficient delivery of services, including in healthcare. Smartphones have been explored as a readout method for both optical and electrical signals [123–125]. For optical signals, a smartphone camera is used for data acquisition [126–128]. Using a custom application, the acquired image is compared with reference values and the result is calculated [126–130].

Smart phone-based imaging was used for calculating and displaying results in a multilayered paper-based sensor for cancer screening [131]. The movable layers allowed one to control the flow of the solution. Using the special design and smartphone-based readout, it was possible to achieve a low detection limit of 0.015 ng/mL[131].

Smartphones are also used for coupling with an electrochemical sensing device for the readout of signals [132]. A screen-printed sensor with multi-walled carbon nanotubes (MwCNT)/thionine (Thi)/gold nanoparticles (AuNPs) electrodes is capable of detecting cancer antigen (CA125) with a limit of detection of 2mU/mL [132]. The sensor uses a electrochemical detector powered using a smartphone, and it transfers data to the smartphone, where it is readout using a custom app (Figure 6) [132].

Table 2 summarizes recent works using paper-based sensors for cancer screening. It provides the breakdown of the sensor in terms of the biomarker(s) detected, recognition element, readout method used, and the types of cancer detected.

#### **Table 2.** Paper-based sensors for cancer screening.

**Figure 6.** Sensor for detection of cancer antigen (CA125). The screen-printed sensor used electrochemical deterrence for signal measurement and a smartphone with a custom application for readout (reproduced with permission from [132] Copyright 2022, Elsevier).

#### **7. Limitations of Current Paper-Based Methods**

Despite the promising future, there are currently several limitations that hinder the large-scale acceptance of paper-based sensors. Ranging from fabrication methods, a requirement of measurement equipment, to regulatory requirements, many limitations need to be addressed before paper sensors are actually put into service. Sensitivity and accuracy are also concerns in paper-based devices.

However, paper-based sensors are excellent for qualitative and semi-quantitative screening. One can further improve the accuracy of paper-based testing through implementing paper-based testing with artificial-intelligence-based analysis. Machine learning and deep learning can even predict the sequence of DNA and RNA that can point to cancer mutations. They could also potentially detect cancer cells in blood droplets. The ability to differentiate between cancer versus normal cells in blood based on AI can be a very powerful approach in making paper-based testing a reality for mass screening [152].

With the exception of laser printing and screen printing, different fabrication methods used for the fabrication of paper-based microfluidic devices such as photo-lithography, e-beam lithography, reactive ion etching, and metal or oxide deposition require extensive

capital investment and training, making them non-feasible for use in low- and middleincome countries.

For devices with qualitative measurement of signals, despite the low cost of single paper-based sensing device, expensive equipment such as electrochemical workstations, photo detectors, or electrochemiluminscence detectors are required for a results readout. This not only affects the portability of the device but also increases the overall cost of the screening, and it requires a trained technician to carry out the readout.

Nevertheless, paper-based sensors can make a significant impact in terms of testing blood for infectious diseases and even in the fight against cancer. The qualitative assessment of whether a person has cancer based on biomarkers in blood can reduce the cancer clinical burden in low- and middle-income countries. The paper-based testing method will definitely have an advantage here to reduce the clinical burden through the large-scale screening of populations. People with cancer undergoing chemotherapy may be prone to infectious diseases. Life-threatening infectious diseases can kill people in a few days. Here, POC systems based on paper and colorimetric detection methods for parasites, viruses, and other agents would be highly valuable. Before vaccination, people, including doctors, were dying of COVID-19 within a few days to two weeks. Low-cost qualitative paper-based sensors with instant results through color change are an absolute necessity to differentiate between population who has the virus from those who do not. This may be cheaper than the current PCR test that is used and is done in the laboratory. In the era of the pandemic, low-cost sensors for personal safety are very important. Qualitative paper-based sensors with instant read out will be highly useful in such pandemics as one cannot run tests in the laboratory frequently.

#### **8. Conclusions**

Paper-based sensors have immense potential to to act as low-cost tools in the mass screening of cancer. They can be used in cancer-screening camps, especially in low- and middle-income countries. Due to the health care resources being highly stressed in these countries, screening using paper-based sensors will act as a filter. The samples of the patients who test positive in the screening stages can then be treated as a priority. This will in a way help in resource allocation and management in these countries. It must be mentioned that the paper-based sensors with their current state of the art are insufficient for providing data to healthcare professionals in making important decisions. Further, testing will be required to ascertain the stage of the disease before any treatment plan is decided. Many times, the same biomarker is produced for multiple types of cancers. CEA is a common marker for multiple types of cancer such as lung, breast, ovaries, stomach, and intestine, to name a few. Thus, it is necessary to determine the type of cancer and the stage of disease progression before starting the treatment. Normal CEA levels are 2.5 ng/mL. A CEA level of 10 ng/mL would indicate the presence of cancer, and anything above 20 ng/mL would indicate the spread of cancer.

Patients with continuously decreasing levels of CEA do better after treatment than patients with increasing CEA levels. A regular paper-based CEA test with an instant read out can tell the doctors the potential for cancer progression. Their use will reduce the number of expensive tests (CT scans, PET-scans, etc.) a patient undergoes during follow-up visits and also help reduce the cost of treatment. In the era of telemedicine, it is becoming increasingly convenient to deliver healthcare at home. Cancer-screening tests can be conducted at home, and the results of the test can be emailed or texted to a doctor automatically using smart phones. So, paper-based testing is important here for home-based low-cost sensors.

Although paper-based sensors may not potentially be the knight in shining armor against humanity's fight against cancer, they can potentially be the important foot soldier in the fight. Future integrated paper-based sensors where all the sensing and electronic circuitry are printed in paper can make an impact on low-cost testing. Paper-based cantilevers with optical sensors and electronics integrated in a hand-held chip could enable the

detection of cancer biomarkers such as prostate-specific antigen (PSA) from blood samples. Thus, there are exciting opportunities for paper-based sensors in the fight against cancer. The places where paper-based sensors along with mobile phones and AI-based techniques could make an impact in the fight against cancer are (1) the low-cost qualitative screening of large populations; (2) reducing the clinical burden through proper resource allocation; (3) estimating cancer prognosis; (4) monitoring cancer treatment; and (5) detecting cancer recurrence qualitatively or semi-quantitatively. With the ability to miniaturize anything, from detectors to spectrometers, one can implement miniaturized low-cost detectors along with paper-based sensors for cancer detection. Such miniaturized sensors are already made by many companies and could be bought off the shelf and integrated with a paper-based sensor. Finally, paper-based methods with artificial intelligence techniques can enable low cost and further improve the sensitivity and accuracy of paper-based sensors.

**Author Contributions:** Conceptualization, A.D. and A.J.N.; writing—original draft preparation, A.D.; writing—review and editing, A.D., A.J.N. and B.P.; and supervision, A.J.N., B.P., and T.-H.O. All authors have read and agreed to the published version of the manuscript.

**Funding:** B.P. acknowledges funding from the United States-India Educational Foundation through the Fulbright-Nehru Academic and Research Professional Excellence Award, Grant Number: 2021/ APE-R/63. A.J.N. acknowledges the financial support from the Department of Biotechnology, Government of India, through Ramalingaswami Re-entry fellowship (D.O. No. BT/HRD/35/02/2006) and VIT SEED GRANT 2021-2022 (SG20210234).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** A.J.N. would like to acknowledge the financialsupport from the Department of Biotechnology, Government of India, through Ramalingaswami Re-entry fellowship (D.O. No. BT/HRD/35/02/2006) and VIT SEED GRANT 2021-2022 (SG20210234). B.P. acknowledges funding from the United States-India Educational Foundation through the Fulbright-Nehru Academic and Research Professional Excellence Award, Grant Number: 2021/APE-R/63.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **Abbreviations**

The following abbreviations are used in this manuscript:

VEGF-C Vascular endothelial growth factor C NMB New methylene blue NH2-SWCNTs Amino-functional single-walled carbon nanotubes AuNPs Gold nanoparticles

#### **References**

### *Review* **Recent Advances in Microfluidic Platform for Physical and Immunological Detection and Capture of Circulating Tumor Cells**

**Mahesh Padmalaya Bhat 1,2, Venkatachalam Thendral 1, Uluvangada Thammaiah Uthappa 3, Kyeong-Hwan Lee 2,4, Madhuprasad Kigga 1, Tariq Altalhi 5, Mahaveer D. Kurkuri 1,\* and Krishna Kant 6,\***

**Abstract:** CTCs (circulating tumor cells) are well-known for their use in clinical trials for tumor diagnosis. Capturing and isolating these CTCs from whole blood samples has enormous benefits in cancer diagnosis and treatment. In general, various approaches are being used to separate malignant cells, including immunomagnets, macroscale filters, centrifuges, dielectrophoresis, and immunological approaches. These procedures, on the other hand, are time-consuming and necessitate multiple high-level operational protocols. In addition, considering their low efficiency and throughput, the processes of capturing and isolating CTCs face tremendous challenges. Meanwhile, recent advances in microfluidic devices promise unprecedented advantages for capturing and isolating CTCs with greater efficiency, sensitivity, selectivity and accuracy. In this regard, this review article focuses primarily on the various fabrication methodologies involved in microfluidic devices and techniques specifically used to capture and isolate CTCs using various physical and biological methods as well as their conceptual ideas, advantages and disadvantages.

**Keywords:** circulating tumor cells (CTCs); microfluidic device; physical method; biological method; cancer diagnostics

#### **1. Introduction**

Cancer is defined as the uncontrolled proliferation of aberrant cells in the human body, and it is classified into two types: benign and malignant cancers. A benign tumor that grows slowly and has no negative effects on the human body. Malignant tumors, on the other hand, are aggressive, grow quickly, spread rapidly and eventually kill the patient. During metastasis, some tumor cells at the primary tumor's borders undergo a process known as epithelial-mesenchymal transition (EMT), in which the cells lose their epithelial traits and gain migratory mesenchyme properties [1]. These migratory tumor cells enter adjacent arteries and start travelling along with red and white blood cells throughout the body. CTCs enter the bloodstream through the vasculature and circulate alongside healthy hematological cells before metastasis [2,3]. However, these can only be diagnosed if the patient has progressed to the metastatic stage [4]. These CTCs stop internally at

**Citation:** Bhat, M.P.; Thendral, V.; Uthappa, U.T.; Lee, K.-H.; Kigga, M.; Altalhi, T.; Kurkuri, M.D.; Kant, K. Recent Advances in Microfluidic Platform for Physical and Immunological Detection and Capture of Circulating Tumor Cells. *Biosensors* **2022**, *12*, 220. https:// doi.org/10.3390/bios12040220

Received: 11 March 2022 Accepted: 4 April 2022 Published: 7 April 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

some organs and trigger secondary tumors; from this stage onwards, the cancer enters its deadliest form, and the patient could face fatal consequences [5,6]. Hence, the early detection of these cells or the monitoring of their presence in the bloodstream is required and important for the accurate diagnosis and prognosis of cancer [7]. A survey has shown that malignant tumors will be the major cause of death worldwide by 2030, expected to grow to 20.3 million new cancer cases and 13.2 million deaths [8].

However, CTCs are extremely rare among hematological cells. There are only a few CTCs in a 1.0 mL blood sample, where nearly 5 billion red blood cells (RBCs) and 10 million white blood cells (WBCs) are present. In addition, the CTCs may exist in a single-cell or cluster form, with varied phenotypic properties. Based on the changes in protein expression on CTCs, they can be classified into epithelial-mesenchymal, epithelial, and mesenchymal types [9]. Therefore, collecting and isolating them from other components in the bloodstream is quite difficult and challenging [10]. Detection of these rare cells using sensors would be beneficial. Sensors have previously been used for environmental applications [11–17]. On the other hand, sensors would be ideal for the detection of these rare CTCs. Currently, several techniques such as flow cytometry, enzyme-linked immunosorbent assay (ELISA), Western blotting, quantitative polymerase chain reaction (Q-PCR), magnetic-activated cell sorting (MACS), fluorescence-activated cell sorting (FACS) and centrifugation techniques, and laserbased technology are widely used for the biomolecular or cellular analysis of cancer [18–26]. Although these techniques have several limitations, such as substantial sample consumption, low throughput, lack of real-time monitoring, and high overall operational expenses, there are no other alternative simple techniques available for CTC isolation. As a result, there is a great scientific desire to improve cancer diagnosis using low-cost procedures [27].

In our opinion, microfluidic devices are one of the most intriguing methods for capturing and isolating CTCs from blood samples. Microfluidic devices have many advantages, including their high throughput, low cost, miniaturization, quick analysis, high sensitivity, precise operation, high efficiency, portability, low sample consumption, and accuracy [28–34]. As the name implies, microfluidics is concerned with accurate fluid flow management in microliters (10–6) to picoliters (10–12) within micro-volume channels [35]. Various techniques like 3D printing [36], molding, laminating, and high-resolution nanofabrication are used to create these devices. S.C. Terry reported the first lab-on-a-chip (LOC) analysis system in 1979, which was investigated for gas chromatography applications [37]. Since then, microfluidic devices have been investigated for a variety of applications, including biosensors [38], separation [39], analysis [40], drug delivery [41,42], optoelectronics [43], cell manipulation [44], and chemical synthesis [45,46]. There has been much advancement in surface chemistry, which has enabled the development of smart surfaces and devices for various applications [47,48]. In comparison to other approaches, microfluidic channels have a high surface-to-volume ratio [49]. Microfluidic devices are usually made from polymers such polyethylene glycol diacrylate (PEGDA) [50], parylene [51], and polydimethylsiloxane (PDMS) [52,53]. There are two types of microfluidic technologies for capturing and isolating CTCs: physical and biological methods [54]. The different intrinsic features of cell populations, including their density [55], size [56], compressibility [57], deformability [58], dielectric properties [59], and viscosity [60], are used to physically separate CTCs [61,62]. Deterministic lateral displacement, inertial microfluidics, micropores, micropillar arrays, vortex-mediated deformability cytometry (VDC), inertial focusing dielectrophoresis, acoustic waves, and optical approaches have all been reported for the detection and separation of CTCs [63–69]. Biological approaches, on the other hand, rely on specific surface proteins produced on tumor cells to act as molecular recognizers such as transferrin, peptides, sialic acid, and antibodies to trap and isolate CTCs [70–72]. The two primary kinds of biological techniques are positive and negative sorting. The epithelial cell adhesion molecule (EpCAM) is a one-of-a-kind biomarker for positive sorting, which uses CTCs as target cells. Negative sorting, on the other hand, uses CD1513, anti-CD6647, and anti-CD45 biomarkers to identify leukocytes as target cells. Physical approaches are simple to use and do not require expensive biomarkers or a long incubation period. Nonetheless, they lack specificity and isolation

purity. Biological approaches, on the other hand, need a more involved, time-consuming, and costly procedure. Yet, they have a high level of specificity, purity, and efficiency [73]. As a result, a physical-based approach is one of the most effective and straightforward methods for capturing and isolating CTCs. In this review, we have highlighted and critically examined the recent relevant literature on the fabrication of microfluidic devices for CTC isolation and the most promising elements of CTC capture and isolation, employing innovative microfluidic devices such as physical and biological techniques. The general technologies involved in the physical and biological separation of CTCs are depicted in Figure 1.

**Figure 1.** Range of methods involved in physical and biological approaches for early-stage detection and isolation of CTCs.

#### **2. Fabrication of Microfluidic Devices for the Isolation of CTCs**

In a very short span of time, microfluidics has emerged in several technological advancements. There are a variety of materials for microfluidic device fabrication, each with different properties according to the requirements. Based on the required specific characteristics of the fabrication material and product requirements, different techniques are used for the development of the device. Another major aspect is the cost of the involved material. In most cases, used devices are disposed of. Thus, the method involved should be economically feasible. Herein, we have classified the most recent techniques adapted in the fabrication of microfluidic devices for the isolation of CTCs.

#### *2.1. Additive Manufacturing*

Molding techniques involving PDMS and other thermoplastics are the most common approaches to fabricating microfluidic devices [74]. The disadvantages of traditional fabrication approaches are that they require a cleanroom, are expensive, utilize timeconsuming wafer processes, and require the labor-intensive manual assembly of multiple layers. These factors have limited their wide application [75,76]. Furthermore, it is difficult to efficiently fabricate true 3D structures with large surface areas to increase CTC capture efficiency [77,78]. In recent years, 3D printing, which can create 3D objects layer by layer, has received a lot of attention as a potential replacement for the PDMS-based conventional molding process. In the additive manufacturing (AM) approach, the device is fabricated using a 3D printer and computer-aided design (CAD) software to design the desired shape in a short amount of time. Chu et al. created monolithic microfluidic devices to separate CTCs from whole blood samples [79] (Figure 2a). The fabricated device has a 100 mm channel length, 20.5 mm breadth and 19.2 mm width. The microfluidic device is comprised of two inlets for a sample, a buffer, and an outlet for collecting the waste. The main advantages of this device are that during the filtration process, potential cell damage

due to handling the sample was eliminated, and the desired pore size could be attained with high resolution in commercially available membrane filters. Further, Gong et al. developed controlled-compression integrated microgaskets (CCIMs) and simple integrated microgaskets (SIMs), which are bound with small chips to form a wider connection of chips accomplished by a microelectromechanical system (MEMS) and nanoelectromechanical system (NEMS) [80]. SIMs or CCIMs are 3D printed as part of the device's fabrication. Thus, no additional materials or components are needed to connect to the larger 3D-printed interface chip. Later, Chen et al. developed a microfluidic device with 3D-printed internal structures to facilitate high fluid flow and surface area [81]. The printed structure was functionalised with EpCAM antibodies to capture CTCs.

#### *2.2. Etching Technique*

Etching is the process of protecting the desired area of a substrate while treating the other in order to remove a particular depth of material. The parts that we do not want to etch are usually protected. Liu et al. used wet etching and thermal bonding to create a pyramid-shaped microfluidic device with one inlet and six outlets [82]. The microchamber is a critical functional component of microfluidic devices for CTC separation. A layer of chemical-corrosion-resistant adhesive tape was pasted on a standard glass slide, and a laser ablation system was used to transfer the desired prototype onto the adhesive tape. The first round of tape was then peeled off, and the glass slide with patterned tape was immersed in the etching solution for 25 min at an etch rate of 1 μm/min. The second and third annular tapes were peeled off, and the glass slide with patterned tapes was dipped in the etching solution for 7 and 8 min, respectively. After the device was completed, a laser was used to punch one inlet and six outlets to allow the blood samples to flow. Each outlet was located on a different layer at different heights of the microfluidic device. The first, second, and third steps were respectively 40, 15, and 4~8 μm high. The device showed a throughput of ~99%. The device has the advantages of being simple to set up, having high isolation efficiency, demonstrating improved throughput and not requiring an expensive capture reagent. Further, Yang et al. reported a wet etching and thermal bonding process to create a unique, low-cost, wedge-shaped microfluidic device made of two glass pieces with appropriate specificity and sensitivity [83]. The device is comprised of two inlets, a linear reservoir, and an outlet. After coating a standard glass slide with a chemical-corrosionresistant adhesive tape, the laser ablation system was used to transfer the microchannel design onto the adhesive tape. The glass slide coated with patterned tape was immersed in a glass etching solution to create a microchannel with a continuously decreasing height (from 60 to 5 μm). Then, two inlets and an outlet (0.5 mm in diameter) were drilled on the glass slide to obtain the final chip. After a dynamic heating and annealing process in a programmable muffle furnace, the two glass slides were bonded together.

#### *2.3. Mold Punching Technique*

The fabrication of microstructures via conventional techniques can be costly due to the need for expensive equipment set up and maintenance and the time-consuming nature of the process. If micro- or nano-scale processes can be replicated, manufacturing costs can be drastically lowered. In this technique, micro/nanostructure molds are fabricated once, and products can be duplicated from them. The inverted or negative aspects of the device construction are present in the masters [84]. Liao et al. created an optically induced dielectrophoresis (ODEP) microfluidic device with a T-shaped microchannel made up of four layers: layer A (PDMS), layer B (indium-tin-oxide glass substrate), layer C (doublesided adhesive), and layer D (indium-tin-oxide glass substrate coated with photoconductive material) for the isolation of CTCs using EpCAM/CD45 markers [85]. To facilitate cell suspension transfer, the main channel and side channel's dimensions (L × W × H) were set to 2500 × 1000 × 60 μm and 2500 × 400 × 60 μm, respectively. The junction area in the T-shaped microchannel that was specified for CTC separation was 1400 × 1000 × 60 μm. The device consisted of three punch holes for tubing connections, with each hole used for

loading the sample, harvesting the fresh, waste cell suspension samples, and collecting the separated cells. The advantages of this device included the fact that the cell manipulation process was simpler and easy to operate.

#### *2.4. Photolithography Technique*

Photolithography has been widely used in the fabrication of microfluidic devices. It entails exposing a photoresist-coated substrate to light so that the selectively developed regions can be protected from/subjected to subsequent fabrication processes like etching or deposition [86,87]. This process, however, necessitates the use of costly photolithographic facilities with specialized lighting for working with ultraviolet (UV)-sensitive materials [88] and uses light-sensitive photoresist to transfer a geometric design from a photomask to a smooth surface. On a glass slide, Kwak et al. reported a spiral-shaped channel microfluidic device [89]. Each circular channel measures 250 μm in width and has a gap between them, with a channel depth of 130 μm. The distance between the spiral channel and the magnet (i.e., radius) was reduced from 3500 μm to 500 μm. High throughput and selectivity are two advantages of this design (Figure 2b). Further, Fan et al., on the other hand, devised a novel size-based separation approach for the rapid identification and isolation of CTCs [90]. The authors created a microfluidic device based on a polydimethylsiloxane (PDMS) membrane filter. The device had a thickness of 60 μm, a diameter of 6.9–10.8 μm, and a gap of 25 μm between two holes. The microfilter produced using lithography has several advantages, including precise, uniformly dispersed pores, high porosity, low cost, and quick processing. However, this method is not suitable for mass production. Later, Yan et al. fabricated an electrochemical microchip for high-efficiency CTC isolation to address the limitations of prior efforts [91]. The PDMS micropillar-array-based electrochemical microchip had hierarchical structures spanning from μm to nm, which were created using a traditional soft lithography approach and then gold layer plating for the electrochemical capture and lysing of captured cells. Similarly, Zhou et al. created a PDMS-based multi-flow microfluidic system using dry film resist instead of SU-8, followed by soft photolithography [62]. The developed straight channel had a length, width, and height of 20 mm, 150 μm, and 50 μm on PDMS, which was bound to a glass slide followed by plasma treatment.

Kulasinghe et al. designed a multi-flow straight microchannel of 50 μm height and 150 μm width, with two inputs and two outputs for inertial cell migration [92]. The device uses size-dependent separation from the inertial movement of a mixture of WBCs and CTCs, allowing for the isolation of larger CTC clusters as the channel length increases. The sample was injected through the outer inlet, while the phosphate-buffered saline was injected through the inner inlet (PBS). Cells migrated transversely from the sample zone into the clean buffer flow channel as a result of inertial force. Yoon et al. designed a 4.5 × 4.5 cm2 microfluidic device on a surface-oxidized silicon chip [93]. The device has two inlets for the sample and the buffer, which are followed by two outlets for waste and isolated cells. The main channel measured 500 μm in length. The slanted weir runs from the upper side of the main channel wall to the branch point. The height of the slanted weir was 7 μm lower than the height of the main channel. A double-layer photolithographic technique was used to pattern the slanted weir-integrated microfluidic channel. Initially, the first layer was spin-coated with a thickness of 23 μm using negative photoresist SU-8 2050, and the second layer with a weir gap was spin-coated with a thickness of 7 μm using negative photoresist SU-8 2007 to get the expected slanted weir-designed device. In other work, Chen et al. fabricated a PDMS-based microfluidic design consisting of gallium nitride (GaN) and aluminium gallium nitride (AlGaN) layers integrated with a field-effect-transistor (FET) chip of 1.2 × 0.8 mm by the plasma etching and metal deposition method followed by a molecular beam epitaxy process [94]. Photoresist SU-8 was spin-coated on a silicon wafer with a thickness of 30 μm; the length, width and height of the channel were set to 60, 20, and 30 μm, respectively. The upper layer of the device was composed of two inlets for cells and buffered saline with four trapping microchambers for cell capturing. The bottom layer was embedded with an FET sensor array on the epoxy substrate. Raillon et al. printed a circuit board coated with a

positive photoresist to fabricate a label-free PDMS microfluidic device for the isolation and enumeration of CTCs from human blood samples [95]. At first, a glass wafer was coated with lift-off resist and positive photoresist, followed by printing electrodes using a laser writer to achieve a glass chip with electrodes. Secondly, an SU-8 mold was used to develop a PDMS chip using standard photolithography. The glass chip and PDMS chip were combined to form a single PDMS impedance chip. The channel dimensions were 70, 16, and 40 μm in depth, length, and width, respectively. The device consisted of a plastic vortex chip and PDMS impedance chip, which were connected for fluidic flow. Syringe pumps were used for the injection of the sample into the channel. The flow rate was optimized for the vortex chip at 7 mL/min and reduced to 100 μL/min for the impedance chip. Further, captured CTCs were flushed out with an increased buffer flow rate of 8 mL/min. An excitation voltage was applied between two electrodes at 500 mV and 460 kHz frequency with a flow rate of 100 μL/min, 10 kHz bandwidth, and 100 kHz sampling frequency to detect cancer cells. The advantages of the specific electrode design chip included its high-frequency measurements, ease of fabrication, and fast particle counting (Figure 2c).

**Figure 2.** Microfluidic device designs fabricated using various techniques. (**a**) 3D-printed device showing the microchannels with layers and the filter holder; reprinted with permission from ref. [79], 2019, Royal Society of Chemistry. (**b**) Microfluidic device fabricated by photolithography showing spiral channel and cell trapping segments; reprinted with permission from ref. [89], 2018, Elsevier. (**c**) Representation of label-free enumeration of CTCs using a vortex chip connected to an impedance cytometry chip; reprinted with permission from ref. [95], 2019, John Wiley and Sons. (**d**) Schematic of the detection strategy of the micro-aperture chip system for CTC detection; reprinted with permission from ref. [96], 2015, Royal Society of Chemistry.

Similarly, Chen et al. fabricated a PDMS-based hybrid magnet-deformability CTC chip patterned through a photolithographic technique [97]. The thickness of the silicon wafer post-spin coat was 7 μm, where the CTCs were isolated using a magnetic force. The 12 rows of micro-elliptical pillars were designed within the channel. The distance between adjacent micropillars was gradually reduced from 18 to 5 μm for effective CTC removal, while the width between adjacent arrays remained constant at 1500 μm. The presence of a magnet beneath the device aided in increased the capturing efficiency. The micro-ellipse was comprised of three parts, which include a half-ellipse with a semi-long axis of 30 μm, a half-circle with a radius of 15 μm, a rectangle with a length of 30 μm and a device with a depth of 55 μm. Furthermore, Varillas et al. developed a PDMS-based geometrically enhanced mixing (GEM) microfluidic chip with two layers of SU-8 coating (main channel layer and herringbone mixer layer) for the isolation of CTCs using EpCAM antibodies [98]. For the main channel, the thickness of the SU-8 2035 photoresist was 50 μm. The herringbone mixer layer was formed by adding a second layer of SU-8 after UV light exposure and post-soft baking. A precise arrangement between the main channel and the mixer was maintained to create the herringbone mixer pattern. The inlet and outlet

wells were created by punching the holes in PDMS after a second exposure was performed. Shamloo et al. fabricated a new integrated Y-shaped microfluidic device consisting of two subunits, a functional unit and a mixing unit, through SU-8 photoresist patterning and a wet etching process for the immunomagnetic separation of CTCs [99]. The blood samples spiked with CTCs were passed through a 500 μm wide inlet channel. The functional unit and mixing unit had dimensions (L × W) of 12 × 4 mm and 9 × 1 mm, respectively, in which the channel was subjected to an alternative voltage by 10 electrodes arranged in a zigzag pattern. The sample flowed for 3 mm before reaching the diverging region, where it extended for 7 mm towards the outlets. Non-tagged cells were collected through the upper outlet, while magnetic-particle-tagged cells were isolated through the lower outlets, which had a magnet beneath them. The important features of the device were its simple geometry, high efficiency, and high feasibility. However, it was lacking in high performance.

Chang et al. used a silicon fabrication process to create a PDMS-based microfluidic chip to capture CTCs [96] (Figure 2d). The device was made up of 8 microchips with dimensions (L × W) of 40 × 20 mm. Each chip had a 9 mm by 3 mm porous area in the center, with a pore area thickness of 50 μm. These microchips were covered with a 1 mm thick glass slide. A PDMS layer of ~2 mm thickness was used as a spacer between the glass slide and microchips to form the fluidic chamber. The dimension of the fluidic chamber was defined by a laser cutter with a 30 mm by 3.8 mm grove. The entire setup was placed on an acrylic stand where a magnet was placed. The inlet and outlet were connected to the sample source and peristaltic pump, respectively. This parallel flow micro-aperture chip system has several advantages, including compatibility, ease of use, and the ability to reuse the chip for cell analysis. Later, Chen et al. used soft lithography to create a microfluidic device with a microwell-structured array for the analysis and isolation of targeted tumor cells [100]. The length and depth of the channel were 10 mm and 60 μm, respectively and the width of the chamber was 2.3 mm. The depth and diameter of four various-sized microwell structures were 5.0 μm/18 μm, 5.0 μm/20 μm, 5.0 μm/22 μm, and 8.0 μm/20 μm, respectively, with excellent selectivity for CTCs. Hoshino et al. designed a PDMS-based immunomagnetic microchip for the capture of CTCs from spiked cultured cancer cell lines by magnetic nanoparticles (MNPs) functionalized with EpCAM antibody [101]. UV-patterned SU8-photoresist coated on the silicon wafer was used as a master. The developed microchannel on PDMS was bonded on a glass substrate with a thickness of 150 μm. The developed microchannel measured 30 mm in length, 20 mm in width, and 500 μm in height. Fallahi et al. used photolithography to create a stretchable, flexible microfluidic device for the size-based separation of CTCs [102]. The channel dimensions of the developed device were 100 mm, 100 μm, and 45 μm in length, width, and height, respectively. There were sample and buffer flow inlets as well as waste, large-cell outlets, and small-cell outlets. The entire chip was placed on a specially designed stretching platform (Figure 3a). However, when compared to other size-based microfluidic separation techniques, the device setup was complicated. Further, Jiang et al. demonstrated the use of microbubbles to extract CTCs in a label-free, high-throughput acoustic microstreaming technique [103]. SU-8 2075 photoresist and soft lithography were used to construct the device on a 4-inch silicon wafer. The system was made up of 101 pairs of lateral cavity acoustic transducers (LCATs), each with one inlet and two outlets. The device had a width of 750 μm and was mounted on a piezoelectric transducer with ultrasonic gel between them. The isolation of CTCs by LCATs depended on the oscillation of trapped microbubbles in lateral slanted dead-end side channels to generate a first-order oscillatory flow at the air–liquid interface followed by a second-order streaming flow that consisted of an open microstreaming flow and a closed-looped microstreaming vortex. The dead-end of the channel was tilted at 15◦ to allow bulk flow through the microstreaming. The narrow gap in the flow area between the looped microstreaming vortex and the air–liquid interface was controlled by the voltage, which regulated the particle size that flowed through. Cells that were smaller than the gap moved forward along the flow by trapping large CTCs. This method allows rapid isolation with the potential to isolate multiple types of CTCs.

Furthermore, Jou et al. used photolithography to create a silicon-based V-BioChip with nano-pillar arrays with a chip dimension (L × W × H) of 32 × 34 × 0.7 mm [104]. A metal-assisted chemical etching technique was used to create nano-pillars within the microchamber. The chip surface was coated with a layer of polyethylene glycol-biotin (PEG-biotin) using a vapor deposition method. Streptavidin was attached to the biotin end using a liquid deposition method to improve capture efficiency. The surface-modified chip with nano-pillars promotes antigen-antibody interaction between the surface and CTCs, resulting in cell capture. Furthermore, Zhang et al. created a label-free microfluidic device for isolating CTCs from breast cancer patients' blood samples [105]. The photolithography technique was used to develop microchannels to fabricate the microfluidic device. An inlet, a cell intercept area and an outlet were present on the chip. The impurities were filtered through two layers of hexagonal columns in the microchannels. The first and second narrow channels were 50 μm and 20 μm long, respectively, with cell filtration occurring in 30 main channels and 31 side channels. The channels consist of 40 μm cylindrical wells separated by a 100 μm separation distance. Reinholt et al. created a PDMS microfluidic device using photolithography to isolate CTCs using aptamer and extract and to amplify DNA for gene mutation analysis [106]. The device consists of two orthogonal microchannels with two micropillar arrays for CTC isolation at the intersection of the two microchannels and the genomic DNA isolation array downstream of the cell capture array. The cell channel was 1 mm wide, whereas the DNA channel was 500 μm to 1 mm wide and 25 μm deep. Micropillars with a diameter of 50 μm made up the cell capture array. The DNA micropillar array was spaced in a gradient starting at 10 μm and ending at 7 μm. Nasiri et al. developed a hybrid PDMS microfluidic device for CTC isolation via inertial and magnetic separation [107]. For the isolation of CTCs from blood samples, the device consists of an asymmetric serpentine inertial channel, an inertial focusing channel and magnetic cell separation zones. The dimension of the inertial channel was set to 400 μm in width and 80 μm in height, followed by a magnetic separator channel width of 650 μm (Figure 3b).

**Figure 3.** Microfluidic device designs fabricated using various techniques. (**a**) Schematic of the working setup of the stretchable microfluidic device, with an inset showing the multi-flow microchannel with a real stretchable microfluidic device; reprinted with permission from ref. [102], 2021, Royal Society of Chemistry. (**b**) Schematic of hybrid microfluidic cell separation device showing CTC sorting by an inertial focusing microchannel followed by magnetic separation [107]. (**c**) Photomasks fabricated using offset printing showed better resolution and smooth surface over other laser printing techniques [108]. (**d**) Schematic illustration of the optically induced dielectrophoresis (ODEP) microfluidic system assembly where Layer A was composed of fabricated polydimethylsiloxane (PDMS) components; Layer B was composed of indium-tin-oxide (ITO) glass; Layer C was composed of double-sided adhesive tape with microfabricated microchannels; and Layer D was composed of ITO glass substrate coated with a layer of photoconductive material [85].

#### *2.5. Printing Technique*

Despite the fact that 3D printing is a cheap, robust and scalable method for producing master molds [109,110], there are still challenges that have prevented microfluidic developers from adopting 3D printing, including resolution, throughput and resin biocompatibility [111]. Attempts to reduce the cost of the technique have focused on UV lighting, laser/offset printing, etc. [112–114] Laser printing and offset printing could be cost-effective alternatives to expensive photolithography technology. Nguyen et al. investigated methacrylate (MA) gel, a type of nail polish that has been shown to work as a photoresist material instead of SU-8, to develop a master mold with additional benefits such as low cost, rapid production, high resolution (100 μm thickness, 100 μm feature size), high accuracy, and reproducibility [108] (Figure 3c). They used laser and offset printing techniques for photomask generation. The fabricated microfluidic device had a diameter of 100 μm and a height of up to 1 mm. They devised a cost-effective method for fabricating microfluidic devices. To save money, standard procedures like spin coating, plasma etching, and aligners were kept out of the device fabrication. Xu et al. created a microfluidic device out of polymethyl methacrylate (PMMA) by using a laser engraving machine to create microchannels on the surface for CTC isolation [115]. The device was divided into two major components: a filtration system on top and a magnetic microfluidic chip at the bottom. The filtration system used a micropore array membrane to isolate CTCs before filtering out the waste cells. The filter membrane measured 20 × 20 mm and had a pore size of 10 μm. The CTCs with trace WBCs were rinsed off the membrane after filtration for further purification. The magnetic microfluidic chip with a magnetic base of 70 mm with a diameter of 50 mm was used for the negative sorting of CTCs. This device showed low capture efficiency and needed two steps for the isolation of CTCs. Recently, Gurudatt et al. fabricated an electrochemical microfluidic channel modified with conducting polymers by a screen printing approach using carbon ink on a glass slide [116]. The developed microchannel exhibited a width and height of 95 ± 2.5 μm and 15 μm, respectively. The screen-printed channel was dried at 60 ◦C for two days. Further, the channels were covered using a glass slide. Later, for the amplification of separation, the channel wall was modified with a DAT monomer to covalently attach lipids. Further, Nieto et al. fabricated microchips with pillars on a soda-lime glass substrate using a laser-direct writing technique followed by thermal treatment [25]. An aluminum film was placed on the rear side of the soda-lime glass to increase the ablation. A cylindrical array of micro-posts with 420 μm diameter with a pitch and depth of 245 μm was formed. Further, the pillars were functionalised with EpCAM to facilitate CTC isolation.

#### *2.6. Overall Summary of the Fabrication Process*

Several fabrication methods have been discussed, each with its own set of characteristics. One must know the minimum feature sizes that the above approaches can produce, as well as a variety of other criteria such as surface roughness, aspect ratio and normal working size, in order to get benefit from the available techniques. Factors such as fluidic outcomes, pressure drop, microchannel, and process time play a major role in the development of the device. Though there are several techniques available for the fabrication of microfluidic devices, photolithography-based devices are determined to be promising in terms of channel dimension precision based on the aforesaid results. The technology, however, can only be used for two-dimensional devices. Additive manufacturing, on the other hand, is cutting-edge, with the potential to create three-dimensional monolithic devices. The main disadvantage is that they are not precise enough for micrometer channels. Therefore, in the near future, 3D printing technology could overcome the challenges and replace the traditional photolithography process for fabricating microfluidic devices. In general, microfluidic devices are fabricated from a variety of materials, including silicon, glass, metals, ceramics, and hard plastics, and they require several fabrication processes, including thermal bonding, chemical etching, and reactive ion etching, which require more

time and effort. PDMS-based microfluidic devices, on the other hand, have advantages due to their low cost, optical transparency and biocompatibility.

#### **3. Isolation of Circulating Tumor Cells (CTCs) by Microfluidic Devices**

During metastasis, cancer cells detach from the primary tumor and intrude apart into tissues in the bloodstream. To detect and isolate CTCs, various techniques including centrifugation, magnetic separation, microchips, filtration, micro/nano substrates and biomarkers have been used. With the widespread adoption of microfluidic techniques, a large number of researchers have worked hard to develop more efficient and reliable CTC separation technologies ranging from immunomagnetic beads to size-based microfluidic devices. Currently, the major commercialised products for CTC separation techniques include the CellSearch system, which uses immunomagnetic beads, and the CelarCellFX1 system, which uses size-dependent isolation [117]. The methods for isolating CTCs are mostly based on biological qualities of the tumor cells, such as specific antigen expression and receptor, or physical properties of the tumor cells, such as size and deformability. Inertial focusing, acoustics, microfluidic filters, optics and dielectrophoresis are some of the size and deformability-based approaches.

#### *3.1. Size-Based Isolation*

Due to its visibility and ease of management, CTC separation based on size and flexibility is one of the oldest approaches. The principle of separating cells from the main flow channel through filtration is, in fact, rather simple. Membrane devices are designed to act as a filter, allowing blood to flow while separating CTCs based on their size and deformability. When diluted blood travels through the main channel, cells greater than a certain size are captured by this membrane filtration set-up within the device, while smaller cells continue on their course and are separated. The risk of clogging, the requirement for frequent maintenance, cleaning, and the incapacity of cells to recover after filtration are all prevalent issues with these devices. This method's most serious flaw is that it can't separate more than one particle type in a single stage. Microfluidic systems capture tumor cells more efficiently via filtering because pore sizes and geometries are carefully controlled by microfabrication. Filters are divided into four types based on their structures: weir-type, pillars, crossflow, and membranes [118]. Size-based filtration using polymer membranes or microsieve membrane filter devices has been shown to extract CTCs from whole blood samples based on the morphological size differences between cancer cells (~15–40 μm in diameter) and leukocytes (~10 μm in diameter) [119]. The size, geometry, and density of the pores in the microfilters can be controlled uniformly and precisely. In addition, this technology can provide maximum sample processing capability via parallel arrays of multiple flow cells, which reduces processing time, cost, and filter clogging while facilitating mass production and high-throughput screening for large-scale clinical studies.

Yoon et al. developed and reported a slanted weir microfluidic channel to reduce haemocyte contamination during CTC isolation [93] (Figure 4a). With a flow rate of 2.5 mL/h and 3.8 mL/h for a breast cancer cell line (LM2 MDA-MB-231) at 0.8◦ weir to 0.5◦ weir, a high separation efficiency of ~97% was achieved. The viability of the collected tumor cells was also determined using the trypan blue assay, and it was found to be 97.1% for the 0.8◦ weir and 95.8% for the 0.5◦ weir. The viability of the 0.5◦ weir was slightly lower depending on the high flow rate and shear rate. This chip showed high separation efficiency with minimal contamination. However, the major drawback was its low throughput. Furthermore, Liu et al. developed a simple pyramid-shaped microchamber that is feasible, cost-effective, and highly efficient for CTC separation from breast carcinoma patients [82]. With an optimised flow rate of 200 μL/min, the capture efficiency of the device was assessed with a fresh blood sample in five sequence concentrations of 25–200 cells/mL using four different cancer cell lines (BGC823, H1975, PC-3, and SKBR3) spiked into DMEM medium. As a proof of concept, polystyrene beads with diameters of 10 μm (red beads) and 20 μm (blue beads) were allowed to pass through the pyramid-shaped channel at a flow rate of 200 μL/min. When the flow rate was increased to 300 μL/min, the capture efficiency

increased to 92% and 89%, respectively, at different outlet heights of 6 μm and 8 μm. This method has advantages, including lower sample consumption, a simple experimental procedure, high capture efficiency, and ease of observation. Finally, from the DMEM medium, the SKBR3 cell line had a capture efficiency of 93%, while the healthy blood sample had a capture efficiency of 89%. Further, Fan et al. designed and developed a PDMS membrane filter-based technique for the isolation of CTCs [90] (Figure 4b). At a flow rate of 10 mL/h, >90% cancer cell recovery was achieved from a blood sample spiked with lung cancer cells. Later, Zhang et al. created a label-free microfluidic device for isolating CTCs from breast cancer patients [105]. At a flow rate of 10 mL/h, the device demonstrated 73.6% capture efficiency and an 82% recovery rate. The main and side microchannels were 80 μm and 50 μm and 50 μm and 50 μm in width and height, respectively; the filter microchannel was 40 μm in width 10 μm in height. The device was used to isolate CTC cell strains such as SKBR3, MCF-7, and MDAMB231. Immunofluorescence staining was used to identify the cultured cells.

**Figure 4.** Schematic illustration of microfluidic devices for the isolation of CTCs using various techniques. (**a**) An overview of a slanted weir device; separation of CTCs over a slanted weir based on distinct size and deformability [93]. (**b**) Schematic of the microfluidic device integrated with a PDMS microfiltration membrane for CTC capture; reprinted with permission from ref. [90], 2015, Elsevier. (**c**) Top view of the multi-flow effect of a size-dependent inertial migration microfluidic system representing rotation-induced lift force (FΩ) for the isolation of CTCs [62]. (**d**) Enrichment of CTCs using spiral microfluidic technology utilizing inertial lift force [120]. (**e**) Illustration chip, self-amplified inertial-focused cell bifurcation of CTCs in the microfluidic channel; reprinted with permission from ref. [121], 2020, American Chemical Society.

#### *3.2. Inertial Focusing Microchannel-Based Isolation*

Inertial focusing is a phenomenon that occurs when suspended particles in a fluid stream migrate across flow lines and arrange themselves in equilibrium positions at specific cross-sectional positions. This behavior is caused by inertial forces within the channel and is controlled by channel geometry and flow conditions [122,123]. This phenomenon occurs in straight channels due to a balance of two dominating forces such as shear gradient inertial lift force (FSL), caused by the curvature of the fluid velocity profile and wall induced inertial lift force (FWL), caused by the particle's interaction with the nearby wall. The particles are pushed toward the channel walls by FSL, while they are moved away from the walls and toward the channel center by FWL [124,125]. As a result, the particles tend to attain a state of equilibrium where these forces are equal.