Despite decades of intensive research, cancer remains one of the leading causes of death worldwide, with over 8 million deaths reported annually [1]. Metastasis, the primary cause of cancer-related mortality, involves tumor cells detaching from the primary site, entering the blood or lymphatic vessels, and spreading through the circulatory system to distant organs [2,3]. On reaching these organs, the cells adhere to the endothelium of the target tissue, infiltrate the tissue, and then proliferate to form secondary tumors [4]. Early diagnosis and treatment can increase the five-year survival rate up to 90 %, but this rate drops to 26 % once metastasis occurs [5]. The scarcity of circulating tumor cells (CTCs) and heterogeneity of primary tumors make early detection extremely challenging [6]. Liquid biopsy has emerged as a promising technique in precision oncology, offering a non-invasive and real-time approach to monitor treatment response, therapeutic resistance, and minimal residual disease [7]. Moreover, it enables the analysis of circulating biomarkers, such as cell-free DNA [8], extracellular vesicles [9], and CTCs [10,11]. Consequently, it offers significant potential for detecting cancer at an early stage, sometimes even before symptoms appear or tumors are visible on imaging scans.

Circulating tumor cells (CTCs), originating from primary tumors and carrying genetic information, are powerful liquid biopsy markers for cancer diagnosis, treatment monitoring, and prognosis [[12], [13], [14]]. However, CTCs are extremely rare in peripheral blood, particularly during early-stage cancer, making their efficient isolation a major challenge [15,16]. Common commercial techniques for isolating and characterizing CTCs include fluorescence-activated cell sorting (FACS) [17], magnetic-activated cell sorting (MACS) [18], and density gradient centrifugation [19]. Despite their widespread use, these approaches often require costly equipment and complex sample pre-processing. Furthermore, they are labor-intensive and may result in cell loss or damage. Once isolated, the CTCs must be lysed to release the intracellular components, such as DNA, RNA, and proteins, for downstream analysis [20]. However, current lysis techniques face several challenges, including incomplete membrane disruption, loss of target biomolecules, and the risk of contamination. Thus, optimizing lysis protocols is essential to ensure accurate and reliable biomarker analyses in both clinical and research settings [21].

Microfluidic devices offer significant advantages for CTC isolation and processing due to their rapid response, low cost, portability, minimal sample requirement, and high cell-handling efficiency [[22], [23], [24], [25]]. Current microfluidic approaches for CTC isolation can be broadly categorized into labeled and label-free methods. Labeled techniques rely on surface modifications within microchannels that exploit specific proteins expressed on CTC membranes for selective capture. Common techniques include immunocapture using chaotic mixing channels [26], aptamer-based capture [27], and micropost arrays [28]. However, chaotic mixing complicates downstream CTC recovery, while aptamers suffer from low throughput, and micropost arrays impose high shear forces that may damage the cells and are expensive to fabricate. Immunomagnetic capture methods employ magnetic particles functionalized with antibodies or aptamers to bind the CTCs, which are then isolated using an external magnetic field [[29], [30], [31]]. Although effective, this approach often results in the CTCs adhering to the particles, thereby hindering their subsequent analysis. In addition, immunofluorescence techniques, which use fluorescent probes to detect specific DNA sequences, offer high sensitivity and specificity but are prone to false-positive or false-negative results [32].

In contrast, label-free microfluidic methods rely on the intrinsic physical properties of the CTCs themselves, such as their size [33], deformability [34], and other biophysical characteristics [35], to avoid the need for surface markers. These methods are broadly classified as either active or passive, depending on the mechanisms used to induce cell motion or separation. Active techniques use external force fields such as dielectrophoresis [36], acoustics [37], or optically induced dielectrophoresis [38] to manipulate and separate the cells. However, while these methods offer a high throughput and separation efficiency, they require additional equipment, which increases the system complexity and cost. Moreover, the need for precise exposure to external fields often results in longer residence times and lower flow rates. To address these issues, passive methods rely entirely on cell-fluid interactions and the channel design. Typical methods include microvortices [39], deterministic lateral displacement [40], and inertial focusing [41,42]. Specifically, prior studies have demonstrated the use of microvortices to alter cell trajectories, vortex-induced flows to enhance mixing, and deformability-based enrichment to achieve label-free cell separation [[43], [44], [45]]. These approaches eliminate the need for external forces, thereby enabling faster, lower-cost processing and improved cell viability. As a result, label-free separation has attracted growing attention for the development of next-generation microfluidic devices for biomedical applications.

Centrifugal microfluidic platforms, also known as lab-on-a-disc (LOD) systems, utilize the inertial forces generated by rotation to drive and control the fluid flow within microchannels patterned on the disc surface. These systems eliminate the need for external pumps or complex interconnections while effectively removing air bubbles and residual volumes. Furthermore, precise fluid manipulation can be achieved through the incorporation of valves, microchannels, and reservoirs into the system [46]. LOD devices support many functions, including valving [47], metering [48], and mixing [49], and are thus well suited for diagnostic and point-of-care (PoC) applications. Additionally, they enable density-based fluid transport and separation [50]. Recent studies have demonstrated their success in various component separation tasks, including leukocyte isolation [51], immune cell enrichment [52], blood cell counting [53], and CTC separation [54].

Centrifugal microfluidics facilitates label-free cell separation and alignment through the inertial effects induced in specially designed microchannel geometries. Among the various designs that have been proposed, contraction-expansion arrays (CEA) have attracted particular attention owing to their high cell separation performance. Shamloo et al. [55] demonstrated the successful separation of MCF-7 cells in a hydrodynamic CEA in a centrifugal platform. However, the separation efficiency was rather low (76 %) due to the high sample concentration (target-to-nontarget cell ratio of 1:10 with approximately 5000 cells/μL). Bakhshi et al. [56] proposed a label-free continuous CTC separation method that combined inertial focusing with dielectrophoretic forces. The numerical results indicated a high efficiency of 99.5 %; however, the separation performance was not confirmed experimentally. Farahinia et al. [57] developed a centrifugal microfluidic system based on multiple CEAs and biocompatible magnetite-arginine nanoparticles. Although the device achieved an experimental separation efficiency of 89.1 %, its practicality in clinical contexts was undermined by the need for time-intensive labeling, significant spatial demands on the disc, increased operational costs, and a high design complexity.

Following CTC isolation, genomic and transcriptomic profiling is often required to analyze the gene expression patterns, identify mutations, and evaluate biomarkers relevant to cancer diagnosis and treatment. However, to access this genetic material, the cell membranes must be disrupted using lysis reagents [58,59]. Effective on-disc chemical lysis requires efficient mixing of the target cells with a lysis reagent to ensure complete membrane disruption. Thus, properly designed microfluidic mixers that provide adequate channel lengths and residence times are essential [60,61]. Jahromi et al. [62] developed a centrifugal microfluidic system that used passive pneumatic and inertial forces to mix the cell samples with lysis reagents and maintain precise fluid control. However, the design utilized a multilayer disc architecture, which increased the system complexity and fabrication cost. Nasiri et al. [63] proposed an integrated centrifugal microfluidic device combining a CTC separation unit and an on-disc micromixer for lysis. While promising, the system was evaluated only through numerical simulations without experimental validation.

Given the limitations described above, there remains a critical need for simple and efficient integrated platforms for cancer cell enrichment and subsequent lysis. This study meets this need by developing a cost-effective, centrifugal microfluidic device capable of isolating and lysing cancer cells from whole blood samples without the need for time-consuming and labor-intensive labeling techniques. The separation module incorporates a Y-shaped inlet for sheath and sample introduction, a trapezoidal CEA microchannel for inertial focusing, and a bifurcated outlet for size-based cell separation. A high-speed actuated siphon valve is employed to direct the separated cancer cells into a square-wave serpentine microchannel, where they mix with lysis buffer and then flow into a downstream chamber. The performance of the proposed integrated device is investigated both numerically and experimentally. The results confirm that the proposed device enables rapid, automated, and cost-effective CTC isolation and lysis and thus has a strong potential for clinical applications in cancer diagnostics and personalized medicine.