Isolation, Extraction, and Analysis of Cells After Confined Migration

Abstract

Cell migration through confined microenvironments is a critical biological process that underlies numerous physiological and pathological events, including immune cell trafficking, tissue morphogenesis, and cancer metastasis. Although polydimethylsiloxane‐based microchannel devices have enabled detailed studies of confined migration, the efficient collection of cells post‐migration for downstream molecular analyses remains a major challenge. Existing approaches often rely on harsh mechanical dissociation that compromises cell viability and integrity and do not permit in situ collection of cell lysates. To overcome these limitations, we have developed the Trap‐based Recovery After Permeation (TRAP) chip, a pump‐free microfluidic platform that integrates controlled confined migration with efficient post‐migration cell or lysate collection. The TRAP chip incorporates microchannel arrays terminating in a precisely engineered trap region that enables gentle recovery of cells or cellular components without exposing them to high shear forces or requiring large buffer volumes. This innovation ensures the viability of recovered cells and expands the applicability of confined migration assays beyond imaging‐based studies. We demonstrate that the TRAP chip facilitates the extraction of post‐confinement cells for mechanical characterization, including measurement of Young's modulus, as well as the isolation of proteins and RNA suitable for downstream assays such as western blot and qPCR. The TRAP chip thus represents a significant advancement in microfluidic technologies, offering a robust, reproducible, and minimally invasive approach for studying the mechanobiology of confined migration, with broad potential for applications in basic research, cellular engineering, and translational studies where cell behavior under physical confinement is of critical importance. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1: Fabrication of TRAP and control chips

Basic Protocol 2: Cell seeding and live cell isolation from TRAP chips

Alternate Protocol: Biomolecular extraction from TRAP chips

INTRODUCTION

Cells in vivo encounter complex confining microenvironments throughout the body, ranging from the narrow pathways of capillary beds to the diverse interstitial spaces of tissues (Paul et al., 2017). Cell migration through these confined spaces is a multifaceted phenomenon that influences a variety of biological processes, including organ development (Chitnis et al., 2024), immune responses (Moreau et al., 2018), and cancer metastasis (Okeyo et al., 2015). To better understand the cellular responses to these complex in vivo confining microenvironments, it is crucial for bioengineers to develop in vitro experimental systems capable of recapitulating key variables of these tissue tracks (Doolin & Stroka, 2019; Liu et al., 2015; Paul et al., 2016). Of these, polydimethylsiloxane (PDMS)‐based microchannels are particularly popular due to their straightforward fabrication, ease of extracellular matrix (ECM) protein adsorption, and ability to precisely control the degree of confinement, allowing for a more accurate analysis of cellular responses to specific levels of confinement (Doolin & Stroka, 2018; Hsia et al., 2022; Irimia & Toner, 2009; Li et al., 2024; Todorovski et al., 2023; Zhao et al., 2021). Furthermore, these microfluidic devices are particularly compatible with multiple microscopy modalities (Afthinos et al., 2022; Holle et al., 2019; Zhao et al., 2021), allowing for timelapse imaging of cellular interactions with confining microenvironments (Nguyen et al., 2013). This compatibility with microscopy allows microchannel systems to be utilized for a wide variety of image‐based analyses of both fixed and live cells (Lee & Holle, 2024). Such analyses include detailed morphological analyses of the area, aspect ratio, or volume of the cell and nucleus (Dahl et al., 2008; Denais et al., 2016); determination of the localization of specific proteins (Hsia et al., 2022); tracking of cell migration dynamics (Holle et al., 2019); and observation of ion flow dynamics (Stroka et al., 2014). Despite these advancements and their usefulness for a wide range of image‐based assays, it remains extremely difficult to extract cells from these devices after they have undergone confined migration due to technical challenges in accessing cells within exit reservoirs. This highlights a major limitation of the microchannel assay in general, as many biological assays useful for understanding mechanosensitive cell behavior require large numbers of post‐confined cells, ranging from simple western blots and qPCR to more advanced and modern techniques like flow cytometry, RNAseq, and ATACseq (Lu et al., 2024). Furthermore, if confinement is to be utilized as an approach to mechanically pre‐condition cells destined for downstream applications like tissue engineering or immunotherapy, then the gentle isolation of large numbers of these cells post‐confinement is necessary. Thus, there is a great need for a microchannel platform that allows for the extraction of cells or cellular components post‐confinement.

Current research on the collection of cells following confinement in microchannel‐based systems remains limited. One attempt at addressing this challenge involved the development of a microchannel device that allows for confined migration prior to physical separation of the PDMS‐based microchannel array from the glass substrate (Bui et al., 2018). In this system, manual separation of the microchannels from the glass substrate results in cells remaining attached to either the glass or the microchannels. This implies that physically ripping apart the cell‐ECM junction is required by this system, which will inherently alter cells due to the harsh mechanical stimulus. Finally, this approach is only capable of population‐level analysis of cells that were inside the channels at the moment of harvest, meaning that no distinction can be made between a cell that has just entered the confinement and one that has migrated an extremely long distance over a long period of time. More recently, a microchannel device incorporating a cell collection chamber has been developed to facilitate the harvesting of cells via trypsinization, followed by their transfer to well plates for subsequent lysate collection (Bastianello et al., 2024). Although this method enables downstream analysis, it does not accommodate in situ lysate collection. The enzymatic treatment and replating processes inherent in this approach might compromise the integrity of cellular omics data. Consequently, there is a critical need for a microchannel system capable of in situ isolation and collection of cell lysates. Such a system should support analysis of cells that have experienced confinement over a defined journey and offer the ability to assess cellular responses to varying degrees of confinement.

To address the challenges of cell extraction from microchannel‐based devices, we developed the Trap‐based Recovery After Permeation (TRAP) chip, an improved system that enhances cell recovery and collection efficiency through key modifications in chip cutting and assembly. Unlike standard microchannel confinement devices (Fig. 1) (Gao et al., 2025), the TRAP chip features four arrays of identically designed microchannels to ensure uniform confinement. A stereomicroscope‐guided cutting technique removes excess PDMS in the outer reservoir precisely near the microchannels. Additionally, the incorporation of a PDMS‐based trap wall during assembly creates a small uncovered “trap region” at the end of each microchannel array, specifically designed to facilitate efficient in situ cell or lysate collection post‐migration (Fig. 1). This refined approach significantly reduces shear stress and pipetting inefficiencies, providing a more effective solution for post‐confinement cell recovery and analysis.

Figure 1.

Pictures and schematics of a standard chip, TRAP chip, and control chip.

In this article, we demonstrate that the TRAP chip functions as a versatile and user‐friendly platform that enables efficient collection of post‐confinement cells while preserving cell viability and the integrity of cellular components. The chip also facilitates direct collection of cell lysates for downstream biomolecular assays. This design innovation simplifies the process of cell recovery, making the TRAP chip an effective tool for conducting downstream biochemical assays. These assays can provide valuable insights into cellular behaviors and responses as a function of confinement, enhancing our understanding of cell dynamics under these circumstances. Herein, the fabrication procedures for both the TRAP and control chips are described in Basic Protocol 1. Basic Protocol 2 details the cell seeding process and the subsequent isolation of viable cells following confinement. The methodology for collecting cell lysates for downstream biomolecular analysis is described in the Alternate Protocol.

NOTE: When handling PDMS blocks containing microchannel patterns, avoid direct contact with the patterned surface. Always use fine‐tipped tweezers for transfer and refrain from clamping or applying pressure to the microchannel regions.

NOTE: During medium changes within the chips, minimize mechanical stress on the PDMS. After cell seeding, avoid pressing, poking, or deforming the chips to prevent disruption of cells.

Basic Protocol 1. FABRICATION OF TRAP AND CONTROL CHIPS

This protocol outlines the step‐by‐step procedure for the fabrication of TRAP chips as well as control chips (Fig. 1). Control chips were designed to replicate the microenvironment of the trap region in TRAP chips while omitting microchannels. These control devices serve to approximate the cell density and environmental conditions present in the trap region following confined migration, thereby providing a baseline for comparison in downstream analyses. Chips are fabricated using PDMS replica molding, followed by precision cutting and bonding to glass substrates via plasma treatment. The bonded chips are subsequently coated with collagen to enable cell adhesion.

Materials

• PDMS and curing agent (SYLGARD^TM 184 Elastomer Kit, Dow Corning, 04019862)

• Collage type I (Rat‐tail collagen I, Gibco, A10483‐01)

• 0.1% (v/v) acetic acid (see recipe)

• 70% (v/v) ethanol

• Phosphate‐buffered saline (PBS), sterile (1^st BASE, BUF‐2040‐10X4L)

• Silanized silicon wafer with microchannel patterns

• 15‐cm Petri dishes (Thermo Fisher Scientific, 150468)

• Double‐sided tape (3M Science, 7000042028)

• Analytical scale (Precisa, 321 LG)

• Spatula

• Nitrogen gun

• Vacuum degassing chamber (Memmert, V029)

• 80°C oven (Memmert, INB200)

• Scalpel (handle No. 3, Swann‐Morton, 0933; blade No.11, Swann‐Morton, 1103)

• Large glass plate or cutting mat

• Razor blades (VWR^®, 55411‐050)

• Stereomicroscope (SDPTOP, SZM)

• Fine‐tipped tweezers (Electron Microscopy Sciences, 78319‐4)

• Customized 13 × 13–mm and 18 × 18–mm metal punches

• 15‐ml Falcon tube (Corning, 352096)

• Adhesive tape (e.g., cello tape)

• Glass coverslips (Marienfeld, 0112700)

• Plasma cleaner (Harrick Plasma, PDC‐001‐HP)

• 6‐well plates, sterile (Thermo Fisher Scientific, 140675)

• Parafilm

• 37°C, 5% CO₂ cell incubator (Thermo Fisher Scientific, 371)

• Biosafety cabinet [ESCO Airstream ^® NS (S‐series) Class II Type A2 Biological Safety Cabinet, model no. AC2‐4S8‐NS‐PORT]

• Aspirator

• 1Secure the silanized silicon wafer with microchannel patterns to the base of a 15‐cm Petri dish using double‐sided tape to prevent movement during casting. Weigh the PDMS base and curing agent to ensure a 10:1 weight ratio. Thoroughly mix the components with a spatula for ∼1 min, ensuring the mixture is homogeneously blended and evenly aerated with small bubbles.

  The required weight of PDMS mixture depends on the number of chips and specific experimental requirements. For standard fabrication, ∼2 g of the mixture is needed per chip. If casting the silicon wafer for the first time, pour ∼75 g PDMS mixture to ensure complete coverage. During the cutting process (step 5), cut only along the outer edges of the wafer. This approach preserves the marginal PDMS bulk, allowing subsequent casts to require only ∼2 g PDMS mixture per chip design to fill the wafer regions. When molding chip walls in a 15‐cm Petri dish, ∼75 g PDMS mixture is typically sufficient. The PDMS thickness may be adjusted as needed for different applications.

• 2Clean the surface of the silicon wafer by blowing compressed nitrogen to remove any dust or debris. Carefully and evenly pour the prepared PDMS mixture over the silicon wafer containing the microchannel mold. Simultaneously, pour 75 g of the PDMS mixture into a separate 15‐cm Petri dish designated for fabricating chip walls.
• 3Place the Petri dishes containing the wafer and PDMS mixture into a vacuum degassing chamber. Degas for ∼5 min at 10 mb.

  The degassing time may vary depending on the power of the vacuum system. Ensure that most of the air bubbles are removed. A few minor surface bubbles can be accepted, as they typically dissipate during the curing process.

• 4Transfer the degassed wafer and PDMS mixture to an oven. Cure at 80°C for 90 min.
• 5After curing, use a scalpel to peel the PDMS block containing the microchannel features from the silicon wafer. Keep PDMS block in a clean 15‐cm Petri dish.

  Avoid touching the microchannel features on the PDMS, as any touch may introduce dust or compromise the integrity of the microchannels. Always keep the microchannel features facing up in Petri dishes and keep containers sealed when not in use to maintain sterility and surface cleanliness.

• 6Place the cured PDMS block containing the microchannel designs onto a large glass plate or a cutting mat. Using a scalpel, carefully dissect the PDMS into individual pieces, each containing one complete chip design.
• 7Cut out a square in the middle of the chip to create an uncovered inner reservoir designated for cell seeding.
• 8Using razor blades, under a stereomicroscope, precisely trim the excess PDMS from the outer reservoir regions along all four sides of the microchannels on each chip. Using fine‐tipped tweezers, place the processed chips into a 15‐cm Petri dish.

  Avoid contact with the microchannel surface during cutting to prevent contamination or damage. Use fine‐tipped tweezers to handle the chips by the outer reservoir region only because such precision instruments with narrow, pointed ends are suitable for accurate handling while maintaining minimal contact. Be cautious with sharp tweezer tips, as accidental contact with the microchannel area may compromise device integrity. Keep the chip with microchannel features facing up.

  After trimming excess PDMS, inspect the cut TRAP chip under a microscope with measurement capability. Discard any chip with a remaining outer reservoir length exceeding 50 µm, aiming to minimize the residual length as much as possible.

• 9To prepare the TRAP chip wall, first use a customized 18 × 18–mm metal punch to extract a PDMS block from the cured PDMS mixture from step 4. Then, employ a customized 13 × 13 –mm metal punch to remove the central portion of the block, creating a wall‐like structure suitable for cell culture compartmentalization and compatible with 30‐mm glass coverslips.
• 10To prepare the inner part of the control chips, use razor blades to cut PDMS blocks of dimensions comparable to those of the processed chips in step 9.

  Steps 1 to 10 can be repeated as needed, depending on the number of chips required for the experiment.

• 11Prepare a fresh solution of collagen type I at a concentration of 100 µg/ml in 0.1% acetic acid in a 15‐ml Falcon tube.

  Each TRAP chip requires ∼300 µl collagen solution, whereas each control chip requires 500 µl. Calculate the total required volume based on the number of chips to be coated.

• 12Clean the PDMS chip and control PDMS block surfaces using adhesive tape (e.g., cello tape) to remove dust and debris.

  Avoid applying excessive pressure on the microchannel side to prevent damage to the microchannel structures.

• 13Clean glass coverslips by rinsing with 70% ethanol and then dry them thoroughly using compressed nitrogen.
• 14Place the cleaned coverslips, PDMS chips, control PDMS blocks, and PDMS walls into a 15‐cm Petri dish.
• 15Insert the open Petri dish into the plasma cleaner chamber and perform oxygen plasma treatment following the manufacturer's guidelines.

  Plasma parameters may vary depending on the equipment model used. Here, we use the Harrick Plasma machine (PDC‐001‐HP), where 1 min of oxygen (20 mb) is introduced and the surface is activated for 30 s under the highest power level (30 W).

• 16Use fine‐tipped tweezers to invert each control PDMS block and PDMS chip and carefully align them at the center of a glass coverslip to assemble the inner components of the control and TRAP chips, respectively.
• 17Gently press down on the control PDMS block and PDMS chip, ensuring firm contact at all four corners as well as over the microchannel regions of the PDMS chip to promote uniform bonding.
• 18Similarly, flip the PDMS wall and align it properly over the chip to form the trap region.
• 19Press firmly on top of the PDMS wall to enhance bonding.

  Plasma activation is transient, with complete bonding occurring quickly. For more than six chips, it is recommended to divide the bonding process into batches to maintain bonding efficiency.

• 20Place the assembled TRAP and control chips at 80°C for 10 min to strengthen the bond between the PDMS and glass.
• 21Transfer each assembled chip to a sterile 6‐well plate and add the prepared collagen type I solution (see step 11) to fill the reservoirs and trap regions.

  When adding solution into the reservoirs, position the pipet tip at a corner of the uncovered inner reservoir and dispense slowly to minimize bubble formation. If bubbles appear, gently press on the PDMS from above to release them.

• 22Degas the chips for 3 min in the vacuum degassing chamber with the pressure set at 10 mb.

  This step ensures that no air bubbles remain inside the narrow microchannels, allowing for a uniform collagen coating.

• 23Seal the 6‐well plate with Parafilm and incubate the chips overnight in a 37°C, 5% CO₂ cell incubator to allow collagen adsorption onto the PDMS surfaces.
• 24Rinse the chips three times with sterile PBS in a biosafety cabinet using an aspirator.

  To remove liquid from the chips, attach a fine pipet tip (e.g., a P20 tip) to the catheter of the aspirator and position the tip at the corner of the uncovered inner reservoir to facilitate efficient aspiration. If any residual liquid is difficult to remove, gently press on the PDMS to help dislodge it for complete removal. When adding PBS, dispense the solution slowly at the corner of the reservoir to minimize bubble formation. If persistent bubbles appear, briefly degas the chip again, as there are no cells present at this stage. The PBS should be diluted to 1× with Milli‐Q water and filtered through a 500‐ml Steritop bottle‐top filter unit with pore size 0.22 µm.

• 25Seal the chips with Parafilm and store them in PBS at 4°C if not using immediately.

  Ensure that the reservoirs and microchannels remain filled with PBS and add sufficient PBS to each trap region to prevent drying. Drying can cause collagen bundling, which may adversely affect cell migration. Properly stored chips are stable for up to 3 weeks at 4°C.

Basic Protocol 2. CELL SEEDING AND LIVE CELL ISOLATION FROM TRAP CHIPS

This protocol describes the step‐by‐step procedure by which cells are seeded into TRAP chips (Figure 2A) and how cells can be collected from the trap regions by trypsinization for downstream analysis (Figure 2B).

Figure 2.

Schematics of cell seeding and cell isolation in the TRAP chip system. (A and B) Workflow of cell seeding into TRAP chips (A) and isolation of cells post‐confinement from TRAP chips (B). i: Covered inner reservoir, ii: Uncovered inner reservoir, and iv: Trap region.

Materials

• PBS, sterile (1^st BASE, BUF‐2040‐10X4L)

• Cell culture medium (see recipe), 37°C

• HT1080 cells (ATCC, CCL‐121)

• 0.05% (w/v) trypsin

• TRAP and control chips (see Basic Protocol 1)

• Biosafety cabinet [ESCO Airstream ^® NS (S‐series) Class II Type A2 Biological Safety Cabinet, model no. AC2‐4S8‐NS‐PORT]

• Cell culture flasks (Thermo Fisher Scientific, 156499)

• Aspirator

• Tissue culture microscope (Invitrogen, EVOS M5000)

• 15‐ml Falcon tubes, sterile (Corning, 352096)

• Standard tabletop centrifuge

NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly.

NOTE: All culture incubations are performed in a 37°C, 5% CO₂ cell incubator (Thermo Fisher Scientific, 371) unless otherwise specified.

• 1Remove the TRAP and control chips from the refrigerator, rinse all chips once with sterile PBS, and keep them submerged in PBS within the biosafety cabinet.

  Refer to the annotation on adding solutions to chips in Basic Protocol 1 for detailed instructions on media addition and removal.

• 2Sterilize the chips by exposing them to UV light for 30 min in the biosafety cabinet.
• 3Remove PBS completely and add warm cell culture medium to all reservoirs and trap regions. Ensure the entire chip is submerged in culture medium.

  Refer to the annotation to Basic Protocol 1, step 21, for detailed instructions on medium addition and removal.

• 4Incubate the chips in a 37°C, 5% CO₂ cell incubator for ≥30 min prior to cell seeding.
• 5Passage the HT1080 cells (cultured in cell culture flasks) with 0.05% trypsin and prepare a single‐cell suspension (at a density of 1 × 10⁶ cells/ml) for seeding in cell culture medium.
• 6Remove the chips from the incubator and completely aspirate all cell culture medium from the reservoirs and trap regions.
• 7Seed 7 µl HT1080 cells (at a density of 1 × 10⁶ cells/ml) into the covered inner reservoir of each TRAP chip using a pipet. Seed 1,000 cells in 200 µl medium into the trap region of each control chip.

  Refer to the annotation to Basic Protocol 1, step 21, for the detailed procedure for adding solutions to chips. If bubbles form in the covered inner reservoir during cell seeding, do not press down on the PDMS, as mechanical force may damage the cells. Small bubbles are generally not problematic; however, large bubbles should be avoided, as they create a longer distance for cells to migrate before entering the microchannels. If numerous large bubbles appear, completely aspirate the medium from all regions, rinse the chips with culture medium until no cells are visible, and then reseed the cells.

  The optimal cell seeding density should be determined based on the specific cell line and experimental goals. In this protocol, HT1080 cells are used, and the chosen seeding density typically results in confluency within the inner reservoir after ∼4 days. Control chips are included to mimic the microenvironment of the trap region in TRAP chips, but without any microchannels. These controls are intended to reflect the cell number and conditions observed in the trap region after migration, specifically following the 4‐day incubation period used here.

• 8Check the cell seeding under a tissue culture microscope. If no cells have entered the trap region and the seeding appears uniform in the covered inner reservoir, gently add cell culture medium to the uncovered inner reservoirs and trap regions of the TRAP chips. Ensure that sufficient medium is also added to the control chips.

  If cell seeding is uneven and cells have entered the trap region, completely aspirate the medium from all regions, rinse the chips with culture medium until no cells are visible, and then reseed the cells.

  Avoid overfilling, as immersing the entire TRAP chip at this stage can cause cells to float from the uncovered inner reservoir into the trap region. Ensure the medium height is approximately equal between the uncovered inner reservoir and trap region to minimize hydrostatic pressure differences, which could introduce confounding effects. For control chips, which contain only a trap region, directly add approximately 100 to 200 µl medium.

• 9Incubate all chips for ≥2 hr to allow complete cell attachment. After confirming attachment under the microscope, top up the cell culture medium to fully immerse both the TRAP and control chips.

  Cell culture medium replenishment is needed every 2 days. The medium underneath the PDMS in the covered inner reservoir should be retained to maintain cell viability. During medium replenishment, avoid using the pipet or aspirator near the corners of the uncovered inner reservoir, as the strong suction force may damage cells through shear stress. Rather, place the pipet or aspirator at the corners of the trap region to minimize potential damage to the cells within the trap region.

• 10On the day of cell extraction, pre‐warm the 0.05% trypsin and sterile PBS to 37°C.
• 11Use a pipet or an aspirator to carefully aspirate cell culture medium from the uncovered inner reservoir and the trap region.

  Refer to the annotation for step 9.

• 12Rinse the trap region and uncovered inner reservoir twice with pre‐warmed PBS.

  Unlike previous PBS rinsing (step 1), where complete medium removal is required before PBS addition, here, it is sufficient to directly add PBS to the trap region and uncovered inner reservoir. This is because the objective is solely to collect cells from the post‐confinement region, not to fully exchange the medium.

• 13After rinsing, add around 100 to 150 µl pre‐warmed 0.05% trypsin into the trap region and place a few drops (50 to 100 µl) of PBS into the uncovered inner reservoir. For control chips, directly add 400 µl pre‐warmed 0.05% trypsin into the trap region.

  When adding PBS to the uncovered inner reservoir or trypsin into the trap region, avoid overfilling to prevent it from spilling into another region. Because different solutions are placed across the microchannels, a gradient will naturally form. To minimize cell exposure to trypsin in the inner reservoir, perform this step quickly and transfer the chips to the cell incubator immediately and gently. The volume of trypsin can be adjusted according to the thickness of the chip. Ensure that the trypsin fully covers the trap region.

• 14Incubate the chips in the cell culture incubator for 3 min and then check whether most of the cells in the trap region have rounded up and detached.

  The recommended trypsinization time is <5 min, as extended exposure may reduce cell viability. Gentle shaking of the plate or tapping the bottom of the plate can help dislodge the cells.

• 15Using a P200 pipet, gently and slowly resuspend the cells within the trap region and then aspirate and transfer them into a sterile 15‐ml Falcon tube containing 1 ml pre‐warmed cell culture medium.

  While pipetting, take care to avoid spilling the suspension into the uncovered inner reservoir, as this can significantly reduce cell collection efficiency.

• 16Centrifuge the cell suspension for 3 min at 150 × g.
• 17Carefully remove the supernatant and resuspend the cell pellet in 200 µl cell culture medium for downstream experiments.

  The pellet may not be visible due to the low number of collected cells. When aspirating the supernatant, avoid placing the aspirator tip too close to the bottom of the tube. Gradually tilting the Falcon tube while aspirating helps maximize supernatant removal without disturbing the pellet. The resuspension volume may be adjusted as needed. Perform a cell count to estimate the total number of collected cells. Optimization of seeding density, culture duration, and the number of pooled chips is necessary and may vary by cell line due to differences in migration ability.

BIOMOLECULAR EXTRACTION FROM TRAP CHIPS

In addition to live cell isolation (Basic Protocol 2), cell lysates, like protein lysates or RNA lysates, are also instrumental in understanding confinement mechanobiology. This alternate protocol describes how to directly extract protein lysates or RNA lysates from TRAP chips via a freeze‐thaw approach (Figure 3).

Figure 3.

Pictures and schematics of biomolecular extraction from TRAP chips. i: Covered inner reservoir, ii: Uncovered inner reservoir, and iv: Trap region. Cell lysate collection should be performed on ice; the setup shown here is for illustrative purposes only.

Materials

• Protease inhibitor cocktail (Halt Protease and Phosphatase Inhibitor Cocktail, 100×, Thermo Fisher Scientific, 1861281)

• 0.5 M EDTA stock solution (Thermo Fisher Scientific, 1861274)

• RIPA lysis buffer (Thermo Fisher Scientific, 89900)

• TRIzol lysis buffer (Qiagen, 79306)

• Seeded TRAP and control chips (see Basic Protocol 2, step 9)

• PBS, sterile (1^st BASE, BUF‐2040‐10X4L), 4°C

• Aspirator (optional)

• 1.5‐ml Eppendorf tubes

NOTE: Depending on the cell migration ability and culture duration, different numbers of chips may need to be pooled to obtain sufficient cell lysates. In our work, three control chips and six TRAP chips per confinement pattern were cultured for 4 days and pooled separately to generate a single lysate per condition.

NOTE: All steps are recommended to be performed in a biosafety cabinet, except for the freezing step (step 5).

• 1Prepare fresh protein lysis buffer by diluting protease inhibitor cocktail in RIPA lysis buffer and adding 0.5 M EDTA stock solution to a final concentration of 5 mM and then place it on ice. Alternatively, place TRIzol lysis buffer on ice for RNA extraction.

  Use 100 µl protein lysis buffer per experimental condition. For RNA extraction, use 100 µl TRIzol lysis buffer per chip.

• 2Remove the seeded TRAP and control chips from the cell incubator and use an aspirator or pipet to carefully aspirate all cell culture medium in the uncovered inner reservoir and the trap region.
• 3Rinse the uncovered inner reservoir and the trap region with cold sterile PBS, without aspirating the medium in the covered inner reservoir. For control chips, proceed directly to step 7 after rinsing the trap region with cold PBS.
• 4After PBS rinsing, add a small volume of cold PBS (50 to 100 µl) to the uncovered inner reservoir of each TRAP chip.

  Avoid overfilling, as excess PBS can spill into the trap region and interfere with the lysate.

• 5Immediately transfer the TRAP chips to a –80°C freezer.

  This step should be performed as quickly as possible to prevent PBS from diffusing through the microchannels into the outer reservoir. PBS typically takes ∼10 min to freeze completely. The purpose of frozen PBS in the open inner reservoir is to minimize diffusion of the lysis buffer into this region. The solid ice acts as a physical barrier, slowing fluid exchange and thereby enhancing the purity of the collected cell lysates.

• 6After 10 min, remove the TRAP chips from the freezer and place them on ice to thaw.

  Once the ice in the uncovered inner reservoir begins to melt and reflect light, proceed to the next step. Avoid moving forward too early, as residual ice crystals in the trap region may block buffer addition and hinder efficient lysate extraction.

• 7a For protein extraction: Add 100 µl cold protein lysis buffer (see step 1) to the trap region of the first chip and use a P200 pipet tip to vigorously scrape the cells. Transfer the resulting lysate to the trap region of the next chip and repeat the scraping process. Continue this pooling process until all chips for a given confinement condition have been processed.

  Thanks to the trap region design, extracted proteins can be concentrated in a small volume. To minimize PBS diffusion into the trap region during extended handling, consider dividing the chips into two sets. For example, extract three chips first and then thaw and process the remaining three. Any bubbles formed during scraping may be aspirated and added to the final lysate tube for the corresponding condition.

• 7b For RNA extraction: Add 100 µl cold TRIzol lysis buffer (see step 1) to the trap region of each chip and use a P200 pipet tip to thoroughly scrape the cells. Transfer the resulting lysate from each chip into a 1.5‐ml Eppendorf tube.

  Unlike for protein extraction, concentrating the RNA lysate during scraping is not necessary, as further processing (e.g., RNA purification) will be performed downstream.

• 8Store all the lysates at –80°C until further downstream processing.

REAGENTS AND SOLUTIONS

Acetic acid. 0.1%

• 0.1 ml acetic acid (Sigma‐Aldrich, 33209‐1litre)

• 99.9 ml Milli‐Q water (Merck, Direct B)

• Filter with 500‐ml Steritop bottle‐top filter unit with pore size 0.22 µm (Merck, S2GPU05RE)

• Store ≤3 months at 4°C

Cell culture medium

• 445 ml Dulbecco's modified Eagle's medium (Gibco, 10569‐010)

• 50 ml fetal bovine serum (Gibco, 26140079)

• 5 ml penicillin‐streptomycin (Gibco, 15140‐122)

• Filter with 500‐ml Steritop bottle‐top filter unit with pore size 0.22 µm (Merck, S2GPU05RE)

• Store ≤1 month at 4°C

COMMENTARY

Background Information

Cell migration through confined microenvironments is fundamental to many biological processes, including immune surveillance, tissue morphogenesis, and cancer metastasis. Microfluidic systems, particularly PDMS‐based microchannels, have been widely used to study confined migration because they allow precise control of geometry and compatibility with advanced imaging. These platforms have enabled detailed analyses of cell morphology, migration dynamics, and protein localization under confinement. However, a key limitation of existing devices is the difficulty in recovering viable cells or lysates after migration. Conventional approaches often rely on harsh mechanical disruption or enzymatic treatment, which can compromise cell viability and distort molecular signatures. Furthermore, most systems are designed primarily for imaging‐based assays and do not support efficient in situ collection for downstream analyses such as western blotting, qPCR, RNA‐seq, or ATAC‐seq. This restricts their broader application in mechanobiology, cellular engineering, and translational studies. To address these challenges, the TRAP chip was developed. This pump‐free microfluidic platform incorporates engineered trap regions that enable gentle, efficient recovery of cells or lysates following confined migration. By minimizing shear stress and bypassing the need for harsh dissociation, the TRAP chip preserves cell integrity and expands the potential of confinement assays to include both mechanical characterization and molecular profiling.

Critical Parameters

Several critical factors influence the successful fabrication of microfluidic chips (Basic Protocol 1). Precise cutting along the boundaries of microchannels in the outer reservoirs under the guidance of a stereomicroscope is required to make sure the majority of cells will be in the open trap region after migration through the microchannels. Cleanliness of both the coverslips and the PDMS surfaces is also essential, as residual debris can impede proper bonding between the two surfaces and may result in chip detachment. The parameters of plasma treatment, including power and oxygen flow duration, also play a key role in determining the efficiency and reliability of the PDMS‐to‐glass bonding. It is recommended that users optimize these settings to achieve robust attachment. Precise alignment during PDMS attachment is crucial. Placing both the PDMS chip and the trap wall at the center of the coverslip ensures uniform alignment on all four sides. Misalignment may lead to an uneven trapping area or insufficient space for structural components such as channel walls. Lastly, maintaining chip moisture is essential, as drying can compromise the functionality of the collagen coating, potentially impairing cell attachment and migration.

During cell seeding (Basic Protocol 2), particular care must be taken while pipetting cells into the chip. Rapid pipetting may force small cells to flow through the wider microchannels; therefore, gentle and controlled pipetting techniques are strongly recommended. For downstream analysis, such as cell lysate collection (Alternate Protocol), rapid freezing of the chip is preferred to minimize biomolecular degradation. Once the ice melts at the central uncovered reservoir, immediately perform lysis on ice to prevent PBS from flowing into the trap region and diluting the lysate. When processing multiple chips, it is advisable to divide them into separate freeze‐thaw groups to maintain consistency and minimize degradation.

Troubleshooting

Potential problems in conducting the protocols, along with the possible causes and solutions, are shown in Table 1.

Table 1.

Troubleshooting Guide for TRAP Chip Fabrication and Cell Seeding, Isolation, and Extraction

Problem	Possible cause	Solution
Chip detachment	Surface of PDMS or coverslip is not clean	Use cello tape to clean the PDMS multiple times and clean the coverslips
Chip detachment	Power of plasma is low	Increase the duration of the plasma treatment or the power of the plasma
Chip detachment	Oxygen is not enough	Increase the duration of the oxygen introduction or the pressure of the oxygen
Cells do not adhere or migrate actively across the microchannels	Collagen coating is insufficient	Dissolve collagen properly in fresh acetic acid and check the collagen coating by immunofluorescence staining or do a couple of rinses with acetic acid if there is too much collagen blocking the entrance of the microchannels
Cells do not adhere or migrate actively across the microchannels	Chips have partially or fully dried up	Make sure the chips are always immersed in PBS when stored
Contamination in the chips	Sterilization of chips by UV is not sufficient	Increase the duration of UV exposure
Cells migrate under the attached PDMS region	Chip detachment	Please refer to the causes and solutions for chip detachment above
Cells flow into the trap region during cell seeding	Pipetting is too harsh	Practicing gentle pipetting
Cells flow into the trap region during cell seeding	Chip detachment	Please refer to the causes and solutions for chip detachment in the previous sections
Low yield when isolating cells	Cell generally do not migrate often	Increase the number of chips being pooled or seed more cells
Low yield when isolating cells	Trypsinization is not efficient	Appropriately increase the duration of trypsinization or the concentration of trypsin
RNA or protein degradation	Freeze‐thaw duration is too long	To prevent sample degradation, either perform the freeze‐thaw and cell lysis steps quickly or process the chips in smaller batches

Statistical Analysis

All data are reported as the mean ± 95% confidence interval (unless specified otherwise) and were analyzed in GraphPad Prism (9.0). Welch's ANOVA (for normally distributed data with unequal variance) with Dunnett's T3 post‐hoc test (when the sample size was less than 50) and one‐way ANOVA (for normally distributed data with equal variance) with Tukey's post‐hoc test (for groups with an equal size) were used when appropriate. P‐values less than 0.05 were considered statistically significant.

Understanding Results

The objective of these protocols is to establish an easy‐to‐use microchannel system (Basic Protocol 1) that enables the collection of cells following confinement, either as live cells (Basic Protocol 2) for functional assays or as lysates (Alternate Protocol) for downstream biomolecular analyses. Depending on the experimental requirements, researchers can select the appropriate assays to perform on the isolated cells or harvested lysates. Utilizing this platform, we report for the first time the post‐confinement isolation of HT1080 cells, an invasive fibrosarcoma cell line, and the subsequent measurement of their Young's modulus—a key mechanical parameter that is instrumental in elucidating confinement‐related mechanobiology. The nanoindentation setup used for these measurements is described in Figure 4A and in Method 1 in the Supporting Information. Our results revealed that HT1080 cells exhibited a progressive decrease in Young's modulus with increasing degrees of confinement, with an approximate 40% reduction observed in cells that migrated through long, 3‐µm‐wide microchannels (Figure 4B). These findings demonstrate that the TRAP chip system provides a robust and versatile platform for investigating the mechanobiological consequences of confined cell migration.

Figure 4.

TRAP chips enable live cell isolation after microchannel confinement and post‐confinement mechanical analysis. (A) Schematic of the nanoindentation experimental setup. (B) Young's modulus measurement for cells before and after migrating though long microchannels with different widths. (n = 36, 38, and 40). Welch's ANOVA with Dunnett's T3 multiple comparisons was used. Error bars represent 95% confidence intervals. *P < 0.05, *** P < 0.001.

Furthermore, cell lysates were collected post‐confinement to enable downstream biomolecular assays. Notably, we demonstrated for the first time that confined migration can induce early osteogenic differentiation in human mesenchymal stem cells, as evidenced by qPCR and western blot analyses at both the transcript and protein levels (Gao et al., 2025). Additionally, this platform was employed to investigate cancer cell confinement mechanobiology. Lysates from HT1080 cells showed no statistically significant differences in Lamin A or YAP mRNA levels following confined migration (Figure 5A), although significant differences in protein abundance were present for both, as measured via western blot (Figures 5B to 5D). This mismatch between transcript levels and protein levels suggests that cells modulate their gene expression in complex ways and further supports the need for new tools like the TRAP chip that allow for more complete analysis of confinement‐sensitive responses. As this tool can also be integrated with sequencing technologies, deeper insights into the mechanobiological responses associated with cell confinement will be an important focus moving forward.

Figure 5.

TRAP chips support on‐chip cell confinement and lysate collection for biomolecular analysis. (A) qPCR analysis of Lamin A, YAP, and Actin mRNA expression in cells before and after migration through long microchannels with different widths (n = 3). (B) Representative western blot images of YAP, Lamin A, and GAPDH. (C and D) Quantification of relative protein expression for YAP (C) and Lamin A (D) (n = 3). Data are the mean ± SD; one‐way ANOVA with Tukey's post‐hoc test. *P < 0.05, **P < 0.01.

Time Considerations

Basic Protocol 1 can be finished within 4 hr on average. The PDMS replica molding might take more time depending on the number of chips needed. However, do note that the chips need to be rinsed the second day. Therefore, the overall duration of Basic Protocol 1 is 2 days. Basic Protocol 2 requires around 4 to 5 hr to accomplish. The Alternate Protocol can be finished within 2 hr.

Author Contributions

Xu Gao: Conceptualization; data curation; formal analysis; investigation; methodology; writing ‐ original draft. Yixuan Li: Conceptualization; investigation; methodology. Jia Wen Nicole Lee: Conceptualization; investigation; methodology. Jianxuan Zhou: Investigation; methodology. Vaishnavi Rangaraj: Data curation; investigation; methodology. Avery Rui Sun: Investigation. Jennifer L. Young: Investigation. Andrew W. Holle: Conceptualization; funding acquisition; methodology; project administration; resources; supervision; writing ‐ original draft; writing ‐ review and editing.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Method 1 Procedure for measuring the Young's modulus of post‐confinement cells obtained from Basic Protocol 2.

Gao, X. , Li, Y. , Lee, J. W. N. , Zhou, J. , Rangaraj, V. , Sun, A. R. , Young, J. L. , & Holle, A. W. (2025). Isolation, extraction, and analysis of cells after confined migration. Current Protocols, 5, e70204. doi: 10.1002/cpz1.70204

Data Availability Statement

The data, tools, and material (or their source) that support the protocols are available from the corresponding author upon reasonable request.