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Immunomagnetic separation (IMS) allows specific capture and isolation of intact pathogen cells by superparamagnetic beads or polystyrene beads that are coated with iron oxide and antibodies specific for a particular VTEC serogroup.
Immunomagnetic separation (IMS) for isolation of Listeria serves as the basis for assays that are ideally suited to food industry applications. These include hazard analysis and critical control point (HACCP) programmes as well as in-process and finished product testing. IMS can be integrated into such testing in two ways. There are IMS methodologies that have been specifically developed for these applications. Alternatively, IMS can be performed during execution of standard cultural methods to increase sensitivity and reduce test time.
Several attributes of IMS account for its useful properties of speed and sensitivity. Primary among these is the concentrating effect of IMS beads. Whereas enrichment increases bacterial number per unit volume as bacteria multiply over time, IMS immediately concentrates bacteria from a larger volume into a smaller volume. This is because IMS beads can be dispersed into a significant sample volume to bind pathogens, and can then be magnetically concentrated and resuspended in a smaller volume for further analysis. Enrichment can therefore be eliminated or minimized. Because of this, one IMS method for Listeria detection produces results positive and negative in 24h. Eliminating 26 days of test time allows earlier shipping of finished product and faster decontamination when pathogens are detected in the processing environment.
Omission of enrichment renders the combination of IMS with an appropriate detection method capable of enumerating pathogens and detecting injured cells. Enumeration of cells subsequent to IMS provides more information than enrichment-based methods which only report pathogens as present or absent. For example, during sanitizing to eliminate contamination, pathogen numbers may be decreasing, but may still be sufficient to result in continued positive results in tests that give either a positive or a negative result only. This could obscure the effectiveness of the chosen sanitation protocols during the decontamination process. Injured Listeria cells are often present in environmental and food samples. These injured organisms are more readily detected with IMS-based methods than with enrichment-based methods. This is because the separating and purifying properties of IMS minimize the presence of background flora which must otherwise be controlled, as in enrichment-based methods. Background flora are traditionally suppressed by use of selective agents which are lethal to injured cells and which can be omitted from IMS-based methods.
IMS can be performed without culturing organisms, facilitating the detection of the only Listeria species whose presence is regulated Listeria monocytogenes. When Listeria monocytogenes is co-present with other Listeria species, it is often overgrown and rendered undetectable. Accurate detection becomes possible with IMS even under conditions where other Listeria species would overgrow, obscuring detection by other methods. These characteristics make IMS valuable for Listeria control by both food and environmental monitoring throughout the food industry.
IMS and, in some cases, lectin-magnetic separations are often used in the above-mentioned disciplines. In microbiology they are especially used for the detection of pathogenic microorganisms. IMS enables the time necessary for detection of the target pathogen to be shortened. Target cells are magnetically separated directly from the sample or the pre-enrichment medium. Isolated cells can than be identified by standard, specific microbiological procedures. IMS is not only faster but also usually gives a higher number of positive samples. Also sublethally injured and stressed microbial cells can be very efficiently isolated using IMS. The most important microbial pathogens can be detected using commercially available specific immunomagnetic particles; they are used for the detection of Salmonella, Listeria and Escherichia coli O157. New immunomagnetic particles for the detection of other microbial pathogens are under development.
Removal of cancer cells is one of the most important applications of IMS in the area of cell biology and medicine. The first experiments were performed in the 1970s and since then an enormous number of applications have been described. Cancer cells are usually removed from bone marrow prior to its autologous transplantation and using IMS they are detected in blood. Elimination of graft-versus-host disease (GvHD) in allogenic bone marrow transplantation requires an effective removal of T cells from the bone marrow of the donor. A direct method enabled a 103 times depletion of T cells.
Magnetic particles are being increasingly used for isolation of human cell subsets directly from blood and other cell sources. B lymphocytes, endothelial cells, granulocytes, haematopoietic progenitor cells, Langerhans cells, leukocytes, monocytes, natural killer cells, reticulocytes, T lymphocytes, spermatozoa and many others may serve as examples. Cells from other animal and plant species have been successfully separated, too.
Not only whole cells, but also cell organelles can be isolated from crude cellular fractions. Dynal (Oslo, Norway) has developed Dynabeads M-500 Subcellular, which are able to isolate rapidly more than 99% of target organelles.
In the area of parasitology Cryptosporidium and Giardia are the parasites where IMS is of interest. Two commercially available kits can be used for this purpose. Both products are used in the method 1622: Cryptosporidium in Water by Filtration/IMS/FA (December 1997 Draft) of the US Environmental Protection Agency. In very low turbidity samples (clean waters), IMS has demonstrated significantly better results than the standard procedures. When water samples were turbid, the recovery efficiency of IMS diminished.
Immunomagnetic separation (IMS) has proved to be a very efficient method for separating target organisms from food materials and background flora. Antibody-coated paramagnetic particles are mixed with the sample. By exposure to a magnetic field, bound target cells are separated while the sample suspension is removed. A number of procedures may be used for subsequent final detection, such as conventional culturing, microscopy, impedance technology, ELISA, latex agglutination or DNA hybridization, partly involving amplification techniques. In addition to the short separation and concentration time, IMS technology also overcomes the problem associated with unwanted inhibition due to selective media components. Since IMS can be used in conjunction with many final detection technologies, it is expected that several automated analytical procedures will make use of this potent technique in the near future.
Immunomagnetic separation (IMS), using combinations of various magnetic beads and antibodies against specific viral surface antigens, has been used to isolate hepatitis A virus, rotaviruses, enteroviruses,7780 and noroviruses81,82 in seeded environmental water samples. More recently, the IMS coupled with membrane adsorptionelution technique has been shown to be useful for the selective and quantitative detection of enteric adenoviruses directly from river water samples.83
It has been suggested that this antibody capture provides an approach for the detection of intact and thus potentially infectious viruses, because antibodies only capture those viral particles with functional antigenic epitopes on the viral surface.84 However, IMS is probably not economical and practical for large volumes of water samples, and their application to virus concentration from field samples has not been extensively demonstrated.
IMS is now recognized as an essential technique that improves the detection of foodborne pathogens in a relatively short time. Automation of the whole concept has resolved many of the perceived problems of cross-contamination and enhanced the user friendliness of the system. Not much developmental work has been done with respect to new applications of the IMS concept since its initial introduction. Few new products have been launched for food testing perhaps because of the difficulty in developing good enough antibodies that have a sufficiently broad specificity for the mirage of increasing numbers of, for example, verotoxin-producing E. coli. Although IMS has not yet been fully integrated into all conceivable detection systems and adapted for continuous online screening and monitoring of processing plants as envisaged, it is an essential step in the isolation of specific pathogens from foodborne outbreak cases for epidemiological tracing.
The target bacteria are separated from the source and from other bacteria not bound to the particles and may be concentrated by resuspension in a small volume suitable for subsequent cultivation or detection by other means.
Possible growth inhibitory substances in the sample are removed, thus facilitating cultivation. IMS is not a detection system, but may provide a significant time-saving for all existing methods based on selective enrichment.
It enables the flexibility of applying different end-point detection techniques. Depending on requirements, detection techniques including microscopy, plating, enzyme-linked immunosorbent assay (ELISA), polymerase chain reaction (PCR), and impedance methods can be used.
Although IMS is accepted as a reliable alternative to conventional selective enrichment protocols, perceived fears of cross-contamination exist due to the manual nature of the technique. The fact that antibodies are not absolutely specific, mean that some cross-reacting organisms may bind to the beads. Bacteria frequently attach nonspecifically to surfaces and the problems of non-specific binding often arise.
Furthermore, inherent in the IMS procedure itself, is the entrapment of microorganisms within the aggregates formed during the concentrating step. Therefore, when not too specific detection methods are used after IMS, for example, plating onto agar media, the problem can become accentuated.
IMS both purifies and concentrates microorganisms. It is an established technique used for recovery of microorganisms, other types of cells, proteins etc. using specific antibodies immobilized on paramagnetic beads. The process works by incubating the beads and sample, allowing time for reaction to occur between the antibodies and the pathogen of interest. Mixing enhances this binding step. Subsequently, the beads are pulled to one side of the container using a magnet, isolating the pathogen of interest from the rest of the sample, which is removed and discarded. This beadmicroorganism complex can be used for direct plating on selective media or used for enrichment of a bacterium or for nucleic acid isolation. However, many protocols include dissolution of the beadpathogen complex, often accomplished by adding acid. Next, the now dissociated beads are removed from the pathogen sample by using the magnet to once again concentrate them at the side of the container, and removing the solution now containing the isolated pathogen. IMS offers high recovery rates, though the drawbacks are the expensive reagents required, the time necessary for binding and unbinding, and problems with unspecific binding or aggregation of nontarget cells within the bead-pathogen clusters.
IMS is used in the above mentioned standard methods for isolation of Cryptosporidium oocysts and Giardia cysts, and are commercially available from several different suppliers. This protocol includes several IMS steps where the first reduces the sample volume from 10 to 1mL and is followed by two rinses in 1mL buffer. Thereafter, the parasites are eluted in HCl for 10min and transferred to a microscope slide with NaOH for neutralization and subsequent immobilization and straining. Manufacturers report high recovery rates for IMS isolation of (oo)cysts, quoting over 95%.
To the best of our knowledge, beads conjugated with beads against other parasites are not available today. During method development, Hoffman etal. recovered E. intestinalis spores, labeled with FITC-conjugated rabbit polyclonal antibodies using two different commercially available magnetic beads conjugated with antirabbit immunoglobulin antibodies.69 A similar approach was used by Duntre and Dard (2005) for T. gondii using commercially available goat antimouse IgM coated magnetic beads and an in-house produced monoclonal antibody.70 However, this antibody lacked in specificity and a more specific antibody was later used by the same authors (2007) for tap and surface water (1020L) concentrates seeded with approximately 100 sporulated oocysts with recoveries of 74.5% from drinking water and 30.6% and 37.1% from two different surfaces waters. For this antibody, acid elution was not sufficient enough, but sonication was required to release the sporocysts. This antibody crossreacted with sporocysts of Hammondia hammondi, Hammondia heydorni, and Neospora caninum. Obtaining antibodies specific enough to isolate a microorganism from a complex background that also may contain closely related microorganisms is one of the major challenges in development of IMS methods.
IMS is also used for recovery of bacteria, mainly from enrichment broths, in analysis of pathogens in food, environmental, and clinical samples. Magnetic beads with antibodies against common pathogen that can be encountered in water, such as EPEC, E. coli O157 and Salmonella are commercially available from several suppliers. Cationic beads for recovery of viruses are also available, and IMS for viruses has been reported in several research studies.71 Automated IMS systems such as Pathatrix Auto System (ABI) will probably increase the number of pathogens against which antibodies are available.
Flow-through IMS techniques have been developed, with Ramadan and colleagues reporting a continuous flow magnetic separation system for Cryptosporidium and Giardia isolation and concentration.72 Incubation of the protozoan pathogens with the IMS beads occur as prescribed in USEPA Method 1623. Subsequently, the beadpathogen complexes enter the flow-through system. Here, as the magnetic particulate matter passes through the channel it is repeatedly captured and released by the rotation of an external permanent magnet. Finally, the concentrated sample is captured by another magnet at the end of the channel. This is illustrated in Fig. 4.8. The aim of the multiple stages performed away from the wall was to avoid the problems of aggregation sometimes observed in IMS. Due to the relatively high magnetic particle concentrations, large aggregates are formed in which impurities might be trapped. This system was reported to concentrate samples of 50mL down to 1mL, with efficiencies comparable to the existing method, performing IMS in smaller volumes in a tube, for both tap (Fig. 4.9) and secondary effluent water. When testing tap water samples, Ramandan etal. found that a glass channel flow system performed better than a disposable plastic molded cartridge, which they speculate was due to rougher walls in the cartridge offering sites where pathogens could become trapped.72 The authors report that their work is a step toward creating automated sample processing for protozoa, reducing the number of steps and human intervention required. See Chapter 10, Section 10.2.1, for further discussion of how microfluidics is applied to sample processing.
Performing IMS followed by plating onto selective or differential agars, or by rapid techniques such as PCR, ELISA, electrochemiluminescence, flow cytometry, or microscopy, markedly enhances the speed and sensitivity of assays for detection of E. coli O157:H7. Enrichment culturing times can be reduced considerably, thus the entire assay can potentially be performed in 8h or less. Immunomagnetic separation may be useful for the recovery of stressed, sublethally injured E. coli O157:H7 which are not resuscitated during selective enrichment culturing. Specificity is determined by the antibody bound to the beads, thus with the availability of appropriate antisera, improved assay systems for detection of E. coli O157:H7 as well as for other bacterial pathogens can be developed. The IMS technique is easy to perform, does not require elaborate instrumentation, and can easily be applied to isolation and detection procedures for E. coli O157:H7 or other food-borne pathogens.
Immunomagnetic separation offers an alternative approach to rapid identification of culturable and nonculturable microorganisms. The principles and application of the method are simple, but rely on suitable antibody specificity under the experimental conditions. Purified antigens are typically biotinylated and bound to streptavidin-coated paramagnetic particles (e.g., Dynal beads). The raw sample is gently mixed with the immunomagnetic beads and then a magnet is used to hold the target organisms against the wall of the recovery vial, and nonbound material is poured off. If required, the process can be repeated, and the beads may be removed by simple vertexing. Target organisms can then be cultured or identified by direct means. The IMS approach may be applied to recovery of indicator bacteria from water, but it is possibly more suited to replace labor-intensive methods for specific pathogens. E. coli O157 recovered from water samples were detected using this technique in some studies. Furthermore, E. coli O157 detection following IMS can be improved by electrochemiluminescence detection. However, the IMS/culture methods are also accompanied with disadvantages such as the ability of nonspecific binding, the need for physicochemical conditions such as pH and temperature, sensitivity to chemicals in the samples, the high cost of monoclonal antibody production, and limited shelf life.
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Zhang, X.; Tang, B.; Li, Y.; Liu, C.; Jiao, P.; Wei, Y. Molecularly Imprinted Magnetic Fluorescent Nanocomposite-Based Sensor for Selective Detection of Lysozyme. Nanomaterials 2021, 11, 1575. https://doi.org/10.3390/nano11061575
Zhang X, Tang B, Li Y, Liu C, Jiao P, Wei Y. Molecularly Imprinted Magnetic Fluorescent Nanocomposite-Based Sensor for Selective Detection of Lysozyme. Nanomaterials. 2021; 11(6):1575. https://doi.org/10.3390/nano11061575
Zhang, Xin, Bo Tang, Yansong Li, Chengbin Liu, Pengfei Jiao, and Yuping Wei. 2021. "Molecularly Imprinted Magnetic Fluorescent Nanocomposite-Based Sensor for Selective Detection of Lysozyme" Nanomaterials 11, no. 6: 1575. https://doi.org/10.3390/nano11061575
The rapid qualitative assessment of surface markers on cancer cells can allow for point-of-care prediction of patient response to various cancer drugs. Preclinical studies targeting cells with an antibody to activated matriptase conjugated to a potent toxin show promise as a selective treatment for a variety of solid tumors. In this paper, we implemented a novel technique for electrical detection of proteins on surfaces of cancer cells using multi-frequency microfluidic impedance cytometry. The biosensor, consists of two gold microelectrodes on a glass substrate embedded in a PDMS microfluidic channel, is used in conjugation with immuno-magnetic separation of cancer cells, and is capable of differentiating between bare magnetic beads, cancer cells and bead-cell aggregates based on their various impedance and frequency responses. We demonstrated proof-of-concept based on detection of activated matriptase proteins on the surface of cultured Mantle cells.
Survival of patients with cancer has been significantly improved due to the developments in new therapeutics for patients in the past decade, however, once metastatic, the disease remains incurable. Thus, new therapeutic agents as well as diagnostic tools predicting patient response are urgently needed. Overexpressionof matriptase (a membrane-bound serine type II protease) has been found in various epithelial tumor and blood malignancies suggesting that the enzyme can be used as marker for CTC detection. Due to the expression of activated matriptase in Mantle cells, anti-matriptase monoclonal antibody (M69) conjugated to monomethyl auristatin E (MMAE) for selectively targeting Mantle cells has been demonstrated as a promising therapeutic with high levels of efficacy with minimal potential side effects1,2.
One of the key processes playing a role in modulation of the tumor environment is (membrane-anchored) proteolysis3. Matriptase, a type II transmembrane serine protease, is an important pericellular protease that has an effect on tumor microenvironments, as it is responsible for initiating the protease cascade and activating growth factors. Matriptase is broadly expressed in epithelial tissues4, where the enzyme plays a crucial role in forming and maintaining epithelium integrity and epidermal differentiation, and also the placenta development, to give a few examples. There is growing evidence showing that altered matriptase expression is potentially important in hematological cells and also neoplasms5. Matriptase is shown to be expressed on the surface of THP-1 human monocytic cells5. Matriptase has also been detected on the surfaces of a wide range of cells including peritoneal macrophages6, two Burkitt lymphoma (BL) cells, and also human leukemiaand chronic lymphocytic leukemia7,8. In contrast to the situation in epithelial/carcinoma cells, these hematological cells express no or low levels of HAI-1 (Hepatocyte Growth Factor Activator Inhibitor Type 1). Various studies have examined the role and regulation of matriptase in human B-cell lymphomas, and data shows that it is expressed in various non-Hodgkin B-cell lymphomas with implication for tumor behavior9. Given the importance of matriptase in tumor behavior and its expression on a wide variety of tumor cell types, the targeted delivery of cancer drugs to the tumor site shows great promise for enhancing drug efficacy and minimizing toxicity towards non-cancerous cells. Thus, the ability to rapidly isolate tumor cells in blood and qualitatively assess matriptase surface expression levels using inexpensive miniaturized instrumentation can provide guidance and great insights in further developing this novel and promising therapeutic approach. We emphasize here that activated matriptase is present in most but not all epithelial cancers, and therefore the M-69 (anti-matriptase) antibody can identify those tumors that express activated matriptase alone or complexed with its inhibitor Hepatocyte Growth Factor Activator Inhibitor Type 1 (HAI-1).
Current technologies for sorting and assessment of surface markers on cells are bulky and unsuitable for point-of-use analysis and deployment in large-scale clinical studies. Fluorescence cytometry (FCM) and fluorescence activated cell sorting (FACS) are the gold standards for high-throughput rapid cell sorting and surface marker analysis. The technology is, however, bulky and expensive and thus not suitable for point-of-care use.
The gold standard technology for cancer cell isolation and marker assessment is the CellSearch CTC (Circulating Tumor Cells) Test, which uses magnetic bead based pre-concentration and fluorescent tagging of the cells and fluorescently analyzing the cell surfaces10. Various configurations of the CTC chip, developed by Toner and colleagues, utilizes optimal microfluidic geometries for highly efficient immuno-separation of CTCs from whole blood based on using EpCAM (epithelial cell adhesion molecule) capture antibodies11,12. More recently, the MagSweeper, an immuno-magnetic separation technique has been developed which is able to enrich tumor cells from blood by 108-fold and can process 9ml of blood per hour12. While magnetic immuno-separation methods are advantageous in that they allow for highly efficient enrichment of rare cells, yet one of primary drawbacks is that once the cells have been tagged with magnetic particles, it is difficult to separate the bead-cell aggregates from the mixture of bare magnetic beads, since magnetic fields will attract both. Thus, in order to separate the beads from the bead-cell aggregates, a trained biologist has to view the mixture visually under a microscope and manually pick the bead-cell aggregates off the slides. Alternatively, this can be done using bulky robotics with automated image analysis capability, or by staining the cells and using fluorescence analysis. All of the above technologies utilize optical-based detection, which requires bulky instrumentation and is thus unsuitable for point-of-care analysis. Previously, Holmes et al. demonstrated an impedance labelling method for identifying target antigen expressing cell subpopulations from a heterogeneous mixture13. Most recently, Lee and colleagues developed a promising microfluidic electronic sensing platform, which involves tagging CTCs with magnetic nanoparticles and using a Hall Detector to detect the MNP (magnetic nano-particles) tagged CTCs but not the MNPs, since the individual MNPs are too small to accumulate sufficient signal to trigger a response in the Hall Detector, and thus the sensor only responds to the MNP tagged CTCs14.
Various promising technologies have also been developed making use of label-free tumor cell separation based on physical properties such as size15,16,17,18 and dielectric permittivity19,20,21, allowing for downstream molecular analysis of the isolated cells. Although there is great interest in developing technologies that can be used for analysing circulating tumor cells, the ability to analyse dissociated cancer cells obtained from a tissue biopsy is also of great importance in predicting patient response to targeted therapies.
Here, we used Maver cell, a Mantle cell line, as a prototype to test the feasibility of a technique that makes use of immune-magnetic separation to pre-concentrate cancer cells and electrical impedance detection to differentiate between isolated cells and bare magnetic beads to assess matriptase expression on cancer cells. We envision ultimately using this technique to isolate tumor cells from complex samples (like dissociated cells obtained from a tissue biopsy) to predict cancer patient response to novel targeted therapeutics using anti-neoplastic agents conjugated with anti-matriptase antibody. We emphasize that our proposed technique can be used in conjunction with the above mentioned immuno-magnetic based cancer cell separation techniques to either characterize matriptase levels on tumor cells obtained from a biopsy and then dissociated into cell suspension or circulating tumor cells directly from blood.
The basic device is shown in Fig.1a. Magnetic beads, coated with an anti-matriptase monoclonal antibody (M69) that recognizes activated matriptase, are mixed with test sample containing target Mantle cells. The expression of matriptase on the membrane of cancer cells results in bead-cell aggregation. Immuno-magnetic separation is used to extract the magnetic beads and the bead-bound cancer cells from the test sample. The use of multi-frequency electrical impedance cytometry allows for differentiating between unbound beads, non-target cells and bead-cell aggregates. Beads have a relatively flat impedance response with frequency (Fig.2a), whereas cells exhibit a drop in impedance change as frequency increases (Fig.2a). This method can be used for detection and qualitative assessment of surface membrane bound protein (i.e. matriptase) levels, as the size and quantity of peaks corresponding to bead-cell aggregates is proportional to concentration of matriptase expressed on the cancer cells. Previously, we demonstrated the feasibility of using digital electronic detection of protein biomarkers22 and also the ability to fully miniaturize the footprint of the readout instrumentation to portable and wearable platforms23,23. We also demonstrated electronic bead aggregate sizing for protein biomarker detection and quantification of soluble proteins25. We also demonstrated label-free classificationof drug sensitive cells usingmulti-frequency impedance cytometry in conjunction withsupervised machine learning26.Here, we utilize information attained by simultaneous measurement of peak intensity at multiple frequencies, which allows effective differentiation between bare beads, and bead-cell aggregates, allowing for detection apparatus to be integrated onto a portable platform.
Schematic of biochip. The presence of matriptase expressing cultured cancer cells in the antibody coated beads mixture results in beads binding to the cell and aggregating. Impedance based sizing allows differentiation between magnetic beads and bead-CTC aggregates.
(a) Theoretical model for the impedance change for cells and beads as a function of frequency when beads or cells pass over sensor. (b) The electrical equivalent circuit model of a two electrode pair system with a cell suspended in buffer. Cdl: double layer capacitance, Rs: solution resistance, R: representing the occlusion of ions passing between the electrodes due to the cell volume, Cm: membrane capacitance, Rc: cytoplasm resistance.
We model the particle inside the channel and the electrode/electrolyte interface using the circuit model shown in Fig.2b. We consider a cell to consist of a membrane, which we represent as a capacitor (Cm) in series with the cytoplasm resistance (Rc). Using inert electrodes (gold), we assume an ideal polarizable electrode model with no charge transfer resistance at the interface. The model consists of a double layer capacitance (Cdl) from the left electrode in series with the solution resistance (Rs) in series with a network of impedance components representing the cell/particle in series with the double layer capacitance (Cdl) of the right electrode. The impedance of the cell consists of a resistor (R) representing the occlusion of ions passing between the electrodes due to the cell volume in parallel to the membrane capacitance (Cm) and cytoplasm resistance (Rc). Equation1 describes the total impedance across the electrodes.
The resistive component (R) is dependent on the volume of the cell/particle. Thus volume alone may be difficult to differentiate between cells and beads. However, at frequencies greater than 1MHz, as membrane capacitance of cells is significantly larger compared to beads, it results in a smaller peak amplitude compared to lower frequencies (f<1MHz) which is dependent on cell/particle size. For a bead, one can assume a negligible membrane capacitance (Cm) and a significantly higher resistivity (Rc) compared to cells.
The microfabricated biochip (Fig.3ac) consists of two gold microelectrodes on a glass substrate with the channel above it formed in a PDMS cover. The micro-channels are 400m wide and 20m high tapering down to a sensing pore which is 100m wide and 20m high. The smaller cross sectional area of the sensing pore allows for both focusing of particles and also higher electrical sensitivity to the extent where 2.8m beads can be detected at the single bead level. The spacing between the two electrodes is 20m and the width of each electrode is 15m. The PDMS devices were treated with oxygen plasma to render the surfaces hydrophilic. To begin the proof-of-concept study of CTC detection by using anti-activated matriptse antibody, we used Maver cells as a prototype. Unlike epithelial cancer cells, Maver cells are a subtype B-cell lymphoma called Mantle cell lymphoma, that are cultured in suspension. As such Maver cells are readily available for the magnetic bead-binding experiment, without the additional step to detach cultured epithelial cancer cells leading to degradation of the membrane-bound matriptase. Activated matriptase expression is found in four Mantle cell lines including Maver cells. Maver cells were used to test MFIS (multi-frequency impedance cytometry) and demonstrate that M69 mAb formed aggregates between magnetic beads and cancer cells to generate distinguishable signals. Results of the Western blot analysis demonstrate the presence of a 70-kDa active form of matriptase in those Mantle cell lines using M69, a specific monoclonal antibody to the activated enzyme (Fig.4). Cancer Cells were suspended in phosphate buffer saline (PBS).
Western Blotting showing the activated matriptase expression in Mantle Cell lymphoma cells (Jek0-1, MAVER, MINO & Z138)31. Equal amount of protein was loaded in 10% SDS-PAGE. All blots are from same gel.
An anti-matriptase monoclonal antibody (M69) conjugated to MMAE selectively targeting Maver Mantle cells was conjugated to 2.8m super-paramagnetic sheep anti-mouse IgG beads. The beads were mixed and incubated with cancer cells in PBS containing 0.1% (w/v) BSA for 1hour, where beads and cancer cells expressing matriptase formed aggregates as visualized optically (Fig.4). A magnetic separator was used to separate both the bare magnetic beads and the bead-cell aggregates from the mixture. The bare beads and the bead-cell aggregates were resuspended in PBS ready to be analysed using multi-frequency impedance cytometry.
Particles suspended in solution were injected into the micro-channels, where impedance measurements were made at multiple frequencies simultaneously. We performed measurements for three different mixtures. The first was for immune-magnetically separated 2.8m magnetic beads in PBS. The second was for a pure suspension of Maver cells in PBS. The third was for a mixture of immuno-magnetically separated magnetic beads and magnetic bead-cell aggregates. For each mixture, the experiment was run for sufficient time until at least several hundred peaks were obtained. The first and third samples were tested three times. A wavelet filter was used for both baseline drift subtraction and also noise reduction, so that peak amplitudes could be readily determined.
In order to fabricate the microelectrodes onto a glass wafer, we first designed the photolithography mask using AutoCAD. Then we photo-patterned the image on a 3 glass wafer using standard photolithography. At last, we performed electron beam evaporation to deposit thin film gold electrodes and lift off processing using acetone in an ultrasonicator to fully pattern the electrodes. Photoresist patterning is performed through the procedure below. The glass wafer was cleaned with acetone and methanol, then photoresist was spin coated, soft baked, and the wafer was exposed with ultra-violet light through chromium mask. The photoresist was developed and then underwent a hard bake. After photo-patterning, we deposited 100nm gold on top of a 10nm layer of chromium which was used for adhesion.
The micro-channels were made of polydimethyl siloxane (PDMS). The master mold for the channel was an inverted image patterned on a 3 inch silicon wafer. We fabricated the master mold using standard photolithography including wafer cleaning, spin-coating of SU-8 (Microchem Inc., Massachusetts, USA) photoresist, soft baking, ultraviolet light exposure, developing, and hard baking. We poured the PDMS mixture with a ratio of 10 to 1 (pre-polymer to curing agent) onto the silicon wafer. We degased the mixture in a vacuum desiccator to remove the bubbles, and cured the mixture in the oven at 80 C for about an hour.
After curing, we peeled the PDMS slab off of the wafer, punched 2mm and 1.2mm holes as the inlet and outlet respectively. We then aligned and bonded the PDMS channel to our micro-electrode chip after a treatment of both substrates with oxygen plasma. At last, we baked the bonded chip at 75 C for 20minutes.
A monoclonal anti-matriptase antibody (M69) was developed. We used cultured Mantle (Maver cell line) cells (Fig.3), and 2.8m superparamagnetic sheep anti-mouse IgG beads (Life Technology, Carlsbad, CA, USA) (Fig.3) for the assay. The protocol for immune-magnetic capture of matriptase expressing cancer cells went as follows: Sheep anti-mouse IgG beads were washed with PBS containing 0.1% (w/v) BSA three times. 3g of M69 antibody was suspended in PBS. The beads and the antibodies were mixed and rotated for 1h in an 1.5ml tube at 25 degrees Celsius to ensure the binding of M69 and the beads. We then washed the beads three times with PBS containing 0.1% (w/v) BSA to remove the excessive antibody. Cultured mantle cells (1.5 million cells/ml) in PBS containing 5% FBS were then added to the 1.5ml tube and gently rotated for another 1h at 25 degrees Celsius. Then, we placed the tube in a magnetic separator allowing for extraction of the magnetic beads and bead-cell aggregates (Fig.5) from the cell suspension followed by washing three times with PBS containing 5% FBS. Finally, the bead-bound cells were suspended in 200l PBS containing 0.1% BSA.
The specificity of M69 towards activated matriptase in various cancer cells has been demonstrated by a subset of this manuscripts co-authors (Lin et al.) in the referenced publication27. In this report, M69 has been shown to only recognize activated matriptase in complex with the endogenous inhibitor, HAI-1. Whereas, the other anti-matriptase antibody, M32, bound both latent and activated matriptase. Moreover, matriptase is not expressed in blood cells except monocytes5. To get rid of the signals derived from monocytes in the blood samples, the interfering noise can be simply eliminated by negative-absorption using the monocytes-specific antibody. We further emphasize that this is a platform technology. Antibody functionalization to beads and passivation to block non-specific cell capture is well established in cellular/molecular assay, and is beyond the scope of this manuscript, which was to demonstrate the ability to qualitatively assess cell surface markers rapidly with electrical detection technology.
In order to extract the electronic signals from the sensor, the biochip was connected to a lock-in amplifier (Zurich Instrument HF2 Series, Zurich Instruments, Zurich, SI) through two wires bonded to the gold pads on the biochip. The amplitude of the input AC voltage was 1V peak to peak for each channel, and the frequency ranged from 100 KHz to 20MHz. The gain of the amplifier is 1 kVolt/Ampere. While testing the electrical signal, we monitored the biochip under an optical microscope in order to simultaneously monitor the beads while measuring the electrical signal. The recorded data was then processed using a custom-written Matlab code including wavelet filter for denoising and detrending, and an algorithm for identifying the peaks and quantifying them.
Figure6 shows the measured particle impedance change figure as a function of frequency over an ensemble of particles. The impedance we plot here is the impedance change when a particle passes through the electrodes. We plot the average impedance change as a function of the frequency at f = 300 KHz, 500kHz, 1MHz, 5MHz and 20MHz for cells and bare beads. The error bars show the standard deviation for the ensemble of measurements. As seen, cancer cells exhibit a larger peak intensity compared to magnetic beads. The reason for this is because of the larger comparative volume of cells compared to beads. Cells are roughly 23X larger in radius compared to beads thus at least 8X larger in volume, which explains why the average impedance change of cells is roughly 8X larger than the beads at low frequencies. As frequency increases, the impedance change of beads remains relatively steady, whereas the impedance change of cells decreases which fits the behaviour expected from our circuit model in Fig.2a. The impedance starts to decrease at 1MHz in the sample consisting of purified cancer cells. The reason for which the impedance of the cancer cells drops faster with frequency, as opposed to the bare beads, is because cells exhibit a significantly larger membrane capacitance and smaller membrane resistance compared to bare magnetic beads.
As confirmed through visual inspection, bead-cell aggregates show larger diameter compared to both bare cells and bare beads. In the electrical measurements, this manifests itself as larger peak intensity (Fig.7). In Fig.8, we plot the distribution of the peak intensity at a frequency of 500kHz and input AC voltage of 1V, which is where impedance change of the particle is dominated by the effect of its volume. For the solution of magnetic beads, we see a normal distribution of particles with a mean value of approximately 2 V. There is a smaller percentage of magnetic beads that exhibit a mean value of approximately 10 V, which is most likely a result of beads aggregating together non-specifically. The solution consisting of cancer cells primarily exhibits a normal distribution with a mean value of approximately 15 V. In the same solution there is some variation in particle sizes due to either smaller cells or cells clustered together resulting in a larger peak amplitude. Here, we note that the final mixture, where immune-magnetic separation was performed, consists of a large percentage of bare beads and then a smaller number of bead-cell aggregates. This is because we mix the cells with an overabundance of particles to ensure efficient capture. As a result, we observe a bimodal distribution for this plot (black curve), where the smaller intensity peaks correspond to bare beads, and the larger intensity peaks corresponding to bead-cell aggregates, and the larger sized peaks correspond to large bead-cell aggregates forming.
Percentage distribution as a function of peak amplitude for experiments with (1) Pure solution of 2.8m beads (red, 300 particles detected). Higher amplitude levels greater than 10 V are due to beads aggregating, (2) Pure solution of Mantle cells (blue, 300 particles detected), (3) Mixture of bare beads with bead-cell aggregates (black, 200 particles detected). The peaks on the black curve with amplitude greater than 10 V result from bead-cell aggregates. Frequency and amplitude of the input AC voltage were 500kHz and 1V respectively. (p<0.05 for beads and cell).
Observation of the peak intensities as a function of its multiple frequencies at higher dimensions illustrates the ability to differentiate the different particles from each other. The results further demonstrate the ability to classify particle types, when studying the impedance at multiple frequencies simultaneously. The two-dimensional scatter plot for Signal-to-Noise Ratio (SNR) of the peak intensities at 500kHz and 20MHz (Fig.9) are shown. The peak intensities at 500kHz and 20MHz for the three data sets (bare bead, bare cells, and bead cell aggregates + beads) are plotted on the scatter plot, each forming distinct clusters. An ellipse is drawn around each data set, the boundaries of which representing the respective standard deviation. The primary goal is to identify the number of cells, which have aggregated with the magnetic beads. Bare magnetic beads (red) form a cluster at the bottom left corner of the x- and y-axis, because of their small relative diameter. Bare cancer cells (green) exhibit larger peak intensities, particularly at f=500 KHz, yet have lower SNR at f = 20MHz, thus have a slope of m <1. The mixture (blue) of bare beads with bead-cell aggregates exhibits properties of both particle types. On the one hand, the overall SNR (@ f = 500 KHz) is relatively smaller compared to cells, and a good proportion of them fall within the red ellipse (for bare magnetic beads). The bead-cell aggregates in the mixture have a higher SNR both at f = 500 KHz and 20MHz compared bare cells and bare beads. Thus, to quantify the number of bead-cell aggregates (to determine the extent of matriptase expression on the cancer cell surface), the particles in the blue ellipse, which have no overlap with the red and green ellipses, will give an accurate count of the bead cell-aggregates in the mixture. Everything in the blue ellipse that does not overlap with the red ellipse represents cancer cells that have been immuno-magnetically separated.
Scatter plots for SNR at 500kHz and SNR at 20MHz. Red dots correspond to pure sample of bare beads. Blue dots correspond to the mixture of beads and bead-cell aggregates. Green dots correspond to pure sample of cells. Overlap of the blue and red data sets corresponds to unbound beads.
Our experimental results showed that through the combination of immuno-magnetic cell separation and multi-frequency microfluidic impedance cytometry, we are able to effectively assess the expression of target antigens on the surface of cancer cells. Bare magnetic beads, cancer cells, and bead-cell aggregates exhibit different frequency responses as well as varying voltage peak intensity distributions. We demonstrate this for qualitatively assaying the presence of activated matriptase on the surface of cancer cells. The lowest concentration for reliable detection demonstrated in this study was 1.5 million cells/ml. This detection limit is more suitable for detection of dissociated cancer cells obtained from a tissue biopsy as opposed to circulating tumor cells in blood. Further work can be done in developing this novel analytical technique to enable qualitative assessment on matriptase levels. The work shown here, lays the groundwork necessary for developing an integrated biochip with the capability of rapidly assessing whether patients will be responsive to anti-matriptase based cancer therapeutics or not. Future work will be dedicated to isolating circulating tumor cells in blood and determining matriptase levels on captured cells. The combination of this technique along with the use of nanoelectronically barcoded beads28,29,30 can be a potential solution for analysing multiple markers on cell surfaces. Though for our experiment, we focused on matriptase and Mantle cells, we emphasize that this method is applicable to a wide array of markers and cell types.
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This project was funded by PhRMA Foundation Starter Grant for Early Faculty, Breast Cancer Research Foundation Grant to JRB, and also National Science Foundation(ECCS Award no. 1711165) and the National Science Foundation CAREER Award (Award no. 1846740).
Mehdi Javanmard, Joseph Bertino, Zhongtian Lin, and Siang-Yo Lin conceived the idea and designed the experiments. Zhongtian Lin and Siang-Yo Lin led the experiments. Pengfei Xie contributed to device fabrication. Chen-Yong Lin produced the antibodies. Gulam M. Rather contributed reagents and analysed data. Mehdi Javanmard, Joseph Bertino, Zhongtian Lin, and Siang-Yo Lin contributed to data analysis and interpretation. Mehdi Javanmard, Joseph Bertino, Zhongtian Lin, and Siang-Yo Lin wrote the paper and all authors provided feedback.
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Lin, Z., Lin, SY., Xie, P. et al. Rapid Assessment of Surface Markers on Cancer Cells Using Immuno-Magnetic Separation and Multi-frequency Impedance Cytometry for Targeted Therapy. Sci Rep 10, 3015 (2020). https://doi.org/10.1038/s41598-020-57540-7Get in Touch with Mechanic