Article pubs.acs.org/ac
Microfluidic Cytometer for the Characterization of Cell Lysis Jeffrey R. SooHoo,† Joshua K. Herr,‡ J. Michael Ramsey,‡ and Glenn M. Walker*,† †
Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Chapel Hill and Raleigh, North Carolina 27695, United States ‡ Department of Chemistry, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599, United States
ABSTRACT: Blood cytometry and intercellular analysis typically requires lysis as a preparatory step, which can alter the results of downstream analyses. We fabricated a microfluidic cytometer to characterize erythrocyte lysis kinetics. Forward light scatter from erythrocytes was used for enumeration at specific locations on a microfluidic chip. Diffusive transport coupled with laminar flow was used to control the concentration and exposure time of the lysis reagent Zap-OGLOBIN II to erythrocytes. Standard clinical practice is to expose erythrocytes to lysis reagent for 10 min. Under optimum conditions, we achieved complete erythrocyte lysis of a blood sample in 0.7 s. A maximum lysis reaction rate of 1.55 s−1 was extrapolated from the data. Lysis began after 0.2 s and could be initiated with a lysis reagent concentration of 1.0% (68.5 mM). An equation that related lysis reagent concentration, [A], to erythrocyte lysis, [B], was determined to be [B] = −0.77[A]0.29t. literature of 0.1 s for ammonium chloride12 and “instantaneous”13,14 for detergents. Lysis kinetics has been studied for hypotonic15,16 and colloidosmotic12,17 methods. Various other methods have been investigated on individual cells such as by laser, nanoscale barbs, and electrical pulses.18 However, outside of a few investigations on individual cells,18 there is a limited amount of kinetics data for detergent lysis available in the literature. In one case, an instrument for measuring lysis times has been demonstrated but only provided a lumped measurement of lysis time within bulk solutions and not an exact analysis of the underlying lysis kinetics.19 Blood and lysis reagents were mixed in a tank, and absorbance measurements were used to monitor lysis progress. When the absorbance crossed a threshold, buffer was added to the mixture to stop the reaction. Measurements from this bulk lysis device produced lysis times of 8 s. More recently, microfluidic approaches have relied on bulk fluorescence measurements.8 The microfluidic method achieved the same lysis time of 8 s. The goal of this study was to use a microfluidic flow cytometer to provide exact cell counts per unit time that could be used to quantify, at a high time resolution, the time-course
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rythrocyte removal is a common preparatory step for many blood assays and in particular for the analysis of leukocytes. Other cell types, such as bacteria, are also lysed before analyzing their intracellular contents. A typical lysis procedure is to mix diluted whole blood with lysis reagent, incubate the mixture for 10 min, and fix the remaining cells. The goal is to minimize, but not necessarily eliminate, the erythrocyte population because large numbers of erythrocytes can interfere with subsequent assays. However, this procedure can overexpose the leukocytes to lysis reagent, which can reduce leukocyte counts1−4 in some cases as much as 60%5 and alter their phenotype.6 Microfluidic devices provide an attractive approach for minimizing leukocyte damage during erythrocyte lysis. The control that micrometer length scales give over diffusion of molecular species and the positioning of single cells allow one to minimize leukocyte exposure to lysis reagent. Various microfluidic devices have been demonstrated for erythrocyte lysis7−11 and have the added benefit of minimizing nonspecific damage to the leukocytes. However, previous microfluidic work has focused on maximizing the number of recovered leukocytes and not necessarily minimizing the lysis time. In the articles that used lysis reagents,8,9,11 minimum lysis times of 8,8 10,9 and 308 seconds were observed using a detergent, DI water, and ammonium chloride, respectively. However, these times are longer than the theoretical minimum times reported in the © 2012 American Chemical Society
Received: September 16, 2011 Accepted: January 10, 2012 Published: January 10, 2012 2195
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flood UV exposure using a custom photomask (Photoplot Store, Colorado Springs, CO). After photoresist development (MF-319; MicroChem Corp., Newton, MA) and removal of exposed chromium (Chromium Etchant; Transene, Danvers, MA), channels were etched into the glass substrate using a dilute HF/NH4F solution (10:1 Buffered Oxide Etch; TransenE). The etched substrates were then diced, and access holes were made by powder blasting (Comco Inc., Burbank, CA). All glass was then thoroughly cleaned by sonication in a 5% solution of Contrad 70 (Fisher, Waltham, MA) for 10 min, followed by immersion in Nanostrip 2X solution (Cyantek Corp., Fremont, CA) for 30 min. Etched glass substrates and glass covers were then hydrolyzed in a 2:2:1 solution consisting of DI H2O, 30% NH4OH, and 30% H2O2 for 30 min at 70 °C, followed by thermal bonding at 550 °C for 10 h. Three reservoirs, each approximately 100 μL in volume, were bonded to the top of the device with epoxy. A threaded port for connection to a syringe pump via PEEK tubing was bonded at the outlet. Reagents and Sample Preparation. The most common lysis reagents are hypotonic, colloid-osmotic, or detergent based solutions. We used a detergent lysis reagent, ZapOGLOBIN II (Coulter, Miami, FL), because of its lysis efficiency, speed, and ability to completely break down the erythrocyte wall. Hypotonic and colloid-osmotic lysis leaves the cell wall intact producing cell “ghosts” that may be counted as cells by a cytometer. The lysis ingredient is a quaternary ammonium salt: ethyl-hexadecyl-dimethyl azanium bromide, C20H44NBr. Zap-OGLOBIN II also includes cyanides and sodium nitrite for stabilization of hemoglobin into cyanmethemoglobin for colorimetry. In solution, the salt dissolves into a negative bromine ion and a positive quaternary ammonium ion, as shown in Figure 2. The ammonium ion, similar in structure
of erythrocyte lysis using a detergent based lysis reagent. We used computer models to predict the distribution of lysis reagent within the microfluidic flow cytometer and verified the model with data from the device. We explored the effect of lysis reagent concentration and exposure time on erythrocyte lysis rates. Understanding these kinetics is crucial for the rational design of microfluidic devices that will be used for processing blood samples.
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EXPERIMENTAL SECTION Microfluidic Device Design. The device consisted of three inlets and one outlet as shown in Figure 1. Erythrocyte lysis was
Figure 1. Flow diagram of device. Vacuum was applied to the outlet channel, which drew fluid through the three inlets in a predefined ratio based on inlet resistances. Data were collected at measurement points 1 mm apart, providing a repeatable and high time resolution picture of erythrocyte lysis.
performed by injecting diluted whole blood and lysis reagent into separate inlets and allowing their streams to flow side-byside. A third inlet, at 13 mm downstream, was used to inject phosphate buffered saline (PBS) to dilute the lysis solution and slow erythrocyte lysis for the remaining 10 mm of microchannel length. A syringe pump (Pump 11 Pico Plus, Harvard Apparatus) in withdraw mode attached to the outlet was used to pump fluids through the device. Diluted whole blood, lysis reagent, and PBS were all loaded into on-chip reservoirs. The lengths of the inlet microchannels were chosen to yield fluidic resistances that ensured a 4:1:4 lysis reagent/blood sample/ buffer stream width ratio within the main lysis microchannel. All microchannels were 25 μm deep. The first 13 mm of the main channel, which contained the sample and lysis reagent streams, was 90 μm wide. The remaining 10 mm, which contained blood, lysis reagent, and PBS, was 162 μm wide. All inlet microchannels were 90 μm wide. The amount of time that erythrocytes were exposed to lysis reagent was controlled by adjusting the syringe pump flow rate. The syringe pump was used to generate blood sample flow rates of 1.0, 0.5, 0.25, and 0.125 μm/min. Using these flow rates, erythrocytes were exposed to lysis reagent for 0.767, 1.53, 3.07, and 6.13 s, respectively. For each exposure time, erythrocytes were exposed to maximum lysis reagent concentrations of 25%, 5%, 2.5%, and 0% (v/v). The balance of the mixture was PBS. In comparison, lysis reagent concentrations in standard protocols vary from 1% to 10%.20,21 Microchip Fabrication. Glass microfluidic chips were fabricated using standard wet chemical etching techniques as described previously.22,23 Briefly, white crown glass substrates (B270) coated with chromium and positive photoresist (Telic Co., Valencia, CA) were patterned with the chip design by
Figure 2. Ethyl-hexadecyl-dimethyl azanium bromide. The ingredient in Zap-OGLOBIN II that lyses erythrocytes.
to the lipids in the cell membrane, rapidly dissolves the erythrocyte membrane; leukocytes membranes are more resistant. Four hundred microliters of Optiprep (BD Biosciences, San Jose, CA) were mixed in an EDTA coated tube (BeckonDickinson K2-EDTA Vacutainer) with 1 mL of PBS Buffer (Sigma 10 × PBS P7059 diluted with filtered DI water) and 2.8 μL of whole blood taken from a finger prick of a volunteer (IRB study 151-09). This provided a 28% preparation of Optiprep, which was neutrally buoyant for erythrocytes and eliminated cell settling. Because the Optiprep mixture diluted blood 500fold, the four experimental blood flow rates of 1.0, 0.5, 0.25, and 0.125 μL/min yielded cell counting rates of of 1 × 104, 5 × 103, 2.5 × 103, and 1.25 × 103min, respectively. Computer Model of Flow and Diffusion. A twodimensional mass transport model of the device was created in COMSOL4.0a. We modeled the effect of flow rate and starting lysis reagent concentration, C0, on the concentration profile of the lysis reagent throughout the device. The height of the channel was sufficiently small compared to channel width (25 μm vs 90 and 165 μm) that we assumed variation in concentration in this dimension was negligible. The models 2196
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Figure 3. Diagram of experimental setup. The device was placed on an X−Y stage, and a laser was directed to the collection points. At each measurement point, forward scatter data was collected, filtered, digitized, and stored for later analysis.
normalized to allow comparison among the different flow rates, since slower flow rates resulted in lower absolute cell counts. Fluorescein images were acquired with a FITC cube (Chroma Technology, Brattleboro, VT) and a cooled highresolution monochrome digital camera (Hamamatsu ORCAER) mounted to an inverted epi-fluorescence microscope (Olympus IX71). Exposure times were held constant at 500 ms. Images were captured and analyzed using IPLab (Scanalytics, Rockville, MD).
were verified experimentally by substituting Fluorescein for lysis reagent and imaging the fluorescent concentration profile. Fluorescein has a similar molecular weight to the lysis reagent (fluorescein C20H12O5 MW of 332.3 vs lysis reagent C20H44N+ MW of 378.5), so we assume the diffusion coefficient of Fluorescein in PBS (2.7 × 10−6 cm2/S)24 is similar to that of the lysis reagent. Data Collection and Analysis. The microfluidic device was integrated with a laser and photodiode to create a flow cytometer,25 as shown in Figure 3. A 488 nm solid-state laser (40 mW) (Cyan; Newport Corp., Irvine, CA) was focused to an elliptical spot (15 μm × 100 μm) from above the chip using a pair of crossed cylindrical lenses. The major axis of the spot was perpendicular to the flow, minimizing the chance for coincident cells while covering all possible lateral paths a cell might take in the channel. A 40× microscope objective (NA = 0.45) (Creative Devices Inc., Neshanic Station, NJ) was positioned under the microchip for the collection of light from elastic scattering. The reflected scatter signal was directed through a beamstop to block unscattered light and neutral density filter and onto a photodiode (DET10A; Thorlabs, Newton, NJ) for detection. Current output from the photodiode was amplified through low-noise current preamplifiers (SR570; Stanford Research Systems, Sunnyvale, CA). Signals were digitized using a multifunction I/O card (PCI-6251; National Instruments, Austin, TX). The acquisition rate was 20 kHz for the highest flow rate and reduced in proportion to the flow rate. Cell counts were performed three times (n = 3) at each of 22 observation points on the chip. Raw data was collected for 10 s during each cell count. Cell counts were extracted from the raw photodiode data with a custom LabVIEW (National Instruments) program. Before acquiring cell count data, background noise levels for each observation point were determined by averaging the signal amplitude over a time slice when no cells were present. During experiments, peaks with amplitudes five times the background noise level were considered cells. Cell counts were generated for each 10 s interval and each observation point. The data were
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RESULTS AND DISCUSSION Effect of Flow Rate on Lysis Reagent Distribution within the Microchannel. The COMSOL model of flow and diffusion was validated by comparing the computed concentration output with images of a fluorescent model system. Fluoroscein solution was used in place of the lysis solution and PBS in place of the diluted blood sample. By matching the flow rate conditions with the computational model, we could use the model to predict lysis solution concentrations at arbitrary locations in the device. The representative images in Figure 4 show that the modeled concentration profile of lysis reagent agrees well with experimental results. The slight protrusion of the fluorescein stream into the PBS stream in Figure 4A was likely due to the rounded sidewall (i.e., nonvertical) profiles typical of the isotropic wet etching process. Further validation was done by comparing intensity linescans taken from the COMSOL model and the fluorescent images. Figure 5A highlights the two areas we compared: immediately after the intersection and 10 mm downstream. The COMSOL model accurately predicted the diffusion of the Fluorescein across the width of the microchannel, shown by the linescans in Figure 5B,C. The difference between the experiment and model in Figure 5C was due to the rounded walls of the channel from the etching process that caused a drop in fluorescent signal at the walls of the channel. A correction factor was generated from the ratio of how much the signal with the device filled with fluorescein differed from a constant, which is what would be expected in a 2D model. Without the correction, starting 25 μm 2197
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Figure 4. Comparison of computational model against chemical model for lysis reagent flow and diffusion in a microchannel. (A) Section of entire device that the following images represent. (B) Photo using fluorescein as model for Zap-OGLOBIN II. (C) Computational model of lysis solution concentration. Figure 5. The calculated and experimental concentration profiles of fluorescein across the channel width at the inlet channel intersection (B) and 10 mm downstream (C). Locations shown in (A).
from either edge, the signal is attenuated and quickly approaches zero at the edge. We assumed that the correlation between the image data and model data at these specific locations implied a correlation at all locations in the device. After validating the COMSOL model with Fluorescein, we calculated the change in concentration of lysis reagent that cells would see as they flowed downstream during lysis. The linescans chosen in Figure 6A approximate the trajectories that cells followed within the device. For the lysis channel, the trajectory was 9 μm from the wall, which is half the width of the sample stream, and for the quenching channel, the trajectory again followed the middle of the sample stream that occurred in the center of the channel 81 μm from either wall. Figure 6B,C shows the calculated concentrations that the cells were exposed to. Lysis reagent concentrations increased as the residence time between the blood sample and lysis reagent streams increased. The flow rate of the streams provides control over lysis reagent exposure time and concentration. The 0.125 μm/min blood sample stream equilibrated with the lysis reagent stream by the end of the lysis channel, 10 mm downstream. Under this condition, cells were exposed to a lysis reagent concentration of 0.8 × C0. At higher flow rates, the blood sample stream did not fully equilibrate with the lysis reagent stream before arriving at the quenching inlet, resulting in submaximum concentrations. Blood sample streams with flow rates of 1.0, 0.5, 0.25, and 0.125 μL/min were exposed to final lysis reagent concentrations of 1.34%, 1.84%, 2.23%, and 2.48%, respectively. Once lysis reagent quenching with PBS began, a reciprocal but attenuated trend was seen (Figure 6C). The decrease in lysis reagent concentration was attenuated because the diffusion
distances are longer after merging the quenching stream and more time is required to reach equilibrium. Effect of Lysis Reagent Concentration on Lysis Time. Having validated the model, we then investigated the effect of lysis reagent concentration on lysis time. First, control experiments were run at flow rates of 1, 0.5, 0.25, and 0.125 μL/min using erythrocytes suspended in buffer without lysis solution. No lysis occurred, and the cell counts remained constant throughout the length of the device, as shown in Figure 7A. Increasing the flow rate also increased the cell flux, which led to proportionally higher total cell counts. The average standard deviation in cell counts of the four flow rates without lysis solution was ±10.0%. On the basis of this observation, we chose a 25% decrease in normalized cell count as the threshold for the initiation of lysis. We next exposed erythrocytes to 2.5%, 5%, and 25% lysis solution at flow rates of 1, 0.5, 0.25, and 0.125 μL/min to determine how concentration and flow rate affect lysis. Figure 7B−D shows cell counts versus time for three different concentrations of lysis reagent. In these plots, cell counts were normalized to facilitate comparison among flow rates, and the x-axis was changed from distance to time so that lysis rates could be more easily determined and compared. Figure 7B shows how cell counts decrease over time for 2.5% lysis solution. Erythrocytes traveling at a flow rate of 1 μL/min exited the lysis channel at 0.767 s, when only 50% of the cells had been lysed. Erythrocytes flowing at the other three flow rates resided in the channel long enough for 100% lysis. 2198
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To determine the relationship between reagent concentration and lysis time, we assumed that the lysis process occurred in two steps: a delay period with little decrease in counts and a lysis period when cells were lysed at a constant rate. A linear regression analysis was performed on the data in Figure 7B−D. Only data points below the 75% lysis threshold were used in the analysis. The x-intercept of each fitted line was used to extract the time required for lysing 100% of the cell population. Lysis reagent concentrations of 2.5%, 5.0%, and 25% required 1.44, 1.22, and 0.73 s, respectively. We plotted these times and fitted them to a curve proportional to the inverse of reagent concentration (Figure 8), which yielded an r2 of 0.91. As concentration increases, the curve asymptotically approaches 0.7 s. This limit, which we observed at 25% concentration, was over an order of magnitude faster than previously reported experimental lysis times of 8 s. The 0.7 s limit is due to a transport delay of lysis reagent to the cells and a finite reaction rate for lysis. The model in Figure 8 suggests that lysis reagent must be greatly diluted in order to stop lysis reactions. We next determined the lowest lysis reagent concentration required to initiate lysis. The 1 μL/min cell count data at a reagent concentration of 2.5% (Figure 7B) were chosen because they provided the highest resolution cell count data at the lowest lysis reagent concentration. We overlaid the cell count data with the lysis reagent concentration determined by the computational model, as shown in Figure 9. We assumed that, once the percentage of unlysed cells dropped below 75%, lysis had begun. Using the plots in Figure 8, we observed that the percentage of unlysed cells dipped below 75% at 0.2 s. At 0.2 s, the lysis reagent concentration within the channel, the concentration that the cells were exposed to, was 1%. At 0.2 s, the lysis reagent has not reached equilibrium, and the lysis reagent concentration has not yet reached its final concentration. Thus, these results show that lysis can be initiated in as little as 0.2 s with 1% lysis reagent. Using this same curve-plotting technique with other flow rates and lysis reagent concentrations, we found that lysis initiation times were as short as 0.2 s and the majority were approximately 0.5 s. Effect of Lysis Reagent Concentration on Lysis Rate. To determine lysis rate as a function of reagent concentration, we modeled the lysis system as:
Figure 6. Longitudinal model results at lysis solution flow rates of 1.0, 0.5, 0.25, and 0.125 mL/min. (A) Highlighted cell paths. Plots of lysis reagent concentration as seen by cell (B) after lysing solution sample intersection and (C) after intersection of diluting buffer.
Even though cells were not completely lysed at the fastest flow rate, the data all possess the same slope, which suggests that the lysis rate does not depend on flow rate within the channel but rather on the lysis reagent concentration. Increasing the lysis reagent concentration increases the lysis rate, as shown in Figure 7C,D. The flow rate within the lysis channel affects the exposure time of erythrocytes to lysis reagent but not the lysis rate. To ensure complete lysis, the residence time of erythrocytes within the lysis channel should be greater than the x-axis intercept of the fitted line for a given concentration. The x-axis intercept becomes shorter as reagent concentration increases. Thus, 100% of cells were lysed at a 1 μL/min flow rate at 25% reagent concentration (Figure 7D) but not at lower concentrations (Figure 7B,C). Flow rates of 0.5, 0.25, and 0.125 μL/min were all slow enough for 100% lysis at all concentrations tested. This result has an important design implication for lab-on-a-chip devices: high throughput devices should use proportionally higher concentrations of lysis reagent to ensure complete erythrocyte lysis, with the caveat that these concentrations risk additional leukocyte damage or lysis.
k
A+B→A+P
(1)
where A is the lysis reagent, B is the cells, k is the reaction rate, and P is the product of cell lysis. The data in Figure 7B−D show that the concentration of cells, [B], did not influence the lysis rate within our experimental test procedure. Thus we can write the reaction rate, r, as: r=
d[B] = −k[A]m dt
(2)
Rearranging the equation and solving for the cell concentration gives [B] = −k[A]m t + [B0]
(3)
Lysis rates for each concentration were determined by performing a linear regression on the data in Figure 4B−D and calculating the slope of each line. The slopes for concentrations of 2.5%, 5.0%, and 25% were 0.75/s, 1.23/s, and 1.55/s, 2199
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Figure 7. Graph of exposure vs cell count for four different lysis solution concentrations. (A) 0% concentration plotted vs distance downstream. Each line represents a different flow rate. (B) 2.5%, (C) 5%, and (D) 25% concentration plotted against time. For each concentration, the lysis rates were fit to lines taken over as a subset of the entire line starting when the remaining cells dropped below, and stayed below, 75%. Measurements were not taken over the 3 mm bend.
Figure 8. Plot of average times to complete lysis for each concentration. Using the X-intercept of the linear fits of Figure 7, we calculated how long lysis took to complete at each concentration and fitted the data to the equation y = Ax−1 + B.
Figure 9. Graph of the cell counts at the lowest concentration at the highest flow rate. The cell count drops below the 75% threshold, representing the start of lysis, at 0.2 s. At this time, the concentration of lysing solution in the cells is 1%.
respectively. These points were then plotted and fit to eq 3, which yielded k = 0.71 and m = 0.25 with an r2 of 0.89, as shown in Figure 10. We used this equation to predict [B] over time for the three different lysis reagent concentrations. These
predicted concentrations agree well with the experimental data, as shown in Figure 10. The model for lysis rate shows that cell concentration is linear with respect to time. A weak dependence on lysis reagent 2200
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concentration is also predicted; increasing the lysis reagent concentration has a diminishing effect on the lysis rate.
CONCLUSIONS
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AUTHOR INFORMATION
REFERENCES
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Figure 10. Lysis rate vs concentration fitted to power law equation. k = 0.71 and m = 0.25.
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Article
In summary, we have demonstrated a microfluidic device to investigate lysis rates at time resolution not previously achieved. Travel times through the device were as low as 0.767 s, and there were 20 observation points over 23 mm. This provided a means of monitoring the effect of the lysis reagent on the cells in 32 ms increments. Using this device, we then characterized lysis concentrations, times, and rates. The minimum lysis reagent concentration needed to initiate lysis was found to be less than 1.0%. At this concentration, lysis was initiated in 0.2 s. This implies that using a significantly lower concentration than the standard amount of lysis solution is possible but will result in longer lysis times. Lysis rates were then characterized, and an equation that relates lysis rate with concentration was generated. From the rate analysis, we determined that increasing the concentration of the lysis reagent does not increase the lysis rate much beyond 10%, lysis slows rapidly but does not stop by diluting the lysis solution, and the typical lysis reagent concentrations of 1% to 10% yields a wide range of lysis rates. A minimum complete lysis time of 0.7 s was observed at a lysis reagent concentration of 25%. There are many other detergent based lysis agents that this instrument would be useful in investigating such as: Saponin,26 TRITON X-100 and TRIS/HCL,14 formic and acetic acids,27 organic buffers,28,29 enzymatic such as lysozyme, and other quaternary ammonium salts.30 Comparison to nondetergent based lysis agents such as DI water or ammonium chloride would also be interesting. A similar number of quenching and fixing agents can also be investigated. Once the lysis environment is optimized for erythrocyte lysis, the effects on leukocytes can be investigated. Damage to the leukocytes would be minimized due to the extremely short time spent in the lysis reagent. The results presented here could lead to methods that minimize nonspecific cell damage during lysis while maximizing the speed of the lysis step from minutes to less than a second.
Corresponding Author
*E-mail:
[email protected]. 2201
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