Centrifugal (CD) Microfluidic Platforms for Nucleic Acid Analysis
Jonathan Siegrist1, Guangyao Jia2, Horacio Kido2, Jim Zoval2, Gale Stewart3, Dominic Gagné3,
Régis Peytavi3, Ann Huletsky3, Michel Bergeron3, and Marc Madou2
1
Department of Biomedical Engineering, University of California, Irvine, CA 92697
2
Department of Mechanical and Aerospace Engineering,
University of California, Irvine, CA 92697
3
Centre de Recherche en Infectiologie de l’Université Laval, Centre Hospitalier Universitaire de
Québec (Pavillon CHUL), Sainte-Foy, Québec, Canada, G1V 4G2
ABSTRACT
While many diagnostic tools used in hospitals today are able to give results within one hour, the methods
currently used for microbial detection and diagnosis take on the order of days to complete. The concept of
molecular theranostics involves the rapid completion of these microbiology-based assays so that results can be
quickly obtained to give a more immediate and thorough picture of patient conditions, such that therapeutics can
accompany diagnostics. Here we report on the development of novel, automated, compact-disc (CD) based
platforms designed to quickly diagnose infectious diseases, making molecular theranostics a reality. Using a
combination of microfabrication and standard macro-machining processes, microfluidic CD platforms capable of
cell lysis and DNA purification, DNA amplification (PCR), and DNA hybridization/detection have been designed,
constructed, and tested. Total analysis time has been reduced from days to minutes, and because of the type and
amount of materials used, the platforms are also inexpensive. The systems developed meet the demands of rapid
theranostics and perform as well as and often faster than many of the tests and assays used today.
INTRODUCTION
Often upon admission to a hospital, a patient will undergo several tests to help determine their condition. Many
biochemistry and hematology-based tests, such as blood oxygen and pH, involve the processing of a sample in
an onsite laboratory. Radiology and other imaging techniques require taking the patient to an onsite center.
Regardless of the resources and manpower required to perform these diagnostics screenings, almost all of them
yield results within 1-2 hours that can be incorporated into the patient’s diagnosis and subsequent treatment. This
rapid incorporation of diagnostics and therapeutics, termed theranostics, is vital to quickly treating patients in
critical and/or unknown conditions.
Medical-based microbiology tests, however, fail to meet these theranostic requirements. When attempting to test
for and diagnose infectious diseases, a sample must often times be sent to an offsite facility for screening. The
standard tests used today involve growing and screening cell cultures, a process that can take over two days to
complete. When dealing with infectious diseases, this extended processing time can literally mean life or death for
a patient. Thus, a need exists for an infectious disease diagnostic platform that can perform an entire analysis
within one hour, to match the time scales of other diagnostic procedures and make true theranostics a reality.
While common infectious disease screening methods involve culturing cell colonies, other methods exist that
involve tapping into the genetic makeup of these organisms to more quickly specify the exact disease being dealt
with. Because theranostic methods must be rapid, the platform of choice for infectious disease detection should
be one that utilizes the genetic material of, for example, a blood-borne microbial organism present in the patient.
By analyzing the nucleic acid information, the type of microbe present is quickly determined without the need for
cell culture. Typically, a nucleic acid analysis involves the lysing of cells to release the small amount of genetic
material (DNA) present in the microbes, amplification of this DNA to a detectable concentration, and detection of
the DNA using hybridization methods. These tests can be performed in a well equipped molecular biology lab, but
using standard, manual protocols they can require many hours to complete, many skilled technicians to complete
them, and can leave behind a large amount of biological waste. By taking these nucleic acid analysis methods
down to the micro-domain and placing them on an automated platform, rapid infectious-disease diagnostics can
be performed in less time with minimal labor. In addition, the disposables of the system can be inexpensive to
make implementation and use economically feasible.
The Madou BioMEMS Research Group in collaboration with Génome Canada has chosen compact-disc (CD)
based centrifugal microfluidic platforms to meet all of these requirements and achieve true molecular theranostic
nucleic acid analysis. Indeed, CD platforms have been used in the past to efficiently automate analysis [1-4].
Using CD technology, rapid, inexpensive, and disposable nucleic acid analysis platforms have been developed,
with three separate platforms tackling the three main steps involved in genetic analysis. Here, a brief description
of the CD technology is presented, followed by an overview of the 3 nucleic acid platforms developed. More indepth descriptions of CD technology can be found in the literature [3-7].
CENTRIFUGAL CD TECHNOLOGY OVERVIEW
CD Fabrication
Before going into the theory behind how microfluidic CDs function, it is best to briefly review basic fabrication
processes and materials. Most commonly, the CDs consist of multi-layer structures made of inexpensive
polycarbonate plastic and pressure-sensitive adhesives (PSA). Using relatively simple CNC machines, channel
widths of down to 1 mm are machined into 1.2 mm or 0.6 mm-thick stock polycarbonate CDs. A computercontrolled cutter-plotter is used to cut channel widths as narrow as 200 µm in thinner materials such as 100 µmthick PSA or thin plastic films. Once the appropriate pieces have been designed and machined, they are aligned
centrally and radially and laminated together using the PSA layers.
The most simple, standard microfluidic CD
consists of no less than 5 layers: 1) top
polycarbonate CD with CNC-machined
sample loading, sample removal, and air
venting holes (sealed using a thin adhesive
film during operation), 2) pressure-sensitive
adhesive with channel features cut using a
plotter, 3) middle polycarbonate CD with
CNC channel features, 4) pressure-sensitive
adhesive with channel features cut using a
plotter, 5) solid bottom polycarbonate CD to
seal off the channels (Figure 1). Microfluidic
CD platforms can involve more layers to
accommodate more complex fluidics.
Moreover, different devices and substances
can be placed inside the CD during
fabrication, such as beads, lyophilized
reagents, or filters. The CDs can also be
exposed to O2 plasma treatment or
Figure 1. Schematic showing the assembly of a typical
functionalized with bovine serum albumin
5-layer microfluidic CD.
(BSA) to create hydrophilic and hydrophobic
surfaces, respectively. The fabrication process usually ends with running the CDs through an industrial press to
ensure excellent adhesion and sealing between all CD layers.
While the majority of CD platforms developed utilize standard macro-machining processes, microfabrication is
easily integrated onto the CD platform. This usually takes the form of creating microfluidic PDMS molds on 6” Si
wafers using multi-level, thick-resist lithography (Figure 2). Once created, these soft lithography PDMS molds can
be placed on a polycarbonate CD, or in some cases bound to glass slides and secured with a specially designed
slide holder CD. Almost all processes and materials used to create the microfluidic CD platforms are relatively
inexpensive, with the exception of the initial SU-8 mold needed when incorporating microfabrication. This allows
for the mass production of inexpensive parts that can be easily disposed of once contaminated with biological
materials. Once a CD platform has been developed, often only a simple motor and spin stand are needed for
operation. Using these varied fabrication procedures, a vast array of applications can be adapted onto the CD
platform to take advantage of the elegant CD pumping and valving mechanisms.
Pumping & Valving
When developing fluidic devices of any kind, the main concern is how to get liquids to and from the areas of
interest in a controlled manner. This general problem can be filtered down to the need for two technologies:
pumps and valves. Indeed, much of the work on microfluidic devices today focuses on the need for effective
pumps and valves, and is an ongoing area of research. The centrifugal CD platform provides elegant, simple, and
effective modes of pumping and valving.
Fluid propulsion on the CD is performed by centrifugally induced pressure on the fluid as the CD spins, and
Madou et al. and Duffy et al. have characterized this type of flow [3][4]. The volumetric flow rate, Q , of a fluid
element in a CD microchannel can be described by the following:
2
Q = ADb "# 2 r $r /32µL
(1)
where A is the channel cross-section area, Db is the hydraulic diameter
of the channel, " is the liquid density, " is the angular velocity of the
spinning!CD, r is the average distance of the liquid element from the
CD center, "r is the radial extent of the fluid, µ is the fluid viscosity,
! and L is the length of the liquid
! in the channel (Figure 3A). From
!
Equation
(1),
it
can
be
seen
that
the speed at which the disc spins, the
!
distance
from the center of the disc, the channel geometries and the
!
fluid
! properties all play a role in how! the fluid will move through the
microchannels.
By using combinations of different channel geometries
!
and spin speeds, flow rates ranging from 5 nL/s up to 0.1 ml/s can be
well controlled. Typical fluid-pumping rotation speeds used range from
300 to 2000 RPM.
!
Figure 2. An SEM image of a typical
2-level PDMS flow cell structure; the
microchannel is 25 µm deep and the
chamber is 250 µm deep.
Centrifugal pumping forces on the CD provide many advantages over some of the other pumping methods
available, such as syringe, peristaltic and electroosmotic. Current pressure-driven syringe and peristaltic pumps
provide very good control over large flow rates, but can be unwieldy when trying to miniaturize and/or multiplex
[8]. In addition, the pressures needed to move fluids through microchannels can be very large. This makes
implementation into small, high-throughput platforms difficult. Electroosmotic pumping methods overcome these
problems, as they can be easily adapted into microchannels using microfabrication. However, these methods are
highly dependent on pH and ionic strength of the fluid being pumped [8]. The extremely high-voltage power
supplies (>>1 kV) required in these systems make them expensive and unpractical. Centrifugal forces, however,
do not need large power supplies (only
a small motor) and are not dependent
on pH or ion levels of the fluid. They
also provide forces across the entire
length of a fluid element, providing
smooth and controlled flow. In addition,
many individual systems can be placed
on a single CD, making multiplexing
easy. When combined with the simple
valving solutions available on the CD,
powerful platforms can be developed.
Figure 3. Schematic of microchannels on a CD. A) Two reservoirs
connected by a single channel, B) Hydrophobic valve made by a
channel restriction in hydrophobic material, C) Hydrophobic valve
made by functionalization of the channel surface with hydrophobic
material, and D) Capillary valve made by channel widening.
Valving on the CD is performed using
two main valve types: hydrophobic and
capillary (Figure 3b-d). Hydrophobic
valves can take two different forms:
one utilizing changes in channel
geometries and the other utilizing
surface modification. In Figure 3b, it is
shown that when a channel quickly and
drastically changes from a wide to
narrow dimension, this can block the
flow of liquid. In Figure 3c, it is seen that a similar valving scheme can be obtained by functionalizing the channel
surface downstream of the fluid with hydrophobic materials. In both cases, the fluid can be forced past the
hydrophobic valve by increasing the spin frequency past a critical value.
The capillary valve is most commonly used in the CD platforms reported on here, and is a result of the balance
between centrifugal and surface tension forces. When fluid being pumped through a narrow channel by
centrifugal forces reaches an abrupt widening, as shown in Figure 3d, a large surface tension force develops at
that widening. If the surface tension force is greater than that of the centrifugal force, then the fluid flow will stop
even though the CD continues to spin. At a certain higher frequency, known as the burst frequency, the
centrifugal forces will overcome the surface tension forces and the fluid will continue down the channel. The point
at which burst frequency occurs can be described by the following:
Pm = "# 2 r $r > 4 % al sin(& c ) /Db = Pcb (2)
!
where Pm is the pressure at the meniscus (centrifugal force), " al is the surface energy per unit area of the liquidair interface, " c is the liquid contact angle, Pcb is the capillary barrier pressure (surface tension force), and all
other parameters are as!defined in Equation (1). Equation (2) assumes axisymetric channel cross-sections, but
can be adopted for different channel geometries and conditions. See Zeng et al. [9] and Duffy et al. [3] for further
development of these models. Equation (2) shows
! that by designing microfluidic structures with channels of
increasingly
smaller
capillary
valves,
control
of
when
a valve “opens” can achieved simply by increasing the
!
!
rotational speed of the CD. This has in fact been implemented on the CD platforms presented here to obtain
sequential control of valve openings. It is worth noting that none of the CD valving schemes presented act as
vapor valves – they perform liquid valving only. Thus, liquid reagents could not be stored for long periods of time
as evaporation and cross-chamber contamination could occur. While mixing is another vitally important aspect of
microfluidic devices, this topic will not be addressed here, but is reviewed in Madou et al. [5].
CELL LYSIS & DNA PURIFICATION PLATFORM
Intro
The first step in nucleic acid analysis for
infectious disease diagnostics is the
extraction of genetic material from the
microbes of interest. With microbial analysis,
the DNA of interest is located inside the cells,
and so the cell membrane must be penetrated
while keeping the DNA preserved. Once the
cells have been disrupted, other remaining
materials such as cell membrane pieces and
smaller vesicles must be removed from the
lysate. Various forms of cell lysis have been
developed and are in use, including
enzymatic lysis (cell membrane digestion via
enzymes), chemical lysis (cell membrane
breakdown via detergents), plasma lysis (cell
membrane disruption via electrical charge
pulses), and mechanical lysis (cell membrane
breakdown via physical means). Chemical
and enzymatic lysis methods leave behind
Figure 4. Schematic of a single cell lysis and purification
residual substances that can be hard to
mechanism.
remove or filter out, while plasma lysis can
cause damage to cellular structures. Of these
methods, mechanical cell disruption is the most effective at breaking down cells that have thick cell membranes
(Gram-positive) and successfully extracting intact DNA, with the added benefit of leaving behind no residual
substances [10][11].
Indeed, mechanical disruption is commonly used and comes in many different forms, such as homogenization,
French-press, and bead beating, to name a few. Almost all mechanical cell disruption methods rely on breakdown
of the cell membranes through shear forces. Bead beating is the most efficient method in this respect, and
functions by combining cells with an agitated mixture of milling beads [12][13]. As the solution is mixed, cell
membranes are disrupted by shear-forces developed when passing between beads as well as direct collisions.
This method can be easily controlled by altering the bead size and extent of agitation. A bead-beating cell lysis
platform has been implemented on a microfluidic CD platform. It utilizes small magnetic disks placed inside each
lysing chamber that oscillate while spinning by interaction with stationary magnets on the static CD platform. This
oscillation causes shear forces which result in lysis. Once lysed, solids are concentrated by centrifugation, and
the supernatant (containing DNA) is extracted using a unique siphon valve. The platform is capable of lysing cells
with thick membranes while preserving the extracted DNA. Moreover, the platform utilizes small volumes and
inexpensive disposables.
Materials & Methods
The cell lysis CDs were constructed using a similar 5-layer structure as described above (Figure 1). The center
disc thickness was 1.2 mm while the top and bottom discs were 0.6 mm-thick. The same 100 µm-thick PSA was
used. Six individual lysing mechanisms were placed on each CD, with each mechanism consisting of a lysis
chamber with ferromagnetic disk and milling beads, a siphon valve, and a collection chamber (Figure 4). The
ferromagnetic disks (V&P Scientific) used were 5.08 mm in diameter and 0.635 mm thick. The milling bead
mixture volume was 50 µL per chamber and consisted of a 4:1 slurry of 100 µm glass beads (Biospec) in a 1%
solution of polyvinylpyrrolidone in water. Midway through fabrication, the discs were subjected to O2 plasma
treatment to make the polycarbonate surfaces hydrophilic, and then the magnetic disks and slurry solution were
placed inside the CD. After finishing assembly, the CDs were run through an industrial press, and remaining water
was evaporated using a vacuum overnight.
The lysis CD spin-stand platform consisted of a servo motor, a driver, and
ToolPAC control software (Pacific Scientific). This platform included
permanent magnets (McMaster-Carr) fixed just below the CD surface to
move the in-CD magnetic disks during spinning (Figure 5). The placement
of these magnets was optimized using simulation and empirical observation
to ensure that the magnetic disk was drawn completely through the lysis
chamber during each pass.
To validate the system, E. coli cells (known to be relatively easy to lyse)
were incubated overnight at 37° C in LB medium (Q-Biogene/MP
Biomedicals) to an optical density of 0.98 (OD 600 nm). In addition, yeast
cells were also tested, as they are far more difficult to lyse [10]. Dehydrated
S. cerevisiae, Type II, were resuspended in 0.8% dextran in sterile water
(Sigma-Aldrich). The samples were loaded, and tests were run at 30, 60,
Figure 5. Schematic showing
120, 240 and 480 seconds at rotation speeds of 50, 200, 800, and 2000
the
mechanism
behind
RPM (n = 20). After lysing, the CDs were spun at 6000 RPM until all liquid
ferromagnetic disk movement.
was moved into the clarification chambers, and the residual cell debris was
pelleted at the bottom of the chambers (~1-2 mins). The CD was stopped to allow the siphon valves to prime
automatically due to hydrophilic capillary wicking, and then spun at 1500 RPM for 10 s to collect the supernatant
(containing DNA) in the collection chamber.
To further purify the DNA sample from the lysis CD for quantitative testing, a FastDNA Spin kit (Q-Biogene/MP
Biomedicals) was used according to the manufacturers protocol. DNA quantification was performed by taking 150
microliters of eluted, purified DNA and transferring it to individual wells of a Corning Half-Area 96-Well Clear FlatBottom UV-Transparent Microplate. Sample absorbencies were determined at 260 nm in a BioTek Synergy plate
reader (each plate contained an 8-well titration of bacteriophage lambda DNA (Promega) of defined
concentration). Unknown sample concentrations were extrapolated from a linear regression analysis of the
lambda DNA standard curve. Sample concentrations were confirmed by spot-checking random sample 260 nm
absorbencies in a Beckman DU-640 spectrophotometer.
Experimental Results
The CD lysis platform was successful in extracting intact DNA from both bacteria and yeast cells. The permanent
magnet placement allowed for optimal movement of the ferromagnetic disks, resulting in shear forces high
enough to lyse the thick-walled yeast cells while leaving genomic DNA in tact. The restriction channel dimensions
between the lysing and clarification chambers were ideal, such that liquid could be pumped without clogging while
leaving the milling beads behind. The siphon valve, used to draw off the supernatant, worked as expected. When
liquid enters the clarification chamber, wicking action due to the hydrophilic surface (recall the plasma treatment
during CD construction) draws liquid into the siphon valve up to a height equal to that of the liquid in the
clarification chamber. Centrifugal forces keep the liquid from wicking up and over the siphon valve channel. Once
the CD stops, the centrifugal forces are no longer present, and so the liquid can wick the rest of the way down the
channel until it reaches the capillary valve at a height lower than the bottom of the clarification chamber. At this
point, the siphon has been primed. Spinning the CD again causes the liquid to be pumped into the collection
chamber. Because the siphon channel begins at a point higher than the pellet mass, only the supernatant is
siphoned into the collection chamber. The entire cell disruption process takes less than 5 minutes.
Figure 6 shows DNA yields for the two different samples (bacteria and yeast) as a function of rotation time and
speed. As can be seen, DNA yields comparable to those of commercial bead beating mechanical cell lysis
systems were obtained [14][15]. While it is clear that the system must be tested and optimized for the particular
type of cell being lysed (as is true with any similar system), the cell lysis CD is capable of lysing cells that have
both thin and thick cell membranes. Moreover, the DNA extracted from the cell lysis CD is of high quality.
Relatively simple equipment is needed to run the system, and the CDs are inexpensive allowing them to be
disposable. The cell lysis CD platform is perfect for quickly creating near PCR-ready DNA samples from original
cell sources. Further details on the cell lysis platform can be found in previous Madou et al. publications [16][17].
Figure 6. Surface plots showing dependence of DNA yield on time and speed of lysis rotation
for bacteria cells (left) and yeast cells (right).
DNA AMPLIFICATION (PCR) PLATFORM
Intro
Extracting genetic material from a sample is only the first step in nucleic acid analysis. Because the diagnosis of
infectious diseases involves human samples, a cocktail will be obtained of genetic material from the patient, the
microbe(s) of interest, and any other microbes or viruses that might be present. In addition, the DNA of interest
will exist in very minute concentrations compared to that of the “background” DNA, and so the microbes’ DNA
must be isolated and amplified. Polymerase Chain Reaction (PCR) is the process most commonly chosen to
perform selective DNA amplification, and relies on thermocycling of a sample. A typical PCR thermocycling run
involves 30-40 cycles, with each cycle consisting of temperature changes from 95° C (denaturation) to 60 °C
(primer annealing) to 72 °C (extension).
PCR efficiency is most commonly evaluated in terms of temperature transition times and thermal homogeneity of
the sample, as these dictate whether or not a high-quality PCR product can be obtained in a small amount of time
[18]. Current bench-top PCR systems can take on the order of hours to complete a single run, significantly
hindering the speed of nucleic acid analysis. Microfluidics and microfabrication have been previously utilized to
create more efficient PCR systems [19-22], but many of these involve expensive fabrication processes and
materials, making disposable devices uneconomical. Here, a rapid, disposable-card based microfluidic PCR
system has been developed that has a small thermal mass resulting in rapid heating/cooling and excellent
thermal homogeneity. In addition, the materials and processes used to make the PCR cards are inexpensive. 30
cycles can be completed within 10 mins, and the resulting PCR product is of a detectable concentration. Although
no CD technologies are required for the PCR system, the same fabrication processes have been used to facilitate
future incorporation onto the CD.
Materials & Methods
Disposable PCR cards were created using inexpensive thin polycarbonate and adhesive materials. Using a
plotter-cutter, channel structures were cut in 250 µm-thick polycarbonate films that served as the middle layer of
the card. Sample loading/removal holes were cut in 127 µm-thick polypropylene PCR adhesive films (Excel
Scientific) to serve as the top layer, and the bottom card layer consisted of 70 µm-thick aluminum PCR adhesive
films (Excel Scientific). The 3-layer cards were aligned using a custom-made alignment assembly and bound
using the adhesives on the polypropylene and aluminum sealing films. Each card contained 4 separate PCR
chambers, with a single chamber volume of 25 µL (Figure 7). The card dimensions were optimized using
computer-based modeling of heat transfer.
Peltier thermoelectric devices (Ferrotec) were chosen to perform active heating and cooling of the PCR cards. A
copper sheet 0.44 mm-thick was placed across the surface of each Peltier device to increase spatial thermal
homogeneity. T-type thermocouples (Omega) were imbedded inside the copper sheet to provide feedback to a
custom multi-level PID control system implemented using National Instrument’s LabVIEW. Both the copper sheets
and T-type thermocouples were secured using thermally-conductive adhesive compounds (Artic). Programmable
power supplies with polarity-switching capabilities (Agilent Technologies) were used to power the system and
interface with the LabVIEW control program. The entire platform was designed such that two PCR cards could be
accommodated, for a total throughout of 8 independent PCR reactions (Figure 8). A custom-made clamp was
used to ensure good thermal contact between the cards and thermoelectric devices. Thermal behavior of the
entire system was characterized in-situ using thin-film T-type thermocouples (RdF Corporation) inserted into the
reaction chambers in between the PCR card layers.
Keeping future CD integration in mind, it was realized that not only will a liquid valve need to be incorporated, but
a vapor valve will be required during thermocycling as well. Thermocycling causes vapors to accumulate as the
liquids come close to boiling temperature. Large pressures will build up as a result causing the samples to
expand, and so the entire chamber needs to be sealed and isolated. With Peltier devices already integrated onto
the PCR platform, a novel ice-valve scheme was chosen. Using Peltier devices, a small plug of liquid can be
frozen on each end of the reaction chamber, thus preventing the movement of both liquid and vapor during
thermocycling. This idea was integrated onto the PCR platform as well.
Finally, validation of the system’s PCR abilities was performed. Briefly, amplification of a 210 base pair region of
the E. coli tuf gene was performed at three concentrations of purified DNA ranging from 10 to 1000 E. coli ATCC
35401 genomic copies. Teco553 and Teco754 forward and reverse oligo primers were used, respectively. Taq
TM
polymerase (Promega) and TaqStart antibody (Clontech) were used in 10X PCR buffer (Promega). The PCR
mixture and a negative control (without a DNA template) were loaded into separate reaction chambers on a card
TM
using a pipette. Hot start was achieved through denaturation of the TaqStart antibody at 95º C for 1 minute. A
40-cycle amplification was conducted using a temperature profile of 7s, 15s, and 15s (ramping times included) at
95º C, 60º C and 72º C, respectively. After cycling, the samples were unloaded from the chambers and 1.0 µL of
sample was injected into an Agilent BioAnalyser 2100 cartridge (Agilent Technologies) for verification via gel
electrophoresis.
Figure 7. A) Layout of PCR card
channels, B) Schematic of PCR card
assembly made of 1-top polypropylene
film w/access holes, 2-polycabonate
machined layers, 3-aluminum bottom
film, and 4,5-thermocouples.
Figure 8. A) PCR platform
consisting of Peltier devices for
heating/cooling (H1, H2) and icevalving (C1-C4);
inset
shows
thermocouples imbedded in Peltier
device, B) Thermocycler with 2 PCR
cards loaded and ready.
Experimental Results
The PCR platform achieved very rapid heating and cooling ramping speeds of up to 10° C/s. Because the Peltier
devices allow active heating and cooling based on the polarity of voltage used, shorter thermocycling times were
achieved as compared to standard PCR systems that rely on active heating only with passive cooling. A single
cycle time of 20 s was achieved, resulting in a total time for a 40-cycle run of 10 mins (Figure 9).
The card was designed such that the thermal mass of the liquid being
heated would be very low and the surface-area-to-volume ratio would
be very high. This resulted in a very good thermal homogeneity
throughout the liquid sample, as measured by the thin-film
thermocouples. At the 60° C and 72° C temperatures, the thermal
gradient measured from bottom to top of the liquid sample was as little
as 0.2° C. However, at 95° C the thermal gradient measured was
almost 2° C. The reason for this is the higher temperature difference
between the surrounding air and the PCR card, resulting in
disequilibrium on the time scales involved in PCR. It was observed that
the same 0.2° C gradient resulted when the system was allowed to
reach equilibrium at 95° C. However, the desired temperature range for
DNA denaturation is from 91°-97° C, so the large temperature gradient
should not affect the PCR product.
Figure 9. Typical PCR
cycling profile.
thermo-
!
Figure 10. Electrophoresis
gel: 1,6 ladder standards, 2
negative
control,
3-5
amplified products starting
with 10, 100, and 100
genome copies, respectively.
The copper sheets applied to the Peltier device surface reduced the
spatial thermal differences to less than 0.2° C, and temperature control of
± 1° C was obtained using the multi-level PID feedback control system.
The ice valves performed flawlessly. No leaks were observed, and the
liquid plugs froze within 15 seconds. A disadvantage of the ice-valves is
that part of the sample is prohibited from being thermocycled, resulting in
a dead volume. In this case, however, the dead volume per reaction
chamber was only 2 µL, or 8% of the total chamber volume.
From the PCR validation experiments, it was observed that as few as 10
copies of the E. coli tuf gene could be successfully amplified to a
detectable amount (Figure 10). The results show not only that
amplification was successful, but also that the specificity was excellent,
as no other bands appeared on the electrophoresis gel. In addition, a 27minute unoptimized thermocycling profile was used for this test. Thus, a
microfluidic PCR system capable of producing high quality products in a
small amount of time was implemented using inexpensive, disposable
parts.
DNA ARRAY HYBRIDIZATION PLATFORM
Intro
The final step in nucleic acid analysis is detection of the DNA of interest, most often performed using hybridization
methods. Specifically, DNA hybridization microarrays can be used when attempting to detect multiple sequences,
as when screening for many infectious diseases. Briefly, custom-made oligonucleotide capture probes (specific to
the microbe of interest) can be immobilized on a solid support. This can be done, for example, by using amineterminated probes reacting with functionalized glass slides or thiol-terminated probes reacting with gold surfaces
[23]. Once an array of different capture probes has been immobilized on a surface, the sample DNA can be
washed across the array. If a complementary sequence exists, hybridization will occur, and transduction of the
hybridization signal can be performed using optical or electrochemical means.
The rate-limiting step when performing analysis with DNA microarrays is the time for hybridization to occur, which
can take upwards of 18-24 hours [24-26]. Enough time must be given to the DNA of interest to passively diffuse
through the entire solution and be exposed to every capture probe on the surface. By using narrow, microfluidic
chambers, hybridization time can be significantly reduced by making the diffusion distances much smaller.
Moreover, by incorporating active flow, mass-transport of sample DNA to the capture probes can be increased
further. Finally, by automating the entire process, these tests can be performed quickly without the need of
expensive, skilled labor.
Using microfabrication, a PDMS microfluidic device for DNA microarray assays has been developed, and this
device has been incorporated onto the CD for a rapid, flow-through hybridization assay. Optical signal
transduction was implemented by the use of fluorescently tagged DNA. The system showed strong fluorescent
hybridization signal within a short 15 min assay time.
Materials & Methods
The microfluidic devices were made of PDMS using a two-level SU-8 mold. A 6” Si wafer was patterned with
25 µm-thick microchannels using SU-8 25 (Microchem), and reagent chambers 250 µm-thick were patterned
using SU-8 100 (Microchem). PDMS base and curing agent (Dow Corning) were mixed in a 10:1 ratio by weight,
and poured on the SU-8 mold. After degassing, the PDMS was cured on a hotplate at 100° C for 10 mins. Once
cured, the PDMS pieces were peeled off, cut out, and access holes for reagent loading were punched through.
Functionalized glass slides were used as the substrate for capture probe immobilization, allowing for easy
fluorescence scanning using a commercial slide reader. The glass slides were modified with amine groups
following work from Joos et al. [27]. Once functionalized, 4 different types of 25-mer capture probes were arrayed
onto the slides using a spotter. The captures probes corresponded to 4 different bacteria of interest, namely
Staphylococcus aureus, S. epidermidis, S. haemolyticus, and S. saprophyticus. The captures probes were
synthesized with a 5’ amine modification (Biosearch Technologies) to allow covalent bonding to the functionalized
slides. The microarrays were fabricated following work from
Shena et al. [28]. 10 µM capture probe solutions in TE buffer
were diluted 2-fold in Array-it Microspotting Solution Plus
(Telechem International). Using a Virtek SDDC-2 arrayer
(Bio-Rad Laboratories) equipped with SMP2 pins (Telechem
International), spots of 0.6 nL volume and 60-80 µm diameter
were placed on the glass slides. Duplicates of each of the 4
capture probes were spotted. The PDMS parts were then
placed on the glass slides, using visual, manual alignment to
ensure the hybridization chambers lined up with the capture
probe spots. No O2 plasma treatment was used to bond the
PDMS to the glass - the passive sealing properties of PDMS
on glass were enough for this application. The slides were
then placed into a specially designed slide holder CD to
accommodate 5 slides and placed on the same spin-stand
set-up as used for the cell lysis CD (Pacific Scientific). Figure
11 shows a schematic of the entire DNA microarray CD
platform.
The test sample solutions were created using PCR in the
presence of fluorescently-tagged nucleotides, specifically
Cy-5 dCTP nucleotides. This work was done according to
previously established methods [29][30]. Four different
solutions were created to correspond to the 4 different
capture probes. Finally, hybridization tests were performed
using the flow-through CD platform. The capture probe spots
were exposed to target sample solution (10 nM), a wash
buffer (2X SSPE + 0.1% SDS) and then a rinse buffer (2X
SSPE). The CD platform spin speeds used were 375 RPM,
850 RPM, and 1175 RPM to release the 2-µL target sample,
the 10 µL wash buffer, and then the 10 µL rinse buffer
solutions, respectively. After spinning, the PDMS channels
were removed, and then the slides were dried and read using
a ScanArray 4000XL (Packard Bioscience Biochip
Technologies).
Figure 11. Schematic of the entire DNA
microarray CD platform. A) The microfluidic
PDMS device, with 1-hybridization chambers
with spotted capture probes, 2-target sample
chamber, 3-wash buffer chambers, 4-rinse
buffer chambers, 5-flow channel. B) Close up
of A-1, showing the capture probe spots (units
in microns). C) The entire CD with 5 PDMSslide devices.
Experimental Results
It was found that the CD DNA array platform could perform
an entire flow-through experiment in less than 15 minutes.
The PDMS device worked as desired, and by controlling the
channel geometries and spin speeds, ideal flow rates were
obtained, from 0.38 to 3.75 µL/min. This allowed for slow flow
of the target sample across the capture probes to allow
efficient mass transport, followed by rapid flow of the rinse and wash solutions to remove the non-specifically
bound DNA probes with higher pressures. The burst frequencies for the different capillary valves used were
consistent across all PDMS pieces used (data not shown), and the sealing of the PDMS on glass allowed
adequate containment of the fluid during flow through while still allowing the pieces to be removed for
fluorescence scanning in the commercial slide reader.
The system was able to detect the differences between the 4 different bacteria of interest, and the specificity was
high (Figure 12). Moreover, the signal intensities were high, verifying that efficient hybridization did occur due to
the gains in mass transport from a microfluidic flow-through system. Again, this was obtained using a
hybridization time of less than 15 minutes, as opposed to the hour time scales normally required of many passive
hybridization systems. The materials needed were relatively inexpensive, allowing for disposal of both the slides
and PDMS devices. In addition, the entire assay was automated. Further details on the DNA array CD platform
can be found in previous Madou et al. publications [31-34].
Figure 12. Fluorescent scanning results of the 4 different capture probes, showing good specificity
and signal strength.
CONCLUSIONS
There is a huge need for more rapid microbial detection and analysis such that theranostics can become a reality
for use in hospitals and clinics. Here, we have reported on three platforms that allow for rapid nucleic acid
analysis. Using microfluidic CD technology, platforms for cell lysis and DNA purification, PCR, and DNA array
detection have been developed. The cell lysis platform allows for rapid (5 min) mechanical cell disruption using
bead-beating methods, and has proven to lyse thick-membrane yeast cells in a very short time. The PCR platform
allows rapid (10 min) thermocycling and can perform successful amplification starting with only 10 genomic
copies. The DNA array platform can perform flow-through hybridization is less than 15 minutes with resulting
strong and specific optical signals. Moreover, all of the platforms developed utilize inexpensive and easy-tomanufacture materials and methods, such that the devices can be disposed of after use. The platforms
themselves use simple motors, drivers, power supplies, and PC computers for automated control, thus reducing
the labor needed to perform these assays. It has been shown that automated, rapid, and inexpensive nucleic acid
analysis is possible and feasible.
The next steps will be to incorporate the three individual platforms already developed into a single, automated
CD-based platform. At this point, much of the science has been addressed, and now engineering must be used to
put the platforms together. Towards making the CD platforms a single product, reagents will need to be stored on
the CD, such that the only labor needed is the loading of the CD with a sample. Lyophilized reagents could be
used, or wax valves could be incorporated for long-term storage of liquids [35]. Moreover, injection molded CD
devices would be desired, so they can more easily be mass-produced. To this end, injection molded cell lysis CDs
and PCR cards are already in production for their respective individual platforms. The nucleic acid CD analysis
platforms have been extremely successful, proving that when practical engineering and manufacturing is
combined with the miniaturization sciences, problems can be effectively and efficiently solved.
ACKNOWLEDGEMENTS
The authors gratefully thank Génome Canada-Génome Québec for funding of the included projects. We would
like to thank Nahui Kim and Chengwu Deng for their contributions on these projects. In addition, contributions
were made from RotaPrep and Rich Welle and John Anderson from Phasiks.
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