A microfluidic fluorescence measurement system
using an astigmatic diffractive microlens array
Ethan Schonbrun,1,2,* Paul E. Steinvurzel,1 and Kenneth B. Crozier1
1School
of Engineering and Applied Science, Harvard University, Cambridge, MA 02138, USA
Institute at Harvard, Harvard University, Cambridge, MA 02142, USA
*schonbrun@rowland.harvard.edu
2Rowland
Abstract: We demonstrate an opto-fluidic detection system based on an array of
astigmatic diffractive microlenses integrated into a microfluidic flow focus
device. Each astigmatic microlens produces a line excitation across the channel
and collects fluorescence emission from the linear detection regions. The linear
excitation spot results in uniform excitation across the channel and high time
resolution in the direction of the flow. Collected fluorescence from each
integrated microlens is relayed to a subregion on a fast CMOS camera. By
analyzing the signal from individual micr o len ses, we d e mo nstr ate co unti n g
and r eso lu tio n o f 5 0 0 n m a nd 1 . 1 µ m beads at rates of up to 8,300
per second at multiple locations. In addition, a cross-correlation analysis of the
signals from different microlenses yields the velocity dispersion of beads
traveling through the channel at peak speeds as high as 560 mm/s. Arrays of
specifically designed diffractive optics promise to increase the resolution and
functionality of opto-fluidic analysis such as flow cytometry and fluorescence
cross-correlation spectroscopy.
©2011 Optical Society of America
OCIS Codes: (050.1965) Diffractive lenses; (120.7250) Velocimetry.
References and links
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1. Introduction Flow cytometry is a powerful method to optically analyze biological materials in
fluids [1]. In
part because of its success, producing a microfabricated flow cytometer has become one of the
major goals of microfluidic technology [2]. Flow cytometers obtain quantitative analysis by
precisely controlling the intersection of a fluid borne sample stream with a laser focal spot.
Commercial table-top systems are capable of extremely high throughput detection by collecting
fluorescence and scattering data on tens of thousands of cells or beads per second.
Building on recent progress in microfluidics, there have been several successful
implementations of microfabricated flow cytometers using lithographically patterned fluidic
structures [3, 4]. Although microfabricated flow cytometers frequently rely on traditional
microscopes for optical detection, there are clear advantages in integrating the optical components
as well. Microoptical components have dimensions on the same order as microfluidic structures.
Consequently, arrays of both systems can be implemented on the same chip to realize higher speed
through parallelization [5, 6]. Another advantage of integrating optical and fluidic systems onto the
same chip is that their alignment is defined lithographically and then fixed during fabrication.
Alignment of the sample stream to the laser spot is critical in flow cytometry, and as future devices
continue to shrink and parallelize these intersections, off chip alignment will become increasingly
more challenging.
The primary method to integrate optical detection into microfluidic devices has been to use
optical waveguides. Optical fibers can be easily integrated into planar microfluidic devices and
are very effective at delivering light from lasers and coupling collected light to photodetectors
[7– 9]. One major drawback of waveguide optical excitation, however, is that there is limited
control of the excitation field distribution. The field emitted by fibers lying outside the fluidic
channel diverges before it intersects the detection region and frequently exce ed s a wid th gr
ea ter tha n 5 0 µ m, wh ich se ver el y limits spatial resolution. To help decrease spot sizes
and increase local intensity, planar lenses patterned into the channel walls have been used to
focus excitation fields from both fibers [10 ] and LEDs [11]. Although these planar lenses have
resulted in greater signal strength, they have not demonstrated focusing tig hter tha n 5 0 µ m.
I n ad d itio n to li mited excitat io n f i eld co ntr o l, in -plane lenses are inefficient light
collectors. For collecting Rayleigh scatter, efficiency is not crucial, but for collecting weaker
fluorescence signals, high numerical aperture (NA) two-dimensional lenses have much higher
collection efficiency.
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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Fig. 1.
Schematic of the optofluidic fluorescence measurement system. The microfluidic
channel and microlens array are integrated onto the same substrate. The microlens array is illuminated
by a broad fluorescence excitation laser beam and imaged onto a high speed CMOS camera by off
chip relay optics that have unity magnification. The fast CMOS camera is a Phantom V7.1 from
Vision Research. Image stacks are collected by the camera and later analyzed to quantify the
fluorescence signal collected from each detection region.
Instead of in plane integration of optical elements, we demonstrate the use of an array of
diffractive microlenses that operate through the top wall of a channel. The device is operated in an
epi-illumination mode, where both the excitation and emission pass through the same diffractive
microlens on the same side of the sample, as shown in Fig. 1. Each microlens has an NA of 0.73,
which produces a tightly focused excitation spot and efficiently collects fluorescence emission. In
addition to large focusing power, astigmatism is added to the microlens design so that the excitation
focal spot is formed into a line that runs perpendicular to the channel direction. Line excitation is
used in table-top flow cytometers [1] and also has been used in single molecule detection [12 ], so
that analyte traveling through the flow focused stream intersects the same intensity at different
cross-sections. Reducing the coefficient of variation for homogenous samples is extremely
important in order to quantitatively compare the fluorescence amplitude from each particle.
The organization of this paper is as follows. In section 2, we first introduce a mathematical
description of the astigmatic microlens and definitions of the desired focal spot distribution. The
microlenses are then experimentally characterized both in a bulk fluid reservoir and aligned to a
single microfluidic channel. Section 3 shows the results for counting homogenous solutions of
fluorescent beads at low and then high throughput. The coefficient of variation is also discussed for
this homogenous sample. Section 4 demonstrates the resolution of bead size for a heterogeneous
sample and compares the results for each of two detection regions. Finally, section 5 presents a
multiple field of view velocimetry technique using the astigmatic microlens array.
2. Diffractive optic design and fabrication Diffractive elements rely on the geometry of a grating
pattern to modify the phase of
diffracted waves. We use electron beam lithography to pattern microlens masters, and
consequently we have extremely precise control over the local grating geometry [13 ]. By
contrast, it is much more difficult to control the geometry of refractive microlenses that require
the three-dimensional fabrication of curved surfaces. Consequently, the focal length of diffractive
microlenses can be defined with much higher precision than refractive microlenses [14], which is
critical for microlens arrays that are lithographically aligned. In addition, using techniques from
computer generated holography; diffractive microlenses can be designed that produce almost
arbitrary intensity distributions in their focal plane.
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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The diffractive microlenses are designed by first calculating the complex field distribution of the
desired focusing wave at the cross section of the microlens, which is spaced by the focal length
from the microfluidic channel. In this case, we want the focusing wave to form a line excitation
across the channel. This can be produced by the multiplication of a spherical wave with a
cylindrical wave,
medium, f
where w
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2
l length, and f1
2
2
2
b
and purely determined by the spherical wave term. The front focal length (ff
2
1
e
x
p
(
)
e
x
p
(
(
w
2
f
)
f
R
f b f b,
f
)
,
o
i
s
t
h
e
s
p
h
e
r
i
c
a
l
Fig.
2. Astigmatic diffractive microlens. a) Microscope image of SU8 lens master. b, c)
Experimentally-observed focal spot intensity distribution at front and back focal planes,
respectively, taken with a 50× objective lens.
w
a
v
e
f
o
c
a
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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ADW jk x y f jk x f
(1) where ADW is the astigmatic diffracted
wave, k is the wavevector of light in the focusing
o is the cylindrical wave focal length. Imaging systems with a large amount of astigmatism produce
two focal spots that are shaped into lines that are orientated perpendicular relative to each other. The
back focal length (f) is located at fo) is located at
1
1
o
o
f
fff
due to the fact that rays offset in x are focused by both the spherical and the cylindrical
terms. At the front focal plane, the intensity distribution is elongated along
y and diffraction limited in x. Using ray optics, the length of the excitation line at the front focal
plane can be estimated by,
is the half width of the major axis of the elongated focal spot, and R is the radius of the diffractive
lens. The width is defined by the angle of the marginal ray focused by the diffractive lens, which
also determines its NA. Because the focal spot is still Gaussian in both directions, the width of the
spot should be designed to be greater than the experimental detection region, resulting in uniform
illumination.
After the desired phase profile of the focusing wave has been calculated using Eq. (1), we apply
a thresholding function that transforms the continuous valued complex field into two d iscr ete p
hase va lue s. T he thr esho ld f u nctio n i n t his ca se se ts an y p ha se val ue fr o m [ 0 , p) to
0 , and [ p, 2 p) to 1 [ 15]. Two level phase diffractive elements have a maximum diffraction
eff icienc y o f 4 0 . 5 % when t he r elative p ath len gt h b et wee n t he t wo le vels has a d if fer
ence o f p [16 ]. The refractive index contrast in our microlenses is between PDMS and air and has
a value of 0.45. We are interested in optimizing the diffraction efficiency for fluorescence emission
with a wavelength of 575 nm, so the ideal thickness of the diffractive elements is 640 nm.
The designs are patterned into SU8 using electron beam lithography to form microlens masters
that can then be molded into PDMS. While it is possible to pattern thin films of SU8 with
features as small as 30 nm using electron beam lithography [17], thicker films are more
challenging. For a film of SU8 that has a thickness of 450 nm, we have found that we can reliably
produce gratings that have a line width of less than 500 nm and a pitch of 900 nm.
f
First, SU8 2000.5 is spun onto an Indium Tin Oxide coated microscope slide at 3000 RPM. We use
an Elionix ELS-7000 100 kV electron beam lithography system for the exposure. The pattern is
written at 20 pA and with a d o sage o f 4 . 4 µ C/c m. SU8 has a much lower exposure
threshold than other electron beam resists, so consequently larger patterns can be written in smaller
time.
Using an atomic force microscope, we measure that the resulting SU8 pattern has reasonably
straight sidewalls and a depth of 400 nm. Due to electron scattering in the relatively thick resist, it is
difficult to obtain deeper features. However, a thickness of 400 nm is enough to produce good
diffraction efficiency. Using a home built beam propagation algorithm, we estimate that the
diffraction efficiency for the fluorescence excitation with a wavelength of 532 nm is 0.34 and the
diffraction efficiency for the fluorescence emission with a wavelength of 575 nm is 0.31. These
values are somewhat lower than the ideal d iffr ac tio n ef ficie nc y o f a p -phase grating of 0.41,
but are still more than three times larger than the diffraction efficiency of an amplitude grating,
which is 0.10.
Fig. 2(a) shows an SU8 diffractive microlens master that has fo = 2 1 0 µ m, f = 1 0 5 0 µ m, and
2R = 1 9 5 µ m. T he r esulting ff1 is 1 7 5 µ m, wh ich is c h o sen b ec ause the micr o len ses
ar e d esig ned to fo cus t hr o u gh a N o . 1 co ver slip that is 1 7 0 µ m t hick. Fro m Fig.
2(a), we can see that the normally circular Fresnel zones are slightly elongated in the vertical
direction as a result of the added astigmatism. After the microlens masters have been fabricated, the
surfaces are treated with a salinization layer. Then PDMS is poured onto the SU8 masters and baked
at 65°C overnight.
To visualize the focal spot distribution, the microlenses are aligned to a fluorescent dye
(Resorufin) filled reservoir. The microlenses are placed face down on a No. 1 coverslip which acts
as the roof of the reservoir. A single microlens is then illuminated by the fluorescence excitation
laser. From the opposite side of the reservoir, we image the fluorescence emission using a 50×
microscope objective and a fluorescence bandpass filter centered at 575 nm. Images of the emission
at the front and back focal planes of the astigmatic microlenses are shown in Fig. 2(b), and (c),
respectively. The front focal plane lies very close to the coverslipfluid interface. This is the focal
volume that we will use to define our detection region as, by contrast, the back focal plane lies
beyond the microfluidic channel.
Fig. 3(a) shows the completed device, where three astigmatic microlenses are aligned and
reversibly bonded to a microfluidic channel downstream from a flow focus junction. The sample
enters the middle inlet channel on the left and is hydrodynamically focused by two shea t h flo w
c han nel s o n t he t o p and b o tto m. T he main inte r r o gatio n cha n nel is 3 0 µ m
wid e and 6 µ m d ee p . Eq. (2) p r ed icts t hat t he excita tio n sp o t is 3 2 µ m wid e
in t he ver tical d ir ec t io n of Fig. 3(a) and consequently slightly overfills the width of the
main channel.
2
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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Fig. 3. Optofludic
device. a) Brightfield microscope image, taken with a 10× objective lens, of
the flow focus microfluidic device aligned to the diffractive microlens array. The diffractive microlen
ses a re sep a ra t ed from th e mi c roflu i di c ch an n el b y t h e c oversli p thi ck n ess of 170
µm, so they appear slightly out of focus. b) Fluorescent emission resulting from the diffractive lenses
focusing the excitation laser into the dye-filled channel, showing the focal spot distributions of all
three detection regions.
Fig. 3(b) shows an image of the induced fluorescence after flooding the device with Resorufin
and illuminating all three diffractive microlenses with the fluorescence excitation laser. The three
micr o le nses p r o d uce thr ee e xcitatio n li nes s ep ar ated b y 2 0 0 µ m tha t tr ave r se
the channel cross section. From the fluorescence images, the full width at half maximum ( FW HM
) sp o t size o f ea c h fo ca l r egio n is fo und to b e 7 µ m alo ng the cha n nel d ir ec tio n
. Due to diffraction of the excitation beam through the finite depth of the channel, the width of
the excitation spot is significantly larger than the diffraction-limited size of the focal spot, but still
less than one quarter of the channel width.
3. Bead counting Fluorescent beads are commonly used as calibration standards in flow cytometry
because,
among other reasons, they can be made relatively monodisperse. We use polystyrene beads from
Invitrogen that have a diameter coefficient of variation (COV) of 4.5%. The beads have Nile Red
fluorescent dye mixed into their volume, so assuming that all the beads have the same dye
concentration, we expect that the COV of the fluorescence signal from each bead to be 14%. Fig.
4(a) shows a one second trace, collected at 4,000 frames per second (fps), of the signals from
each of the three different diffractive lenses along the microfluidic channel. The plotted signals
are obtained by integrating the fluorescence intensities over the pixels corresponding to each
diffractive microlens in the CMOS camera image. In this experiment the sa mp le co ns ist s o f
a su sp ensio n o f 2 µ m b ea d s at a co nc entr atio n o f 0 . 1 % b y wei g ht i n d e
ionized water, while the sheath fluid is de-ionized water. The sample stream is driven by a
syringe pump at a r ate o f 2 0 µL/ ho ur and t he s hea th fl uid is d r iven at a r ate o f 5 0
µ L/ho ur .
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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Fig. 4.
Bead counting at multiple detection regions. a) Time trace sampled at 4,000 frames per
second (fps) of signals produced at each of three detection regions (DR), corresponding to each of
three diffractive microlenses. Inset shows a higher resolution time trace of seven beads as they flow by
the three detection regions. b) Histograms of the peak heights for each of the three different detection
regions.
From Fig. 4(a), we observe that each bead produces a pulse as it passes by a detection region
because the short focal length diffractive microlenses have well-defined collection volumes
analogous to those of confocal microscopes [18 ]. The relative delay between consecutive
detection regions is approximately 1.5 ms, corresponding to an average velocity of 130 mm/s.
Using an algorithm that counts peaks from the signal traces, we find that over a five second time
span, detection regions 1-3 counted 447, 441, and 440 beads respectively. Fig. 4(b) shows
histograms of the peak heights for each detection region, from which COVs of 0.27, 0.21, and
0.22 can be found. We estimate that a COV of 0.14 can be attributed to bead volume dispersion
as estimated from the manufacturer. We believe the remaining variation to be attributed to the
detection system, primarily caused by the intensity gradient along the depth of the channel. More
sophisticated microfluidic designs are capable of flow focusing in the depth direction as well [2],
and this could further reduce the measured COV. At this flow rate and bead concentration,
counting occurs at 90 beads per second. However, the bead concentration can be considerably
increased before coincidence rates become detrimental. For the data shown in Fig. 4, ca mer a fr a
me s ar e ca p tur ed ever y 2 5 0 µ s, o ver wh ic h ti me the b ea d s tr avel a n aver age o
f 3 2 µ m.
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Received 3 Nov 2010;
revised 15 Dec 2010;
accepted 22 Dec 2010;
published 12 Jan 2011
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Fig. 5. Time
resolution characterization. a) Time trace sampled at 40,000 fps of signals
produced at two detection regions. Inset shows a higher resolution time trace of the signal from two
beads. b) Autocorrelation of each detection region, yielding a half width at half maximum of 2 3 a nd
31 µs, resp ec ti vel y. For a v eloc i t y of 9 0 mm/ s, b ead s t ra vel a n avera g e of 2 . 3
µm in between frames.
In addition to excitation uniformity, the line focus of the astigmatic lenses enables high time
resolution because the excitation spot is narrow in the flow direction. To explore the temporal
resolution, we need to increase the sampling speed of the camera. Fig. 5 shows the time tr ac e fo r
1 . 1 µ m b ea d s tr aveli ng p ast t wo co n sec u tive astig mat ic micr o len ses, wher e the
ca mer a ca p tur e r ate ha s b ee n incr ea sed to 4 0 , 0 0 0 fp s, o r a fr a me e ver y 2 5
µ s. T he sa mp le str ea m is d r i ven a t 2 0 µ L/ ho ur and t he s hea t h fl uid is d r ive n
at a r ate o f 4 0 µ L/hour, resulting in an average velocity of 90 mm/s. With the faster camera
sampling rate, we can see in the inset of Fig. 5(a) that the signal pulses now consist of multiple
samples.
The autocorrelation of the signal from each detection region shows the average time that a bead
spends in the excitation region of each lens. The half width half maximum for the two
autocorrelation peaks, shown in Fig. 5(b), ar e 2 3 and 3 1 µ s, r esulti ng i n f ull wid th s o f 4 6
and 6 2 µ s. B y mult ip l yi n g the b ea d velo cit y b y the fu l l width of the autocorrelation
peaks, we exp er i me ntall y d eter mi ne th e wid t h o f the e xcitatio n r egio n s to b e 4 .1 and
5 .6 µ m, respectively. These values are similar, but slightly lower than the width of the
excitation spot ( 7 µ m) , mea s ur ed b y flo o d ing the ch annel with fluorescent dye. We
estimate that the smaller width is due to the fact that micron sized beads probe the intensity
distribution closer to the center of the channel where the excitation is more tightly focused. In
comparison, the much smaller Resorufin molecules from the previous characterization probe the
entire channel depth, which include regions close to the channel walls that are further away from the
focal plane.
4. High throughput sizing The real power of flow cytometry lies in its ability to quantitatively
measure optical signals of
heterogeneous samples at high speeds. To demonstrate this functionality, we load a mixture
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
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of 500 nm and 1. 1 µ m d ia me ter b ea d s into t he d e vice, at a co nce ntr at io n b y mas
s o f 3 *1 0 5 4 and 2*10, respectively. To obtain high throughput, we drive the sample and sheath
flow at a r ate o f 1 0 0 and 3 0 0 µ L/h and o p er ate the ca mer a at clo se t o its ma xi mu
m sp ee d at 1 0 0 , 0 0 0 fps. Fig. 6(a) shows the time trace for two consecutive detection regions.
We can observe two distinct peak heights in the trace, corresponding to the difference in brightness
of the 500 nm and 1 . 1 µ m b ea d s. T he histo g r a ms i n Fig. 6(b) and (c) show clearly the
resolvability of the two sizes of beads at each detection region. The distributions in the histograms
are plotted as a function of the cube root of intensity because the numbers of fluorescence
molecules approximately scale with the volume of the bead . T he hist o gr a m p ea k val ue fo r
the 1 . 1 µ m beads is slightly greater than twice the peak value for the 500 nm beads, which is
consistent with the expectation that the fluorophore concentration is the same for each bead size.
Fig. 6.
High speed size discrimination. (a) Time trace sampled at 100,000 fps of signals
p rod uc ed a t t wo d et ec t i on regi on s for a mi xt u re of 500 n m an d 1. 1 µm b ead s. (b ) a
nd (c ) Histograms of the peak heights at the first and second detection region, respectively, plotted
as a function of the cube root of peak intensity.
The data shown in Fig. 6 is collected for one second, during which 8359 beads are counted in
the first detection region and 8248 beads are counted in the second detection region. By gating the
histogram in between the two peak values, we determine that the first detection region counts 6790
beads with diameter 500 nm, and 1569 beads with diameter 1.1 µ m. T he sec o nd d etec tio n r
eg io n co un ts 6 6 9 8 b ea d s wit h d ia meter 5 0 0 n m, a nd 1 5 5 0 b ead s wit h d ia
meter 1 . 1 µ m. Fo r the 1 0 µ s exposure used here, we measure a signal to noise ratio of 8
for the smaller 500 nm beads. A threshold of one half the mean signal from a 500 nm bead is used
to discriminate beads from background noise, which corresponds to 4 times the background noise
standard deviation.
5. Cross correlation velocimetry I n ad d itio n to char ac ter izing t he mea sur e men t s yste m’
s p er fo r ma nce , si mul taneo us d etec tio n
at multiple locations enables velocimetry by analyzing the transit time between detection regions.
Particle image velocimetry [19 ] and fluorescence correlation spectroscopy [20,21] both use
correlation techniques to obtain velocity information, and have frequently been used in microfluidic
devices. A major advantage of using our system over previous systems is that we are not limited to
the single field of view of a microscope objective. By using arrays of high NA astigmatic lenses, we
can obtain high time resolution, because the detection regions are small, and a large effective field
of view, because the detection regions are spaced far apart from each other. This large effective
field of view enables a much larger dynamic range in transit time measurements.
Fig. 7(a) shows the cross correlation of two consecutive detection regions found from the data set
that is also plotted as Fig. 6. The curve shows a sharp spike occurring at a time delay o f 3 6 0 µ
s. Co nsec uti ve d etec tio n r egio n s ar e sp ac ed b y 2 0 0 µ m, so thi s r isi ng ed ge o f
the correlation curve corresponds to a maximum velocity of 560 mm/s. The correlation peak has
a finite width, which implies that beads in the channel have a distribution of velocities. Beads
traveling down the center of a microfluidic channel have the fastest velocity, while beads closer
to the channel walls travel slower. From Fig. 7(a), we also observe that the correlation cur ve has
a sec o nd ar y p ea k at a ti me d ela y o f 6 8 0 µ s, almo st t wice a s lo n g as the r i sin g
ed ge.
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
17 January 2011 / Vol. 19, No. 2 / OPTICS EXPRESS 1393
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(C) 2011 OSA
Fig. 7.
Size resolved velocity dispersion. (a) Cross correlation of the total signal of a
h et erog en eou s samp le of 0 . 5 an d 1. 1 µm b ea d s from t wo c on se cutive detection regions. (b)
Cross correlation of the intensity gated signals from the same two detection regions, showing t ha t t
h e 1 .1 µm b ea d s ha ve grea t er v el oc i t y d i sp ersi on .
As is common in flow cytometry, we are able to gate the time signals by intensity [1], and co
nseq ue ntl y a nal yze sep ar at el y t he si gnal s fr o m the 5 0 0 n m and the 1 . 1 µ m b ea
d s. U nli ke flow cytometry, however, we have access to data at multiple detection regions which
enables velocity measurements. Fig. 7(b) sho ws t he cr o ss co r r elation o f the 5 0 0 n m and 1 .
1 µ m b ea d s separately. We observe that the maximum velocity of each bead size occurs at the
same time d elay, b ut t hat a lar ger fr ac tio n o f t he 1 . 1 µ m b ea d s ar r ive at lar ger ti me
d ela ys. W e b elieve that this i s ca used b y t he fa ct that mo r e 1 . 1 µ m b ea d s a r e far ther
fr o m t he ce nter o f the channel. Although a complete investigation is outside the scope of this
paper, this is consistent with recent studies on inertial focusing of particles in microfluidic channels
[22 ]. Inertial forces push particles away from the channel center and have been found to act
stronger on particles that fill a greater fraction of the channel width.
6. Conclusions We have demonstrated a fluorescence detection system based on the integration of
an
astigmatic diffractive microlens array with microfluidics. By shaping the focal spot distribution of
the diffractive microlenses into a line, we obtain high time resolution and reduce the coefficient of
variation of the measured signal for homogenous samples. Arrays of high numerical aperture
microlenses enable efficient light collection over much larger regions than would be possible using
a single microscope objective. Due to the efficient fluorescence collection, we have demonstrated
high throughput counting and sizing of beads at 8,300 per second at multiple regions along the
channel. We have also exploited the multiple detection regions to demonstrate a new velocimetry
technique based on a cross correlation analysis of fluorescence signals occurring at two different
detection regions along the channel. The combination of velocimetry and flow cytometry enables
characterization of the interaction between complex flow distributions and heterogeneous
distributions of particles.
Acknowledgments This work was supported by the Advanced Energy Consortium via the Bureau
of Economic
Geology at the University of Texas at Austin. P.E.S. acknowledges support from the Center for
Excitonics, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office
of Science, Office of Basic Energy Sciences under Award number DESC0001088. Fabrication
work was carried out in the Center for Nanoscale Systems (CNS) at Harvard, which is also
funded by the National Science Foundation.
Received 3 Nov 2010; revised 15 Dec 2010; accepted 22 Dec 2010; published 12 Jan 2011
17 January 2011 / Vol. 19, No. 2 / OPTICS EXPRESS 1394
#137673 - $15.00 USD
(C) 2011 OSA