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Supplementary material: Selective plane illumination microscopy on a
chip
Petra Paiè,a,†
Francesca Bragheri,b,†
Andrea Bassi, a,*
Roberto Osellameb
aDipartimento di Fisica, Politecnico di Milano, Piazza Leonardo da Vinci 32, 20133 Milano, Italy
bIstituto di Fotonica e Nanotecnologie, CNR, Piazza Leonardo da Vinci 32, 20133 Milano, Italy
*Corresponding author: [email protected]
TEXT
Chip fabrication by femtosecond laser micromachining
The chip was realized by using femtosecond laser micromachining 1-3. The
irradiation was performed by focusing through a 50x, 0.6 NA microscope
objective the second harmonic of a commercial femtosecond laser
(femtoREGEN, HIGH-Q Laser) emitting pulses of 400 fs, 1040 nm
wavelength and energy up to 23 μJ at 960 kHz repetition rate. Scan
velocities and pulse energies were varied in relation to the depth of the
irradiated structure with respect to the glass surface to compensate the
spherical aberrations. The geometry of the device was obtained by properly
translating the sample relative to the laser beam with a system of high
precision air bearing translation stage (Fiberglide 3D, Aereotech).
The different components, i.e. the lens and the microchannel network along
with their respective access holes for external tubes connection, were
irradiated in the same fabrication step with pulse energies of 270 nJ for the
lens, 350 nJ for the access holes and 500 nJ for the microchannel. The
translation speed was 1 mm/s for the lens and 2 mm/s for all the other
structures.
Two different irradiation geometries are possible, longitudinal and
transverse, in which the sample is translated, respectively, along and
perpendicularly to the beam propagation direction. In the first case,
exploited for the lens irradiation, the channel develops parallel to the writing
beam allowing a reduction of the wall roughness to few nanometers 4,5. The
desired acylindrical shape was obtained by irradiating from the bottom to
the top of the substrate 50 section of the lens (with 6 μm separation in
depth), each having the optimized profile designed for spherical aberrations’
reduction.
The transverse irradiation geometry was used to fabricate the H-
microchannel with square cross-section. In particular a multi-scan
irradiation approach was exploited: contiguous straight lines (with a
separation of 2 m) were scanned, forming the lateral surface of a
rectangular cross-section cylinder. For each channel branch six coaxial
cylinders were irradiated (with dimensions of 40x30, 90x70,180x150,
280x240, 380x330, 480x400 m2), so as to obtain the desired channel with
a cross-section of 500 m after the etching step. Similarly, access holes
were obtained by irradiating seven coaxial circular helices with diameters
equally spaced from 80 to 560 m. The total irradiation time was
approximately 2 hours.
Light sheet profile characterization
The lens was designed to have a long focal length (0.7 mm) as well as a
long depth of focus (0.4 mm) so as to illuminate the whole microchannel
section in a uniform manner. Being the minimum achievable waist limited
by the long depth of focus, we designed a waist of 5.5 μm, by considering
the lens filled with a high refractive index oil, n = 1.56 (Fig. S2).
In order to evaluate the light sheet profile we fabricated a separate device
with the lens facing a 800 μm wide and 35 μm height reservoir and we
filled it with a Rhodamine solution (Fig. S3a). The reservoir was centered at
the expected focusing position.
The light sheet profile was observed by capturing the Rhodamine
fluorescence signal with a CCD mounted on an standard inverted
microscope (DMI 3000M, Leica), using a 10x, 0.25 NA microscope
objective. The acquired fluorescence images were subsequently analyzed
(Matlab) to retrieve the beam radius along the entire channel and to
determine the waist of the beam and its position.
The routine automatically analyze the acquired fluorescence images first by
sectioning them along the beam propagation direction and then by fitting
the fluorescence signal of each section with a gaussian intensity profile to
find out the correspondent waist (Fig. S3b). A correction in the Gaussian
model has been inserted so as to take into account the effect of the depth of
field of the microscope objective used to acquire the fluorescence image.
The optimized lens shows a minimum waist close to the theoretical one,
equal to 5.8 μm. This result confirms that designed acylindrical profile
allows reducing the impact of spherical aberration so as to obtain a beam
waist size close to the theoretical one.
Pumping scheme
Pressure driven pumps (Fluigent, MCFS Flex) were used to inject and
control the sample and the buffer flow in the device. The H-shaped (Fig.
S1) geometry of the channel was chosen so as to exploit the laminarity of
fluids in microfluidic channels. Indeed thanks to this property the streams of
the two channels, sample and buffer, won’t mix when merging into the
central common branch and the fluids entering at sample or buffer input will
exit at sample and buffer output respectively. The velocity of the flow is
controlled by unbalancing the pressure at the input and output inlets
separately for each stream. The buffer stream was exploited to prevent
spheroids touching the channel wall. Indeed the interface between the two
streams can be moved by accordingly balancing the sample and the channel
driving pressures. This allows one to make the spheroids flowing
orthogonally with respect to the light sheet without touching the channel
wall that might induce a rotation of the sample
Spheroids
H2B-mCherry expressing tumor spheroids were prepared as described in
the references 6,7. Fixed samples were passed in dilutions of 2,2’-
thiodiethanol (TDE, Sigma) and PBS in a stepwise manner (25%, 50%,
68%, for 10 min), to finally match the index of refraction of fused silica
(n=1.46).The liquid used in the fluidic channels was a solution of water and
TDE, to which we added 0.1% (final concentration) low melting point
agarose (LMA Sigma) in order to slightly increase the liquid viscosity and
Electronic Supplementary Material (ESI) for Lab on a Chip.This journal is © The Royal Society of Chemistry 2016
consequently to better control the sample movement: this being particularly
important when the sample was slowly scanned through the light sheet.
Correction for light sheet in-homogeneities
During imaging we observed that the sample was illuminated by an in-
homogeneous excitation intensity. This was primarily due to the non-perfect
alignment of the optical fiber which was manually glued to the chip. In
order to correct non-homogeneous illumination, before or after the
measurement session, the channel was filled with a nanobeads solution,
which was circulated at high speed through the light sheet. This creates an
image on the sensor that mapped the intensity distribution of the light sheet
on the entire field of view (reference image).
All the acquired images were then divided by the reference image in order
to correct in-homogeneous illumination (Fig. S4). The correction was less
effective in high-throughput measurements, where the lower signal to noise
ratio of the data resulted in an amplification of the noise in the upper part of
the image (visible in Fig. 3).
Nuclei Segmentation and visualization
Nuclei segmentation was obtained using grayscale morphological image
processing operations 8, implemented in Matlab.
Sample sections, Maximum Intensity Projections and 3D volumes were
visualized using Fiji 9.
Nuclei segmentation
Nuclei segmentation was obtained using well known grayscale
morphological image processing operations8, implemented in Matlab. In
details we performed:
- volume erosion by a sphere with diameter of approximately half the size
of the nuclei (5 µm), followed by filtering by reconstruction (opening by
reconstruction);
- volume dilation by a sphere with 5 µm diameter, followed by filtering by
reconstruction (closing by reconstruction);
- volume opening by a 2 µm diameter sphere (consisting in image erosion
followed by dilation);
- calculation of the regional maxima of the volume;
- detection of the connected components of the final binary volume, which
correspond to the segmented nuclei.
The procedure was applied to the entire volume (c.a. 1000x1000x200
voxels), requiring c.a. 5 min/volume on an Intel I7-4770K with 32GB of
RAM memory. The procedure was also performed plane by plane,
requiring c.a. 300 ms/plane. For high-throughput acquisitions the
segmentation was applied only to a single plane of the acquired stack
corresponding to a depth of c.a. 70 µm within the sample (Fig. 3 and Fig.
S8). The Matlab code for the segmentation of the nuclei is available on
request.
Spheroid volume quantification
Segmentation of the spheroids and measurement of their volume was
performed by applying a series of binary morphological operations to the
acquired SPIM stacks, using Matlab. The acquired grayscale images were
first converted to a binary image by setting a proper threshold level. Then
morphological opening (image dilation followed by erosion) and filling
(consisting in the removal of regional minima that are not connected to the
image border) were performed, in order to segment the area of the spheroid
(green in Fig. S9) within the image. The batch of samples acquired in high-
throughput mode presented a large central empty region. This region was
obtained as the difference between the image processed by opening and
filling, and the same image processed only with the opening operation (red
in Fig. S9).
The time required for segmentation of a single plane was c.a. 30ms,
compatible with real time processing during high-throughput acquisition.
The procedure was repeated in all planes to measure the entire volume of
the spheroid and the volume of the empty region. The Matlab code for
segmentation and quantification is available on request.
FIGURES
Fig. S1 Schematic of the fluidic channel. (a) Schematic of the H-shaped microfluidic channel, with the microscope objective above the chip. The sample is moved by the flux through the fluidic network. (b) Two-dimensional scheme of the H- shaped fluidic network. The movement of the sample is carefully controlled by four tubes each of which provides a selected pressure to the liquid. The geometry has been chosen to avoid the spheroid hitting a wall due to its inertia after the 90° turn. The sample is observed from above with a microscope objective; due to the symmetry of the chip, observation from below (not shown) or double sided detection are also possible. Figures are not in scale.
Fig. S2 Schematic diagram of the procedure used to optimize the lens shape. The first interface of the lens is designed to be perpendicular to each ray composing the beam so that the beam propagation is not modified. The second interface is designed to focus all the beam radii in the same position. Each coordinate of the aspherical profile is retrieved by geometrical considerations on the known angles. The design parameters are summarized in the table on the right hand side. The desired focal length of the lens is indicated with f, the waist is w0 the and the confocal parameter is 2z0
a b
Fig. S3 Lens characterization. (a) Schematic of the chip used to characterize the focusing properties of the lens. The light emitted by the fiber creates a light sheet in a reservoir filled with Rhodamine. The chip is imaged under an inverted microscope to capture the light sheet
profile, in direction perpendicular to it. (b) Image of the light sheet profile obtained with the acylindrical lens in the Rhodamine-filled reservoir. (c) Results of the waist retrieval. The fluorescence image is sectioned along the beam propagation direction (x axis) and the intensity of each section is fitted with a gaussian profile (red) that allows determining the correspondent waist w0. A minimum value of 5.8 µm is obtained.
a
b
c
Fig. S4. Correction for light sheet in-homogeneities. Results of the correction for non-uniform illumination (see also Online Methods for details). Section of a H2B-mCherry expressing spheroid, acquired at z=100 µm in depth, before (a) and after correction (b). Maximum Intensity Projection (MIP) of the spheroid, before (c) and after correction (d). Scale bar, 100µm.
a b
c d
Original Corrected
xy xy
MIP MIP
Original Corrected
Fig. S5. H2B-mCherry expressing spheroid, acquired at various depths, shown with 14 µm step. Arrows indicate three nuclei undergoing mitosis. Scale bar, 100µm.
z=30 µm z=44 µm z=58 µm z=72 µm
z=86 µm z=100µm z=114µm z=128 µm
z=142 µm z=156µm z=170µm z=184 µm
z=198 µm z=212µm z=226 µm z=240µm
xy
Fig. S6. Segmented nuclei of a 250µm diameter H2B-mCherry expressing spheroid; (a) xy sections of the spheroid, acquired at various z-depths (left to right), shown with 45 µm steps; (b) xy sections with superimposed colored segmented nuclei; (c) xz section of the spheroids at various height (left to right), shown with 45 µm step; (d) xz sections with superimposed colored segmented nuclei. Each color corresponds to a different structure. Scale bar, 100µm.
a
b
c
d
xy
xz
xy
xz
Fig. S7. Details of nuclei undergoing mitosis. (a) Nuclei at the depths z=30um (b) z=128 µm. (c) z=212 µm. Nuclei can be observed in detail along different orientations. Yellow lines indicate the position of the transverse (xy) sagittal (zy) and coronal (xz) sections. Scale bar, 10µm.
xy xy xy zy zy zy
xz xz xz
a b c
Fig. S8. Nuclei count in high-throughput acquisitions. (a) Single section of a H2B-mCherry expressing spheroid (sample #1 in Fig.3) at z=70 µm in depth, acquired in high-throughput mode. Scale bar, 100µm. (b) Image in a with overlapped segmented nuclei (c) Statistical analysis of the fluorescence intensity in n=25 spheroids: histogram of the fluorescence intensity showing different intensity ranges (intensity 1 corresponds to the maximum signal detected in all samples) and the number of nuclei whose fluorescence intensity fits in each range. The nuclei count in each range is the sum over the n=25 samples. Standard deviation, calculated as the deviation in nuclei count within the n=25 samples, is shown for each range of the histogram.
Intensity (a.u.)
Nu
clei
#
a b
c
xy xy
High-throughput High-throughput
Fig. S9. Quantification of the volume of spheroids in high-throughput measurements. The acquired samples presented a central empty region whose volume was also quantified. (a) Single section of a 300µm diameter H2B-mCherry expressing spheroid acquired, at z=150 µm in depth, in high throughput mode. The green and red curves indicate the borders of the spheroid and its central empty region respectively. (b) Maximum Intensity Projection (MIP) of a stack of 200 planes, highlighting the entire volume of the spheroid (green) and empty region (red). Scale bar, 100µm. (c) Statistical analysis of n=25 spheroids: histogram of the spheroids’ volume (green) and histogram of the volume of the internal empty regions (red) showing different volume ranges and the percentage of samples whose volume fit in each range.
xy MIP
a b
Volume (x10-3
mm3)
Per
cen
tage
of
sam
ple
s (%
)
c
Spheroid volume Empty volume
High-throughput High-throughput
MEDIA CAPTION
Movie 1 legend
Side view of sample flow in microchannel, to observe the effect of the 90°
curvature in two microchannel with different shape. Sample rotation
induced in the ‘C’ shaped channel is avoided with the introduction of
auxiliary channels in an ‘H’ shaped configuration.
Movie 2 legend.
Optical sectioning of a cellular spheroid.
Real time acquisition of a H2B-mCherry expressing spheroid, slowly
moving through the light sheet (20µm/s). The images are captured at 12 Hz
and corrected for illumination in-homogeneities as described in the
Methods. Scale bar, 100µm
Movie 3 legend.
3D reconstruction of segmented nuclei.
Three dimensional reconstruction of a H2B-mCherry expressing spheroid.
The sample is virtually rotating around itself and after one rotation the
image of the 3D reconstruction of the segmented nuclei (red) is overlapped
to the original sample.
Movie 4 legend
High-throughput acquisition.
Real time acquisition of a H2B-mCherry expressing spheroid, quickly
moving through the light sheet (c.a. 150µm/s). Acquisition starts when the
mean fluorescence signal calculated over the entire image overcome a
certain threshold and continues until the signal decreases below the same
threshold. The acquisition continues for c.a. 2s depending on the sample
size. For visualization, a red circle appears at the frames corresponding to
the acquisition start. Black frames (90 frames) are concatenated to the
acquired stack after each acquisition to visualize the samples separately. No
image correction is applied to the data for illumination in-homogeneities.
Scale bar, 100µm.
† These Authors had equal contribution to this paper.
REFERENCES
1. R. Osellame, H. J. Hoekstra, G. Cerullo, & M. Pollnau,
Femtosecond laser microstructuring: an enabling tool for optofluidic
lab‐on‐chips. Laser & Photonics Reviews 5, 442-463 (2011).
2. R. Gattass, & E. Mazur, Femtosecond laser micromachining in
transparent materials Nature Photon. 2, 219 (2008).
3. R. Osellame, V. Maselli, R. Martinez Vazquez, R. Ramponi, & G.
Cerullo, Integration of optical waveguides and microfluidic
channels both fabricated by femtosecond laser irradiation Appl.
Phys. Lett. 90, 231118 (2007).
4. S. Ho, P. R. Herman, & J. S. Aitchison, Single-and multi-scan
femtosecond laser writing for selective chemical etching of cross
section patternable glass micro-channels. Applied Physics A 106, 5-
13 (2012).
5. T. Yang, P. Paiè, G. Nava, F. Bragheri, R. Martinez Vazquez, P.
Minzioni, M. Veglione, M. Di Tano, C. Mondello, R. Osellame, &
I. Cristiani, An integrated optofluidic device for single-cell sorting
driven by mechanical properties. Lab on a Chip, 15, 1262-1266
(2015).
6. C. Lorenzo, C. Frongia, R. Jorand, J. Fehrenbach, P. Weiss, A.
Maandhui, & V. Lobjois, Live cell division dynamics monitoring in
3D large spheroid tumor models using light sheet microscopy. Cell
Div 6, 22 (2011).
7. A. Ivascu, & M. Kubbies, Rapid generation of single-tumor
spheroids for high-throughput cell function and toxicity analysis.
Journal of biomolecular screening 11, 922-932 (2006).
8. L. Najman, & H. Talbot, Mathematical morphology: from theory to
applications, ISTE-Wiley. (2010).
9. J. Schindelin, et al., Fiji: an open-source platform for biological-
image analysis, Nature Methods 9(7), 676-682 (2012